PRC1 Cooperates with CLASP1 to Organize Central Spindle Plasticity in Mitosis

Abstract

During cell division, chromosome segregation is governed by the interaction of spindle microtubules with the kinetochore. A dramatic remodeling of interpolar microtubules into an organized central spindle between the separating chromatids is required for the initiation and execution of cytokinesis. Central spindle organization requires mitotic kinesins, microtubule-bundling protein PRC1, and Aurora B kinase complex. However, the precise role of PRC1 in central spindle organization has remained elusive. Here we show that PRC1 recruits CLASP1 to the central spindle at early anaphase onset. CLASP1 belongs to a conserved microtubule-binding protein family that mediates the stabilization of overlapping microtubules of the central spindle. PRC1 physically interacts with CLASP1 and specifies its localization to the central spindle. Repression of CLASP1 leads to sister-chromatid bridges and depolymerization of spindle midzone microtubules. Disruption of PRC1-CLASP1 interaction by a membrane-permeable peptide abrogates accurate chromosome segregation, resulting in sister chromatid bridges. These findings reveal a key role for the PRC1-CLASP1 interaction in achieving a stable anti-parallel microtubule organization essential for faithful chromosome segregation. We propose that PRC1 forms a link between stabilization of CLASP1 association with central spindle microtubules and anti-parallel microtubule elongation.

To ensure that each daughter cell receives the full complement of the genome in each cell division, chromosomes move poleward, and non-kinetochore fibers become bundled at the onset of anaphase, initiating assembly of the central spindle, a set of anti-parallel microtubules that serves to concentrate key regulators of cytokinesis (1–3). Chromosomal passengers are a group of evolutionarily conserved proteins that orchestrates chromosome segregation and central spindle plasticity (4, 5). This protein complex containing Aurora B, Survivin, INCENP, and Borealin is relocated from the kinetochore to the central spindle upon anaphase onset (5–9). Perturbation of their function results in defects in metaphase chromosome alignment, chromosome segregation, and cytokinesis (10).

Among the central spindle maintenance components, only two have been reported to mediate the microtubule bundling in the central spindle. One is centralspindlin, a heterotetramer containing CeMKLP1/ZEN-4 and RhoGAP/CYK-4 (11), and the other one is an evolutionarily conserved protein, PRC1 (also named Feo in fruit fry, Ase1 in yeast, and MAP65 in plant cells). PRC1 is a non-motor microtubule-binding and -bundling protein in human cells originally identified as a Cdc2 substrate essential for cytokinesis (12, 13). Similar microtubule regulatory activities have been reported in yeast, fruit fly, and plant cells. It is well known that overexpression of wild type PRC1 in HeLa cells can result in thick microtubule bundles in cells at interphase (13). Bundling activity of PRC1, as well as centralspindlin, is required for the organization of the central spindle as well as for the successful progression of cytokinesis. PRC1 molecules accumulate on the midline of a central spindle with the cell cycle progression to anaphase. As a non-motor microtubule-binding protein, transportation of PRC1 to the midline is promoted by its association to kinesin, KIF4A, and timing of this progression is controlled by the dephosphorylation of Thr-481 on PRC1 when the cell exits metaphase by phosphatase Cdc14 (14). Our recent study shows that prevention of the phosphorylation of PRC1 at Thr-470 causes an inhibition in PRC1 oligomerization in vitro and an aberrant organization of central spindle in vivo, suggesting that this phosphorylation-dependent PRC1 oligomerization ensures that central spindle assembly occurs at the appropriate time in the cell cycle (15).

Spatiotemporal regulation of microtubule organization and dynamics is responsible for the mitotic apparatus such as the central spindle. However, it has remained elusive as to how the central spindle microtubule organization and dynamics are regulated. There are large varieties of microtubule-associated proteins responsible for regulation of the dynamic behavior of microtubules and microtubule-mediated transport. Among these, proteins that associate with the tips of microtubules are called +TIPs, for “plus-end tracking proteins.” These proteins have been shown to be important in different organisms and cellular systems (16). Using yeast two-hybrid assay, CLASPs were identified as interacting partners of the CLIPs and characterized as new +TIP proteins (17).

The microtubule-binding protein CLASP is emerging as an important microtubule regulator in the formation of the mitotic apparatus (18–22). CLASP is required for promoting plus-end growth of spindle microtubules in prometaphase (23). Although the molecular mechanisms underlying its regulation of microtubule dynamics remain elusive, it is generally believed that CLASP orchestrates microtubule dynamics via its physical interacting with EB1, CLIP170, and microtubules (17, 24).

To delineate the molecular function of PRC1 in central spindle organization and spatiotemporal regulation, we carried out a new search for PRC1-interacting proteins. Our studies show that PRC1 physically interacts with CLASP1, and the two proteins cooperate in the organization of the central spindle. Our studies provide a novel regulatory mechanism in which the PRC1 complex operates central spindle organization in mitosis.

MATERIALS AND METHODS

Expression Constructs

Human CLASP1 cDNA was obtained from the Kazuza DNA Research Institute. GFP-CLASP1 contained the whole open reading frame of KIAA0622, subcloned into XhoI-XhoI sites of the mammalian expression vector pEGFP-C1 (Clontech). All the deletions of CLASP1 were generated by PCR. GST⁴ and 3×FLAG plasmids of full-length PRC1 and its various mutants were prepared as described previously (15).

Recombinant Protein Production

Purification of recombinant proteins was carried out as described previously (15). Briefly, 1 liter of LB media (10 g of NaCl, 10 g of tryptone, and 5 g of yeast extract, pH 7.4) was inoculated with bacteria transformed with GST-PRC1 and its mutants, respectively. When the optical density (A₆₀₀) of the bacteria culture reached between 0.6 and 0.7, the protein expression was initiated by addition of 0.5 mm isopropyl d-thiogalactopyranoside for 3 h. Bacteria were then collected by centrifugation and resuspended in PBS containing proteinase inhibitors (leupeptin, pepstatin, and chymostatin; 5 μg/ml) followed by sonication and clarification as described previously (15). The GST fusion protein in bacteria in the soluble fraction was purified using glutathione-agarose chromatography.

To generate TAT-GFP proteins, we recombined the pET-22b vector with an 11-amino acid TAT sequence followed by a GFP gene (25). TAT-GFP-His fusion proteins were expressed and purified as described previously (25).

Generation of TAT Fusion Protein for Functional Analysis

For introducing TAT-GFP fusion proteins to probe for the functional relevance of PRC1-CLASP1 interaction, HeLa cells were cultured to 50–60% confluency before experimentation. Just before introduction, the cells were washed with serum-free media and incubated with TAT-GFP fusion peptides at various concentrations. After incubation, the cells were washed with PBS and then examined directly under fluorescence microscopy.

To analyze the relative level of PRC1 and CLASP1 associated with the cytoskeleton in response to TAT-PRC1 peptide treatment, transfected cells were rinsed twice with PHEM buffer (100 mm PIPES, 20 mm HEPES, pH 6.9, 5 mm EGTA, 2 mm MgCl₂, and 4 m glycerol) followed by incubation in PHEM buffer containing a proteinase inhibitor mixture (pepstatin-A, leupeptin, aprotinin, and chymostatin; final concentration 5 μg/ml for each inhibitor) plus 0.1% Triton X-100 for 2 min at room temperature to allow cytosolic proteins to be released into the extracellular medium. The extracted cells were then harvested from the Petri dish and centrifuged at 1,500 × g for 5 min. The resulting pellets, designated as insoluble materials, were solubilized in 1× SDS-PAGE sample buffer, and the supernatants, called the soluble fraction, were concentrated with 5% trichloroacetic acid followed by reconstitution in 1× SDS-PAGE sample buffer as described (26).

Cell Cultures and Transfection of Plasmids and siRNAs

HeLa and 293T cells (American Type Culture Collection, Manassas, VA) were cultivated as subconfluent monolayers in Dulbecco's modified Eagle's medium (Invitrogen) with 10% fetal bovine serum (HyClone, UT) and 100 units/ml penicillin plus 100 μg/ml streptomycin at 37 °C with 10% CO₂.

Lipofectamine transfection reagents (Invitrogen) were used for plasmid transfection. Cells were transfected with Lipofectamine 2000 pre-mixed with various plasmids (2 μg/ml) as described above. Stable clones were selected in the presence of 0.5–1 mg/ml G418 (Calbiochem). Synthetic siRNAs were transfected, using Oligofectamine (Invitrogen). siRNA oligonucleotides were directed against the following target sites: CLASP1B, 5′-GGATGATTTACAAGACTGG-3′ (24); PRC1, 5′-ATATGGGAGCTAATTGGGA-3′. In the trial experiments, different concentrations of siRNA oligonucleotides were used for different time intervals of treatment, and transfection efficiency was judged based on the uptake of fluorescein isothiocyanate-conjugated oligonucleotides. In pilot experiments, HeLa cells were transfected with different concentrations of siRNA oligonucleotides or control scramble oligonucleotides for different time intervals. Transfected cells were then collected and solubilized in SDS-PAGE sample buffer. The efficiency of this siRNA-mediated protein suppression was judged by Western blotting analyses of target proteins.

Antibodies

His-tagged full-length PRC1 protein, purified by nickel-agarose beads, was used for immunization of BALB/c mice. Three mice were boosted every 3 weeks until the antibody titer reached 1:1,500 as judged by Western blotting. The splenocytes from immunized mice were fused with FO myeloma cells (American Type Culture Collection; Manassas, VA) according to standard protocol. Mono-specific antibodies against PRC1 were selected based on Western blotting and immunocytochemistry. The polyclonal antibodies against PRC1 were generated as described (27). A rabbit anti-CLASP1 antibody was the kind gift from Dr. Fedor Severin (Moscow State University, Russia; see Ref. 22).

Pulldown Assay

The GST-PRC1 (full-length and its mutants) fusion protein-bound Sepharose beads were incubated with 293T cell lysate containing GFP-tagged CLASP1 or deletion mutants for 4 h at 4 °C. After the incubation, the beads were washed twice with PBS containing 0.5% Triton X-100 and twice with PBS and boiled in SDS-PAGE sample buffer. The bound proteins were then separated on 10% SDS-polyacrylamide gel. Separated proteins were then transferred onto nitrocellulose membrane for Western blotting with a monoclonal GFP antibody.

To test whether the recombinant TAT-GFP-PRC1-(486–620) peptide disrupts PRC1*CLASP1 interactions, GFP-CLASP1-bound GST-PRC1 affinity beads were incubated with 5 μm TAT-GFP-PRC1-(486–620) peptide for 30 min on a rotator at room temperature. The affinity beads were then collected and boiled in SDS-PAGE sample buffer. The proteins bound were subsequently subjected to SDS-PAGE analysis as described previously.

Synchronization, Immunoprecipitation, and Immunoblot Analysis

293T or HeLa cells were cultured with Dulbecco's modified Eagle's medium supplemented with 10% fetal calf serum. To obtain the mitotic cells, cells were synchronized by treatment with thymidine for 16 h, washed with PBS three times, and incubated with fresh medium for 10–11 h. The cells or cell extracts at mitosis were subjected to immunofluorescence or immunoprecipitation analysis as described previously (15).

Briefly, for the immunoprecipitation assay, GFP-CLASP1 or its deletion mutants and 3×FLAG-PRC1 co-expressing 293T cells were solubilized in lysis buffer (50 mm Tris-HCl, pH 7.4, 150 mm NaCl, 1 mm EDTA, 0.1% Triton X-100) on ice followed by centrifugation (14,000 rpm for 20 min at 4 °C). 3×FLAG-PRC1 proteins were harvested with Ezview^TM red anti-FLAG M2 affinity beads (Sigma). These beads were washed and boiled in SDS-PAGE sample buffer and then subjected to immunoblot analysis with a monoclonal GFP antibody and FLAG antibody, respectively.

For immunoblotting, proteins were probed by appropriate primary antibodies. Immunoreactive signals were then detected with horseradish peroxidase-conjugated secondary antibodies using ECL kit (Pierce) and visualized by autoradiography on Kodak BioMAX MS film.

Immunofluorescence Microscopy

After transfection or drug treatment, cells on acid-treated glass coverslips were washed with PHEM (60 mm PIPES, 25 mm HEPES, 10 mm EGTA, 2 mm MgCl₂, pH 6.9). Permeabilization was carried out for 1 min in PHEM buffer containing 0.1% Triton X-100 at 37 °C. Pre-extracted cells were fixed in 2% paraformaldehyde in PHEM buffer for 10 min. After washing three times with PBS, cells were blocked with 1% bovine serum albumin (Sigma) in PBS containing 0.05% Triton X-100 for 30 min, then incubated with primary antibodies for 1 h at room temperature, then 1 h at 4 °C, followed by secondary antibodies for 45 min. DNA was stained with DAPI (Sigma).

Deconvolution images were collected using a Deltavision wide field deconvolution microscope system built on an Olympus IX-70 inverted microscope base. For imaging, a ×100 1.35 NA lens was used, and optical sections were taken at intervals of 0.2 μm. Images were processed using DeltaVision Softworx software as described previously (27). Images for display were generated by projecting the sum of the optical sections using the maximum intensity method. For presentation of details of the internal central spindle in Fig. 6, projection images were constructed from a 1-μm section within the cell.

FIGURE 6.

PRC1 collaborates with CLASP1 for faithful mitosis. A, schematic illustration of experimental strategy for assessing the role of PRC1-CLASP1 interaction in central spindle assembly and regulation. Monastrol-arrested prometaphase cells were used to induce mitotic progression by washout of monastrol and release into metaphase by inclusion of 20 μm MG132. To test for the functional relevance of PRC1-CLASP1 interaction, 500 nm TAT-GFP-PRC1-(438–620) peptide and its control peptide TAT-GFP were included for real time imaging of mitotic progression. B, live cell imaging of mitotic progression of a HeLa cell stably expressing mCherry-tubulin. The monastrol-arrested cell was subjected to monastrol wash-out and incubated with 20 μm MG132 and 500 nm TAT-GFP for 60 min before anaphase onset by release into MG132-free but TAT-GFP-containing media. Imaging time begins after the addition of TAT peptides. A representative cell was shown from prometaphase to anaphase onset until telophase. Scale bars, 10 μm. C, live cell imaging of mitotic progression of a HeLa cell-treated TAT-GFP-PRC1-(438–620) (TAT-GFP-PRC1^438∼620) peptide (500 nm). In contrast, in TAT-GFP-treated cells, the duration of anaphase was prolonged for 2 min. Scale bars, 10 μm. D, live cell imaging of mitotic progression of a HeLa cell-treated CLASP1 siRNA for 24 h before monastrol synchronization and TAT-GFP-PRC1-(438–620) peptide was added (500 nm). Scale bars, 10 μm. E and F, quantitative analyses of mitotic progression in the PRC1-CLASP1 perturbed cells. Pre-anaphase progression (from prometaphase to anaphase onset) was analyzed and presented in E, which exhibits no statistically significant difference between TAT-GFP-treated and TAT-GFP-PRC peptide-treated sample (p > 0.05). Post-anaphase progression (from anaphase onset to telophase) was analyzed and presented in E and F. In general, it takes 621.4 ± 16.5 s (n = 8) for TAT-GFP-treated cells to complete the aforementioned process, and it takes 700.6 ± 27.0 s (n = 8) for TAT-GFP-PRC1-(438–620)-treated cells to complete the same process. This difference bears statistical significance (p < 0.01). G, quantification of the duration of mitosis from anaphase onset to midbody formation in PRC1-suppressed cells and CLASP1-repressed cells. On average, control cells transfected with scramble oligonucleotides take 603.5 ± 22.0 s (n = 4) to complete the aforementioned process whereas PRC1-depleted cells take 641.0 ± 27.8 s and CLASP1-depleted cells require 1079.0 ± 92.5 s to complete the same process (n = 4). H, perturbation of PRC1-CLASP1 interaction abrogates chromosome stability in mitosis. Asynchronized HeLa cells were treated with 500 nm recombinant TAT-GFP (panels a″–d″) or TAT-GFP-PRC1-(438–620)⁰ (panels a–d) for 30 min. After treatment, cells were extensively washed, fixed, and stained with an anti-tubulin antibody (red) and DAPI (blue). Recombinant GFP-his protein was added as a control (panels a′–d′). Note that perturbation of PRC1-CLASP1 interaction by treatment of TAT-GFP- PRC1-(438–620) resulted in chromosome lagging (panel b) and formation of a sister chromatid bridge (panel c), a characteristic phenotype of chromosome instability. This phenotype is absent in TAT-GFP-treated and GFP-treated cells (panels b″–d″ and b′–d′). Scale bars, 10 μm. I, quantification of the overlapped microtubule bundles at the central spindle as function of PRC1-CLASP1 interaction. The monastrol-arrested cells stably expressing mCherry-tubulin were subjected to monastrol wash-out and incubated with 20 μm MG132 and 500 nm TAT-GFP for 60 min before anaphase onset by release into MG132-free but TAT-GFP or TAT-GFP-PRC1-(438–620) peptide containing media as illustrated in B–D. Optical sections taken from the middle of the z-stack were scored for the number of overlapped microtubule bundles in the TAT-PRC1-(438–620)-treated cells and expressed as mean ± S.E., n = 12 from three different preparations (p < 0.01 compared with TAT-GFP treatment). Representative images from anaphase A and anaphase B are shown on the left, and statistical analyses are listed on the right. J, quantification of the stable microtubule plus-ends at the central spindle as function of PRC1-CLASP1 interaction. The monastrol-arrested HeLa cells stably expressing mCherry-EB3 were subjected to monastrol wash-out and incubated with 20 μm MG132 and 500 nm TAT-GFP for 60 min before anaphase onset by release into MG132-free but TAT-GFP or TAT-GFP-PRC1-(438–620) peptide-containing media. Optical sections taken from the middle of the z-stack were scored for the number of mCherry-EB3-marked microtubule plus-ends in the TAT- PRC1-(438–620)-treated cells. The number of EB3-marked microtubule plus-ends within the middle one-third of the spindle axis (boxed area) were scored and expressed as mean ± S.E., n = 9 from three different preparations (p < 0.01 compared with TAT-GFP treatment). K, quantification of anaphase defects in PRC1-CLASP1 interaction-disrupted cells. The monastrol-arrested cells stably expressing mCherry-tubulin were subjected to monastrol wash-out and incubated with 20 μm MG132 and 500 nm TAT-GFP for 60 min before anaphase onset by release into MG132-free but TAT-GFP or TAT-GFP-PRC1-(438–620) peptide-containing media as illustrated in B–D. Aliquots of CLASP1-repressed and CLASP1-repressed and TAT-PRC1-(438–620)-treated cells were also included for control. Note that synergistic effect was seen in CLASP1-repressed and TAT-PRC1-(438–620)-treated cells. For quantification, the anaphase-defected cells were expressed as a percentage of mitotic cells (mean ± S.E., n = 100 from three different preparations (p < 0.01 compared with TAT-GFP treatment).

Live Cell Imaging

HeLa cells grown on coverslips stably expressed mCherry-tubulin. After appropriate treatment, the cells were observed with a DeltaVision system (Applied Precision, Issaquah, WA).

Quantification of Microtubule Bundles in the Central Spindle

To evaluate the functional relevance of PRC1-CLASP1 interaction in the central spindle organization, HeLa cells stably expressing mCherry-tubulin were synchronized and subjected to TAT-PRC1-(438–620) treatment before imaging. Optical sections taken from the middle of the z-stack were scored for the number of overlapped microtubule bundles in the TAT-PRC1-(438–620)-treated cells. About 10 cells of each condition from three different preparations were scored and subjected to statistical analyses.

Quantification of Stable Microtubules in the Central Spindle

To evaluate the functional relevance of PRC1-CLASP1 interaction in stabilizing the central spindle microtubule, HeLa cells stably expressing mCherry-EB3 were synchronized and subjected to TAT-PRC1-(438–620) treatment before imaging. Optical sections taken from the middle of the z-stack were scored for the number of mCherry-EB3-marked microtubule plus-ends in the TAT-PRC1-(438–620)-treated cells. To quantify the number of EB3-marked microtubule plus-ends, we selected the region of overlapped central spindle microtubule (middle one-third of the spindle axis; boxed area in Fig. 6J). About nine cells of each condition from three different preparations were scored and subjected to statistical analysis.

RESULTS

CLASP1 Interacts with PRC1 in Vitro and in Vivo

Our early attempt to dissect the central spindle revealed a complex of PRC1-CLASP1.⁵ The distribution of CLASP1 within the anti-parallel inter-digitating microtubules in anaphase is reminiscent of PRC1 distribution in anaphase cells (15). To examine whether CLASP1 interacts with PRC1 in vivo, we co-transfected 293T cells with GFP-CLASP1 and 3×FLAG-PRC1 and immunoisolated FLAG-PRC1 protein and its accessory proteins with an anti-FLAG M2 affinity gel. Immunoblotting with an anti-GFP antibody confirmed the presence of GFP-CLASP1 in FLAG immunoprecipitates (Fig. 1A, lane 6). However, CLIP170, a binding partner of CLASP1, was not recovered from the FLAG immunoprecipitates, demonstrating the specificity of the CLASP1-PRC1 complex.

FIGURE 1.

CLASP1 interacts with PRC1 in vitro and in vivo. A, co-immunoprecipitates with anti-FLAG antibody. 293T cells were co-transfected with 3×FLAG-PRC1 and pEGFP-CLIP-170 or pEGFP-CLASP1. A control was performed using cell lysates from GFP-transfected 293T. Cell extracts were subjected to immunoprecipitation (Immunoprep.) using anti-FLAG antibody, fractionated by SDS-PAGE, and immunoblotted with anti-GFP antibody. Immunoprecipitates contained both 3×FLAG-PRC1 and pEGFP-CLASP1 (lane 6). Lanes 3 and 5 represent the supernatants after incubating with anti-FLAG antibody. The common band in the immunoprecipitates with anti-GFP antibody is antibody-derived protein. In the experiment shown in B, Western blotting verifies co-immunoprecipitation of PRC1 and CLASP1. S, supernatant; P, pellet. B, schematic representation of CLASP1 structure and a series of deletion mutant constructs used in immunofluorescence assay. Compared with CLASP1α isoform, CLASP1 lacks 249 amino acids (aa) on the amino end. N′-terminal region of CLASP1 contains the domain that is responsible for the interaction of CLASP1 with microtubules. The C′-terminal of CLASP1 interacts with CLIPs, the Golgi apparatus, or the cell cortex. C, co-immunoprecipitation of PRC1 and CLASP1 from mitotic cells. Extracts from nocodazole-arrested mitotic cells were incubated with antibodies against PRC1 (lanes 1 and 2) and CLASP1 (lanes 4 and 5), and immunoprecipitates (i.p.) were resolved by SDS-PAGE. Lane 3, immunoprecipitation from PRC1 IP; lane 6, immunoprecipitation from CLASP1 IP. Western blotting verified co-immunoprecipitation of CLASP1 (upper panel, CLASP1 blot) and PRC1 (middle panel). Purified rabbit IgGs were included as for the control immunoprecipitation experiment in which neither CLASP1 nor PRC1 was precipitated (lane 9). Note that tubulin was not pulled down by either PRC1 antibody (lower panel, lane 3) or CLASP1 antibody (lower panel, lane 6). D, GST pulldown assay with GST-PRC1. Full-length GST-PRC1 recombinant proteins purified on glutathione-agarose beads were used as an affinity matrix for absorbing GFP-tagged CLASP1 and its deletions as described under “Materials and Methods.” GST protein-bound agarose beads were used as a control. 293T cell extracts were used. Proteins, retained by glutathione-Sepharose beads decorated with GST or with wild type GST-PRC1, were analyzed by Western blotting. After washing, proteins bound to agarose beads were fractionated on an SDS-polyacrylamide gel followed by transferring onto a nitrocellulose membrane. Western blotting is shown with anti-GFP antibodies for the GFP fusions. GFP antibody reacts with full-length CLASP1 (lane 6), N1003 (lane 7), and N694 (lane 8) fusion proteins from wild type PRC1 pulldown. These results suggest that CLASP1 and two N′-terminal deletions can bind to PRC1 in vitro. E, GST pulldown assay with the indicated GST fusions. GST-tagged full-length and mutants of PRC1 proteins purified on glutathione-agarose beads were used as affinity matrix for absorbing GFP-tagged CLASP1. Western blotting analyses with an anti-GFP antibody are shown for the GFP proteins and anti-GST antibody for the GST fusions. GFP antibody reacts with a 170-kDa protein band of full-length CLASP1 fusion proteins from GST-PRC1 (lane 3), C-terminal (lane 5), phospho-mimicking mutants (lanes 7 and 9), and nonphosphorylatable mutants (lanes 6 and 8) of PRC1 pulldowns, but not GST pulldown (lane 10), indicating that both full-length and C-terminal deletion mutant PRC1 can absorb CLASP1 protein. F, schematic illustration showing pEGFP-tagged CLASP1 and a series of deletion mutant constructs used in the co-immunoprecipitation assays and pulldown assays. PRC1 binding was determined on the basis of GST pulldown data as follows: +, detectable; −, undetectable.

To validate if CLASP1 forms a cognate complex with PRC1, we carried out an anti-PRC1 immunoprecipitation assay. As shown in Fig. 1C, anti-PRC1 antibodies pulled down PRC1 protein from HeLa cell lysates (lane 3). As predicted, CLASP1 protein was presented in the anti-PRC1 immunoprecipitates (lane 6). However, tubulin was not precipitated, validating the specific association of CLASP1 with PRC1. Thus, we conclude that PRC1 forms a complex with CLASP1 in vivo.

To test if CLASP1 physically interacts with PRC1, we performed an in vitro pulldown assay using GST-PRC1 as an affinity matrix to isolate CLASP1 and its deletion mutants from lysates of 293T cells transiently transfected to express GFP-CLASP1 wild type and deletion mutant proteins (Fig. 1B). As shown in Fig. 1D (lanes 6–8), the GST pulldown assay showed that GFP-CLASP1 and its two N-terminal deletions GFP-N1003 and GFP-N694 were specifically absorbed by GST-PRC1, suggesting that PRC1 interacts with CLASP1 via its N terminus.

Our recent studies show that mitotic phosphorylation of PRC1 at Thr-470 is required for PRC1 oligomerization and proper central spindle organization (15). To test whether mitotic phosphorylation of PRC1 at Thr-470 and Thr-481 modulates PRC1-CLASP1 interaction, we expressed phospho-mimicking and nonphosphorylatable PRC1 mutant proteins and used them to absorb the full-length CLASP1 protein. As shown in Fig. 1E, wild type and all phospho-mimetic PRC1 (lanes 6–9) mutants bind to CLASP1 equally well. Thus, we conclude that PRC1 physically interacts with CLASP1, and this interaction is not modulated by phosphorylation at Thr-470 and/or Thr-481.

CLASP1 Co-distributes with PRC1 to the Central Spindle during Mitosis

Given the physical interaction between CLASP1 and PRC1 established in the early studies (Fig. 1), we sought to examine their spatiotemporal distribution profiles and inter-relationship during cell division. To this end, HeLa cells were fixed and stained with antibodies against PRC1 and CLASP1. Both PRC1 and CLASP1 display a typical spindle-like distribution pattern in prometaphase and metaphase cells (Fig. 2A, panels a and b). Compared with that of PRC1, CLASP1 localization to the spindle pole is apparent in metaphase cell (Fig. 2A, panel b). During metaphase-anaphase transition, the majority of PRC1 is relocated to the central spindle, whereas the majority of CLASP1 remains associated with spindle and spindle pole (Fig. 2A, panel c). In anaphase B cells, the majority of CLASP1 relocates to the central spindle where it is superimposed onto that of PRC1 (Fig. 2A, panel d). From telophase to cytokinesis, both proteins are concentrated at the Flemming body (Fig. 2A, panels e and f). The double immunofluorescence study revealed that PRC1 is assembled onto the central spindle prior to the arrival of CLASP1.

FIGURE 2.

PRC1 determines the central spindle localization of CLASP1. A, co-localization of endogenous CLASP1 and PRC1 on the mitotic spindle, spindle midzone, and midbody during mitosis is shown. In prometaphase and metaphase cells, both CLASP1 (red) and PRC1 (green) localize with the entire mitotic spindle (panels a and b). Then both proteins are concentrated as a series of narrow bars at the late mitosis spindle midzone (panels c–e). As cells progress through cytokinesis, CLASP1 and PRC1 are concentrated to the midbody, where the two proteins persist even after the completion of cytokinesis (panel f). Scale bars, 10 μm. B, co-localization of various exogenous GFP-CLASP1 proteins and PRC1 during late mitosis. For each construct, anaphase (panels a and c) and cytokinesis (panels b and d) cells are shown, stained for PRC1 (red) and α-tubulin (blue) to verify the position of the mitotic spindle. Removal of the C-terminal of CLASP1 (CLASP1-(1–1003) (CLASP1^1∼1003)) does not affect its ability to localize on central spindle in anaphase (a) nor midbody during telophase and cytokinesis (panel b). Deletion of the central region of CLASP1 (e.g. CLASP1-(1–694) (CLASP1^1∼694)) failed to localize to the central spindle in anaphase (panel c), and the midbody during telophase and cytokinesis (panel d). Scale bars, 10 μm.

PRC1-CLASP1 Binding Is Necessary for CLASP1 Distribution to the Central Spindle

If PRC1 specifies the localization of CLASP1 to the central spindle, the region where CLASP1 binds to PRC1 should be localized to the central spindle. To test this hypothesis, we generated a series of CLASP1 deletion mutants tagged with GFP. HeLa cells were transiently transfected to express various CLASP1 deletion mutant proteins and examined for their co-localization with PRC1 and tubulin. As shown in Fig. 2B, GFP-CLASP1-(1–1003) co-distributes with PRC1 to the spindle midzone in anaphase and to the midbody during telophase and cytokinesis (panels a and b). On other hand, GFP-CLASP1-(1–694), previously reported to exhibit the microtubule binding activity (17), distributes along with the entire length of microtubules, neither localizing to the central spindle nor to the midbody (Fig. 2B, panels c and d). The GFP-CLASP1-(1–1003) distribution relative to PRC1 in anaphase together with their biochemical interaction established, as shown in Fig. 1, that PRC1 is required for the central spindle localization of CLASP1.

PRC1 Is Essential for Accurate Localization of CLASP1 to the Central Spindle

If PRC1 is required for the localization of CLASP1 to the central spindle, suppression of PRC1 would result in an aberrant localization of CLASP1 in anaphase. To test this hypothesis, HeLa cells were transfected with siRNA oligonucleotides to suppress PRC1. Suppression of PRC1 was successfully achieved by introducing siRNA oligonucleotide into target cells judged by Western blotting analysis (Fig. 3A, top panel) without alteration of CLASP1 protein levels (middle panel). Quantitative Western blotting analyses show that treatment of 25 nm siRNA oligonucleotide resulted in a 93 ± 2% suppression of the PRC1 protein. Immunofluorescence analysis validated the efficacy of the PRC1 protein repression as virtually no PRC1 protein was found in the siRNA-treated cells (Fig. 3B, panels a–c).

FIGURE 3.

Repression of PRC1 eliminates CLASP1 localization to central spindle. A, efficiency of RNA interference of PRC1. HeLa cells were transfected with the PRC1 siRNA oligonucleotide (25 nm) or a mock siRNA oligonucleotide and subjected to SDS-PAGE and immunoblotting analyses. Cell extracts were probed with anti-PRC1 antibody (top panel), anti-CLASP1 antibody (middle panel), and anti-tubulin antibody as internal controls (lower panel). B, HeLa cells grown on coverslips were transfected with PRC1 siRNA (panels a–c) or scramble siRNA control (panels a′–c′). 24 h after transfection, cells were fixed and stained with anti-PRC1 antibody (green), anti-tubulin antibody (red), and DAPI (blue). PRC1-deficient cells exhibited a typical phenotype with segregated sister chromatids and disjoined two spindle halves (panels a and b, arrows, arrowheads). Scale bars, 10 μm. C, HeLa cells, stably expressing EGFP-CLASP1, grown on coverslips were transfected with 25 nm PRC1 siRNA oligonucleotide. 24 h after transfection, cells were fixed and stained with anti-PRC1 antibody (red) and DAPI (blue). Compared with the controls (panels a′–c′), repression of PRC1 protein by siRNA treatment resulted in little distribution of CLASP1 to the central spindle and the midbody (panels a–c), indicating that PRC1 is required for CLASP1 distribution to the central spindle (panels b and b′, arrows) and the midbody during anaphase and telophase. Scale bars, 10 μm. D, HeLa cells stably expressing EGFP-CLASP1 were transfected with 25 nm PRC1 siRNA oligonucleotide. 24 h after transfection, cells were fixed and stained with anti-tubulin antibodies (red) and DAPI (blue). Repression of PRC1 failed to form inter-digital microtubule bundles (panels b and b′; arrows). Scale bars, 10 μm.

As shown in Fig. 3B, the central spindle localization of PRC1 was not altered by the scramble siRNA oligonucleotide treatment (panels a′, b′, and c′). However, in these PRC1-repressed anaphase cells, anti-parallel microtubule organization was perturbed as spindle microtubules from two opposite poles are no longer overlapped (Fig. 3B, panels a and b, arrowheads). The two half-spindles remained disconnected in those PRC1-repressed HeLa cells. Examination of DNA staining revealed an aberrant chromatid segregation (Fig. 3B, panel a, arrows), consistent with previous studies (28). PRC1 suppression did not cause gross change of CLASP1 localization along the mitotic spindle (Fig. 3C, panels a–c). However, repression of PRC1 abolished the overlapping of CLASP1 bundles from anti-parallel configuration (Fig. 3C, arrows, panels a and b). In addition, repression of PRC1 results in disappearance of CLASP1 in the Flemming body (Fig. 3C, panel c) as the integral central spindle microtubules could not be formed in the absence of PRC1. Double labeling of CLASP1 and tubulin revealed that PRC1 is essential for formation of the inter-digital microtubule bridge (Fig. 3D, panels a and b). Thus, we concluded that PRC1 specifies the central spindle localization of CLASP1 during anaphase.

CLASP1 Depletion Causes Cell Cycle Delay and Mitotic Abnormalities

The subcellular distribution of CLASP1 and its physical interaction with PRC1 prompted us to examine if CLASP1 functions in spindle integrity and chromosome dynamics. To this end, we suppressed the CLASP1 protein in HeLa cells by using siRNA oligonucleotide followed by phenotype-based characterization. Quantitative Western blotting analyses indicated that siRNA oligonucleotide suppressed the protein level of CLASP1 up to 70% at 48 h post-transfection (Fig. 4A, top panel). Control siRNA did not alter CLASP1, tubulin, or PRC1 protein levels (Fig. 4A).

FIGURE 4.

CLASP1 is essential for faithful chromosome segregation and mitotic spindle assembly. A, efficiency of RNA interference of CLASP1. Immunoblot analysis of HeLa cells treated with the CLASP1 siRNA oligonucleotide (25 nm) or with a mock siRNA oligonucleotide is shown. Cell extracts were probed with anti-CLASP1 antibody (top panel), anti-PRC1 antibody (middle panel), and anti-tubulin antibody as internal controls (bottom panel). B, repression of CLASP1 resulted in mitotic abnormalities. HeLa cells, stably expressing mCherry-tubulin (red), grown on coverslips were transfected with 25 nm CLASP1 siRNA. 36–48 h after transfection, cells were fixed and stained for DNA (DAPI, blue) and CLASP1 (green). As shown in C and D, compared with control cells (panels a′–c′), CLASP1-depleted cells exhibit anaphase chromatid separation errors (panels a and b, arrows) and had difficulty in chromosome alignment (panel a, arrow). The cells also entered in anaphase with abnormal spindle (panels b and c) and a chromatin bridge (panel c, arrows). Scale bars, 10 μm. C, statistical analyses of cells exhibiting mitotic abnormalities, such as misaligned chromosomes and multipolar spindle, in CLASP1-repressed metaphase cells (black bar) and control preparation (open bar). An average of 100 metaphase-like cells from three separate experiments was counted for HeLa cells that expressed mCherry-tubulin stably and were treated with CLASP1 siRNA oligonucleotide (36–48 h post-transfection). Cells were then examined under a fluorescence microscope, and associated phenotypes were categorized and quantified as the percentage of total mitotic cells. D, statistical analyses of cells exhibiting chromatid bridges or multipolar spindles in CLASP1-deficient anaphase cells (black bar) and control preparation (open bar). An average of 100 anaphase-like cells from three separate experiments was counted after being transfected with CLASP1 siRNA and scramble oligonucleotides (36–48 h post-transfection). Cells were then examined under a fluorescence microscope, and associated phenotypes were categorized and quantified as the percentage of total mitotic cells.

For accurate analysis of the phenotype associated with the repression of CLASP1 activity, we examined the chromosome segregation and mitotic spindle plasticity in tubulin-marked HeLa cells treated with CLASP1 siRNA oligonucleotide under a fluorescence microscope. Knockdown of CLASP1 induced an anaphase chromatid bridge phenotype and disorganized central spindles (Fig. 4B, panels a–c, arrows). We have scored 300 cells from four different preparations, and our quantification shows that the percentage of cells containing misaligned chromosomes and premature anaphase chromatid bridges was significantly greater than those of control cells (Fig. 4, C and D). As shown in Fig. 4C, the chromosome misalignment rate was significantly increased from 3 ± 1% of the control to 17 ± 3% of CLASP1-repressed cells (p < 0.01). The percentage bearing anaphase bridge was increased from 2 ± 1% of the scramble control to 31 ± 5% in CLASP1-suppressed cells (p < 0.01), supporting the notion that CLASP1 is essential for chromosome stability in mammalian cell division.

PRC1-CLASP1 Interaction Is Required for Faithful Mitosis

The functional importance of CLASP1 in mitotic plasticity and PRC1-CLASP1 inter-relationship prompted us to examine the functional relevance of PRC1-CLASP1 interaction in mitosis. To pinpoint the region(s) of PRC1 that binds directly to CLASP1, we generated a series of deletion mutants of PRC1 (illustrated in Fig. 5A) tagged with GST and used glutathione-bound GST PRC1 proteins as affinity matrices. Because our attempt to obtain recombinant full-length CLASP1 in bacteria was unsuccessful, we used cell lysates of 293T cells transiently transfected to express GFP-CLASP1 to incubate with GST-PRC1 affinity matrix. As shown in Fig. 5B, GFP-CLASP1 binds to full-length PRC1 and the C-terminal PRC1-(438–620) judged by GFP antibody blotting analysis (Fig. 5B, upper panel). Neither GST protein nor GST-PRC1-(1–437) absorbed GFP-CLASP1, suggesting that the interaction between CLASP1 and C-terminal 183 amino acids of PRC1 is specific. This binding assay also revealed weak CLASP1 binding activities in the N-terminal deletion mutants PRC1-(438–485) and PRC1-(486–620) (Fig. 5B, lanes 5 and 6).

FIGURE 5.

PRC1-(438–620) involves association of PRC1 with CLASP1 in vitro and in vivo. A, schematic diagram of the functional domains of PRC1. Phosphorylation sites (asterisks) correspond to residues Thr-470, Thr-481, Thr-578, and Thr-616. B, PRC1 binds to CLASP1 via its C-terminal region. Both full-length and deletion mutant GST-PRC1 proteins were expressed and purified using glutathione-agarose beads. Purified GST-PRC1 proteins on Sepharose beads were used as an affinity matrix for absorbing GFP-CLASP1. Retention of GFP-CLASP1 proteins on GST-PRC1 affinity beads was validated by immunoblotting using anti-GFP antibodies. Western blotting analyses revealed that the C-terminal domains of PRC1 (PRC1-(438–620) (PRC1^438∼620)) are responsible for its CLASP1 binding activity (lanes 3, 5, 6, and 7). C, expression and purification of TAT-GFP fusion proteins in E. coli. Molecular size of protein is indicated on the left. D, addition of PRC1-(438–620) recombinant protein disrupts CLASP1-PRC1 association. Purified GST-PRC1 protein on glutathione-Sepharose beads was used as an affinity matrix for absorbing GFP-CLASP1. After extensive washes, aliquots of CLASP1-bound GST-PRC1-Sepharose beads were incubated with 5 μm purified PRC1-(438–620) protein on a rotator for 30 min at room temperature. After extensive washes, the Sepharose beads were boiled in SDS-PAGE sample buffer, and bound proteins were fractionated by SDS-PAGE followed by visualizing in Coomassie Blue stain. Note that incubation of TAT-GFP-PRC1-(438–620) peptide with CLASP1-bound GST-PRC1-Sepharose beads disrupts PRC1-CLASP1 interaction. E, HeLa cells synchronized by mitotic shake off were extracted with 0.1% Triton X-100 solution and separated into soluble (hatch bars) and insoluble (black bars) fractions as described under “Materials and Methods.” Equivalent amounts of protein from the soluble and insoluble fractions were applied to SDS-PAGE. Following separation on 6–16% gradient SDS-PAGE and transblotting onto nitrocellulose membrane, the blots were probed with an anti-PRC1 antibody and an anti-CLASP1 antibody followed by visualization using an ECL kit (Pierce). The signals were quantified using a PhosphorImager with values expressed as a percentage of total (soluble + insoluble). The error bars represent S.E.; n = 3 preparations. F, PRC1-derived peptide TAT-GFP-PRC1-(438–620) increases the extractable CLASP1 proteins. HeLa cells synchronized by mitotic shake off were treated with the TAT-GFP-PRC1-(438–620) peptide followed by a Triton X-100 extraction. The cellular proteins were separated into a soluble and insoluble fraction as described previously (24). Samples were run on SDS-PAGE, blotted to nitrocellulose, and immunoprobed for CLASP1. The signals were quantified using a PhosphorImager with values expressed as percentage of total (soluble + insoluble). Error bars represent S.E.; n = 4. Significant difference is found with respect to the controls (*, p < 0.01).

To explore the functional effect of CLASP1-PRC1 interaction in the mitotic process, we sought to introduce membrane-permeable peptide containing PRC1-(438–620) to modulate endogenous PRC1-CLASP1 interaction. This is achieved by introducing an 11-amino acid peptide derived from TAT protein transduction domain (29, 30) into GFP-PRC1-(438–620) fusion protein. The recombinant protein was histidine-tagged and purified to homogeneity using nickel-affinity beads (Fig. 5C). As predicted, incubation of recombinant GFP-PRC1-(438–620) protein (5 μm) with GFP-CLASP1-bound GST-PRC1 affinity matrix disrupts the CLASP1-PRC1 association in vitro (Fig. 5D, lane 3). Neither GFP nor TAT-GFP interferes with CLASP1-PRC1 interaction (Fig. 5D, lanes 1 and 2), demonstrating the effect of PRC1-(438–620) in competing full-length PRC1 for CLASP1 association.

If recombinant TAT-GFP-PRC1-(438–620) fusion protein disrupts endogenous PRC1-CLASP1 interaction, addition of TAT-GFP-PRC1-(438–620) recombinant protein into HeLa cell culture would compete for endogenous PRC1 and liberate CLASP1 into the cytoplasmic soluble fraction. To determine whether there were any major changes in the behavior of endogenous PRC1 and CLASP1 in response to the addition of GFP-PRC1-(438–620), we measured the partitioning of endogenous PRC and CLASP1 into the Triton X-100-soluble fraction compared with the insoluble “cytoskeletal” fraction based on the Western blotting analyses. As summarized in Fig. 5E, 71.7 ± 4.3% of endogenous PRC1 resides in the Triton X-100-insoluble fraction. Partitioning of CLASP1 is similar to endogenous PRC1; 69.3 ± 4.1% of endogenous CLASP1 is associated with the Triton X-100-insoluble fraction.

Our trial experiments show that the internalization of TAT-GFP and TAT-GFP-PRC1 proteins was seen in virtually all cells 30 min after the addition of TAT-GFP fusion proteins in culture. Our quantitative blotting analyses show that addition of TAT-GFP-PRC1-(438–620) induced a dose-dependent liberation of CLASP1 into cytoplasmic “soluble” fraction. Although TAT-GFP was internalized as effectively as TAT-GFP-PRC1-(438–620), TAT-GFP did not alter the partition of endogenous CLASP1 protein in the cytoplasmic soluble and cytoskeletal insoluble fraction (Fig. 5F). Although 70.3 ± 4.5% of endogenous CLASP1 resides in the Triton X-100-insoluble fraction of TAT-GFP-treated cell culture, only 33.7 ± 7.7% of endogenous CLASP1 is associated with the Triton X-100-insoluble fraction in 500 nm TAT-GFP-PRC1-(438–620)-treated cells (p < 0.01), indicating that TAT-GFP-PRC1-(438–620) fusion protein disrupts endogenous PRC1-CLASP1 interaction in vivo. In contrast, there was no significant difference in the amount of EB1 and tubulin extracted from control or PRC1-(438–620) peptide-treated cells (data not shown).

PRC1 Cooperates with CLASP1 to Orchestrate Central Spindle Plasticity and Chromosome Stability

Having demonstrated the ability of TAT-GFP-PRC1-(438∼620) in perturbing endogenous PRC1-CLASP1 interaction, we sought to test whether the PRC1-CLASP1 interaction is specifically required for central spindle plasticity and dynamics. To overcome the potential pre-anaphase block because of the perturbation of PRC1-CLASP1 interaction, we adopted a protocol to synchronize cells at metaphase and then added TAT-PRC1 peptide to probe for functional relevance in anaphase onset and beyond (illustrated in Fig. 6A). We began real time imaging of metaphase-synchronized HeLa cells expressing mCherry-tubulin. TAT-GFP-PRC1-(438–620) was added into metaphase cells, and time-lapse imaging was started as the cells underwent anaphase-telophase transition. Our trial experiments show that treatment of S and G₂ phase HeLa cells with TAT-GFP-PRC1-(486–620) (500 nm) did not alter cell cycle progression into mitosis or sister chromatid separation (Fig. 6C) as compared with those treated with TAT-GFP control (Fig. 6B). In general, it takes an average of 3103.3 ± 448.5 s (n = 8 cells) for HeLa cells to transit from centrosome separations to the anaphase onset of sister chromatid separation, which is similar to that of TAT-GFP (500 nm)-treated cells (average 2855.8 ± 388.7 s; n = 8 cells; Fig. 6E).

However, treatment of mitotic cells with TAT-GFP-PRC1-(438–620) (500 nm) protein perturbed central spindle assembly plasticity and prolonged the anaphase duration. On average, it takes about 700.6 ± 27.0 s (n = 8 cells) for TAT-GFP-PRC1-(438–620)-treated HeLa cells to transit from the formation of anti-parallel overlapped microtubule organization to midbody formation. However, it takes about 621.4 ± 16.5 s (n = 8 cells; p < 0.01) for control HeLa cells treated with TAT-GFP to complete the aforementioned process (Fig. 6F). Careful examination of time-lapse imaging revealed that the delay took place before the formation of contractile ring (Fig. 6C, asterisks).

To investigate whether the delay in anaphase progress observed in TAT-GFP-PRC1-(486–620)-treated cells was because of the disturbance of endogenous PRC1 or that of CLASP1, we knocked down PRC1 and CLASP1, respectively. Down-regulation of CLASP1 had a detectable prolongation on anaphase duration (on average 1079.0 ± 92.5 s versus 603.5 ± 22.0 s in control cells). The duration from central spindle assembly to contraction ring formation has been extended more than 78.9%. However, suppression of PRC1 by siRNA did not change the anaphase duration (on average 641.0 ± 27.8 s versus 603.5 ± 22.0 s in control cells; Fig. 6G). To test if synergism can be achieved when CLASP1 repression is combined with TAT-PRC1-(438–620) peptide, synchronized HeLa cells were transfected with CLASP1 siRNA followed by incubation of TAT-PRC1-(438–620) peptide as briefed above. As shown in Fig. 6D, a combination of CLASP1 knockdown with perturbation of PRC1-CLASP1 interaction dramatically disrupted the number of overlapped central spindle microtubule bundles in addition to a prolonged anaphase onset, a phenotype seen in TAT-PRC1-(438–620) peptide-treated HeLa cells. Quantitative analyses show that perturbation of PRC1-CLASP1 interactions disrupted the central spindle plasticity as the number of overlapped anti-parallel microtubules is dramatically reduced (Fig. 6I). The number of overlapped microtubules was reduced from an average of 10 ± 2 of control anaphase A cells to an average of 6 ± 1 of TAT-PRC1-(438–620)-treated cells (p < 0.01). A combination of siRNA-mediated repression of CLASP1 with TAT-PRC1-(438–620) treatment further reduced the number of overlapped microtubule bundles (5 ± 2 in anaphase A; p < 0.01). Similar profile was also observed in anaphase B cells in the presence of TAT-PRC1-(438–620) treatment.

To further evaluate the functional relevance of PRC1-CLASP1 interaction in stabilizing the central spindle microtubule, we sought to examine the EB3 distribution in the anaphase central spindle microtubule in the presence of TAT-PRC1-(438–620) peptide treatment. To this end, HeLa cells stably expressing mCherry-EB3 were synchronized and subjected to TAT-PRC1-(438–620) treatment before imaging. Optical sections taken from the middle of the z-stack were scored for the number of mCherry-EB3-marked microtubule plus-ends in the TAT-PRC1-(438–620)-treated cells. EB3 is a plus-end tracking protein that marks the growth and stability of the microtubule. To quantify the number of EB3-marked microtubule plus-ends, we selected the region of overlapped central spindle microtubule (boxed area in Fig. 6J). The statistical analysis demonstrated that perturbation of PRC1-CLASP1 greatly reduced the stability of central spindle microtubules (p < 0.01; compared with that of TAT-GFP-treated cells).

To elucidate the role of CLASP1-PRC1 interaction in the central spindle plasticity and chromosome stability, we carried out immunofluorescence study of mitotic cells treated with TAT-GFP-PRC1-(438–620). As shown in Fig. 6H, perturbation of the PRC1-CLASP1 interaction resulted in chromosome lagging (panel b) and formation of a sister chromatid bridge (panel c), a characteristic phenotype associated with chromosome loss in cell division cycle. This phenotype is rarely seen in TAT-GFP-treated cells (Fig. 6H, panels b″–d″). We have also included CLASP1 siRNA treatment and siRNA treatment plus TAT-PRC1-(438–620) peptide treatment. The typical phenotype seen in PRC1-CLASP1 interaction-disrupted cells is sister chromatid bridges. The quantification from three different preparations was presented in Fig. 6K. Thus, we conclude that PRC1-CLASP1 interaction governs central spindle plasticity and dynamics that are essential for faithful chromosome segregation in mitosis.

DISCUSSION

Our studies reveal a key role for the PRC1-CLASP1 interaction in achieving the stable anti-parallel microtubule organization essential for faithful chromosome segregation. Our characterization demonstrates that the interaction between PRC1 and CLASP1 is essential and sufficient for the localization of CLASP1 to the central spindle. Given the disorganization of central spindle seen in CLASP1-repressed and CLASP1-PRC1 interaction-perturbed cells, we reason that PRC1 specifies the central spindle localization of CLASP1, and CLASP1 is essential for central spindle plasticity and genomic stability in mitosis.

The spindle midzone, a highly organized network of anti-parallel inter-digitating microtubules between separating chromosomes, plays a crucial role in regulating the initiation and completion of cytokinesis. Cytokinesis is a highly orchestrated process that must coordinate precisely with other events of the cell cycle via coupling cytokinesis to Cdc2 inactivation. Down-regulation of Cdc2 activity initiates the accumulation of some proteins in the midline of central spindle. These proteins include MKLP1, MKLP2, MgcRacGAP, PRC1, etc. PRC1 is phosphorylated on Thr-470 and Thr-481 by Cdc2 in early mitosis. In fact, our early study showed that phosphorylation of PRC1 at Thr-470 promotes the formation of PRC1 oligomers (15), which may promote the assembly of PRC1 protein onto overlapping microtubules at the central spindle (31). The fact that phosphorylation of Thr-470 and Thr-480 is not involved in PRC1-CLASP1 interaction suggests that central spindle plasticity is orchestrated by several interactive but perhaps parallel pathways. To this end, it would be of great interest to use super-resolution imaging microscopic analyses to delineate how spatiotemporal control of central spindle assembly is achieved at a finely tuned fashion. In addition, it would also be important to address whether other mitotic phosphorylation sites, other than Thr-470 and Thr-481, at the C-terminal PRC1 regulate the PRC1-CLASP1 interaction (32).

It has been shown that multiple microtubule-associated proteins act in concert to orchestrate cytokinesis. In anaphase cells, many of these proteins establish and/or associate with anti-parallel central spindle microtubules. In addition to interacting with CLASP1 and KIF4 in setting up the central spindle, PRC1 also interacts with and recruits KIF14 to the central spindle (33). Further experimentation will be required to address how multiple PRC1 protein complexes and PRC1 oligomers coordinate in a spatiotemporal manner in establishing and/or maintaining central spindle organization and dynamics. In this regard, selective perturbation of PRC1 association with its binding partners will be essential for delineating the function of specific molecular interaction between PRC1 and its interaction proteins and the molecular pathway of PRC1-mediated central spindle organization. We propose that PRC1 forms a link between stabilization of CLASP1 association with central spindle microtubules and anti-parallel microtubule elongation.

It has been shown that human CLASP1 is an outer kinetochore component that regulates spindle microtubule dynamics (21). Recent studies show that improper microtubule-chromosome attachment (merotely) is a cause of chromosome mis-segregation in unstable cells and that increasing chromosome mis-segregation rates by elevating merotely during consecutive mitoses generates chromosome instability in otherwise stable, near-diploid cells (34, 35). This chromosomal instability is commonly caused by persistent mal-oriented attachment of chromosomes to spindle microtubules. Chromosome segregation requires stable microtubule attachment at kinetochores, yet those attachments must be sufficiently dynamic to permit correction of mal-orientations. To explore how this balance is achieved, Thompson and Compton (34) studied the permissible boundaries of attachment stability versus dynamics essential for genome stability. They identified two microtubule-depolymerizing kinesins that stimulate kinetochore-microtubule dynamics during distinct phases of mitosis to correct mal-orientations. These studies indicate that temporal control of microtubule attachment to chromosomes during mitosis is central to genome stability in human cells. Given the fact that repression of CLASP1 and perturbation of PRC1-CLASP1 interaction resulted in chromosome instability phenotypes such as chromatid bridges (Fig. 4B), it would be of great importance to ascertain if repression of CLASP1 alters the microtubule plus-end dynamics and/or the localization of plus-end tracking such as Kif2b. It would of great importance in the near future to elucidate the functional relevance of CLASP2 in central spindle organization given the homologues seen between CLASP1 and CLASP2.

Taken together, we propose that CLASP1-PRC1 interaction establishes faithful chromatid segregation by specifying central spindle organization and orchestrating kinetochore-microtubule attachment at the kinetochore. It is likely that the kinetochore outer plate protein dynamics is highly coordinated with assembly of the central spindle. The PRC1-CLASP1 interaction established here is a core of this dynamic interaction network that ensures chromosome stability in the cell division cycle.