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[Cancer Research 65, 850-860, February 1, 2005]
© 2005 American Association for Cancer Research

Cell and Tumor Biology

Receptor for Hyaluronan-Mediated Motility Correlates with Centrosome Abnormalities in Multiple Myeloma and Maintains Mitotic Integrity

Christopher A. Maxwell, Jonathan J. Keats, Andrew R. Belch, Linda M. Pilarski and Tony Reiman

Department of Oncology, University of Alberta and Cross Cancer Institute, Edmonton, Alberta, Canada

Requests for reprints: Tony Reiman, Department of Medical Oncology, Cross Cancer Institute, 11560 University Avenue, Edmonton, Alberta, Canada T6G 1Z2. Phone: 780-432-8513; Fax: 780-432-8888; E-mail: tonyreim{at}cancerboard.ab.ca.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Elevated expression of receptor for hyaluronan-mediated motility (RHAMM) within ex vivo diagnostic multiple myeloma plasma cells predicts for aggressive disease and patient survival. Here, we investigate the relationship between RHAMM and centrosomal abnormalities within multiple myeloma patient samples. We report that myeloma patient samples contain pervasive structural and numerical centrosomal abnormalities. Structural, but not numerical, centrosomal abnormalities strongly correlate with elevated RHAMM expression. As others have shown that excess pericentriolar material strongly associates with abnormal mitoses, we modeled centrosomal abnormalities with exogenous RHAMM overexpression. RHAMM overexpression in vitro resulted in centrosomal and mitotic defects. To elucidate a mechanism for RHAMM-mediated spindle defects, we further investigated RHAMM mitotic function. RHAMM mitotic localization mirrors that of targeting protein for Xklp2 (TPX2), and RHAMM interacts with the spindle assembly factors dynein and TPX2. Like TPX2, RHAMM expression is up-regulated during mitosis. Moreover, inhibition of function experiments reveals that RHAMM and TPX2 functions converge to maintain spindle integrity after spindle assembly. We postulate that augmentation of RHAMM expression within human cancers, including myeloma, can directly affect centrosomal structure and spindle integrity and potentially modulate apoptotic and cell cycle progression pathways.

Key Words: multiple myeloma • centrosomes • chromosomal instability • 04-01-17 Hematologic, other


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The receptor for hyaluronan-mediated motility (RHAMM), first described by Turley (1), has cell surface and intracellular distribution and binds hyaluronan (2), ERK kinase (3) and microtubules (4). RHAMM participates in cell motility, signaling (5), and oncogenic events (6). RHAMM also localizes to the centrosome, the microtubule organizing center, and functions in the maintenance of spindle integrity (7). The COOH terminus of RHAMM is essential for centrosomal localization and bears structural similarity to human kinesin-like protein 2, including conservation of a basic leucine zipper domain (8); this motif is essential for the cell cycle–specific interaction between human kinesin-like protein 2 and dynein, which is mediated through targeting protein for Xklp2 (TPX2; ref. 8). Thus, it is intriguing to hypothesize that RHAMM may interact with dynein and TPX2 and function in spindle assembly and integrity.

In multiple myeloma, accumulation of terminally differentiated malignant plasma cells in the bone marrow, despite low plasma cell labeling indices, is suggestive of a clonotypic precursor cell with proliferative potential. RHAMM transcripts are detected in multiple myeloma malignant cells but are weak or absent from normal B cells, bone marrow, or CD138+ plasma cells of control patients (9, 10). MMPCs are characterized by extensive chromosomal instability and elevated RHAMM expression, which is significantly correlated to increased disease-related events and reduced survival (10). Moreover, the prevalence of cytogenetic abnormalities is greater in high, compared with low, RHAMM expressors (10). RHAMM expression is linked to progression and metastasis of a variety of epithelial tumors, including endometrial, stomach, and breast carcinomas (11–13). The association of RHAMM with centrosomes, cell division, and mitotic integrity may explain the correlation observed between RHAMM expression and the progression of malignancies with extensive chromosomal instability.

Centrosomes are composed of two centrioles and an amorphous cloud of pericentriolar material (14). Like RHAMM, many integral components of the pericentriolar material are multiple coiled coil domain proteins that interact with dynein to target centrosomes in a microtubule-independent manner (15). Numerical and/or structural centrosome abnormalities have been reported in a wide range of malignant epithelial tumors with positive correlation to genetic instability and cancer progression (16, 17); to date, however, centrosomal dysregulation has not been investigated within myeloma cells. Centrosomal abnormalities occur early in tumorigenesis (18) with structural (i.e., excess pericentriolar material), rather than numerical, abnormalities most highly associated with abnormal mitoses (16).

Formation of a bipolar spindle is essential to the proper segregation of replicated chromosomes and the maintenance of genetic stability. Spindle assembly can occur through centrosome-dependent and centrosome-independent pathways (19). In some systems, spindle assembly is induced in a Ran-GTP-dependent manner (20, 21). Active Ran, enriched proximal to chromosomes (22), promotes microtubule nucleation through downstream effector proteins, such as TPX2, NuMA, and Aurora A kinase (aurA; refs. 23, 24). TPX2 initiates spindle assembly by nucleating and bundling microtubules (23, 25) and by directly activating aurA in a microtubule-dependent manner (24). The importance of aurA, TPX2, and NuMA has been illustrated in mammalian systems as well (26). Once spindles have been established, their integrity is dependent on the balance of forces generated by microtubule-dependent motors (e.g., dynein). Disruption of this balance, through overexpression or inhibition of structural participants, would be expected to disrupt spindle structure (27).

Because RHAMM is associated with the progression of multiple tumors and, within myeloma, elevated RHAMM is associated with cytogenetic abnormalities, aggressive disease, and shortened survival, we investigated an association between RHAMM and centrosomal abnormalities and the effects of exogenous RHAMM expression on centrosomal and spindle structure. We show that RHAMM gene expression levels correlate with myeloma centrosome volumes and that RHAMM localizes to the centrosome in myeloma plasma cells to a similar extent as other defined centrosome proteins. We show that RHAMM overexpression in vitro induces centrosomal structural abnormalities, similar to those identified in myeloma cells ex vivo, and abnormal spindle architecture. RHAMM interacts with the spindle assembly factors dynein and TPX2, but not NuMA, and dysregulation of RHAMM expression affects G2-M transition and spindle integrity. Based on the work presented here, we postulate that aberrant RHAMM expression in myeloma and other cancers may lead to errors in chromosomal segregation.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Patients and Clinical Data. Bone marrow biopsies from 41 multiple myeloma patients, 8 monoclonal gammopathy of uncertain significance (MGUS, a "premalignant" condition that may precede development of myeloma), and 4 lymphoma patients with uninvolved bone marrow were identified from the pathology records at the Cross Cancer Institute from 1997 to 2000. Bone marrow mononuclear cells were collected from archived aspirates from multiple myeloma patients described above (n = 15), with ≥40% plasma cell infiltration, for quantitative analysis of RHAMM expression. Purified plasma cells were collected from an additional 11 multiple myeloma bone marrow aspirates following CD138, or CD38, selection by fluorescence-activated cell sorting (n = 6 for cytospin analysis of centrosomal RHAMM) or autoMACS Magnetic Cell Sorter [Miltenyi Biotec, Auburn, CA; n = 5 for quantitative reverse transcription-PCR (q-RT-PCR) analysis of RHAMM expression] as described below. All patients consented to the use of their bone marrow samples.

Preparation and Analysis of Multiple Myeloma Bone Marrow Core Biopsies. Bone marrow core biopsies were fixed, decalcified, and paraffin embedded. Sections (4 µm) were mounted on sialinized glass slides. For antigen retrieval, slides were placed in EDTA (1.0 mmol/L, pH 8.0), preheated to 100°C, and placed in a temperature-controlled microwave (TTMega) for 10 minutes. Biopsies were sequentially stained with {gamma}-tubulin (1:200), anti-mouse Alexa 594 (1:200), mouse IgG for blocking, and CD138-FITC (1:20) and mounted with glycerol medium containing 4',6-diamidino-2-phenylindole (DAPI) for DNA staining.

Image Analysis and Three-dimensional Volume Rendering. Following immunofluorescence, confocal z-slices were imaged from core biopsies. For all analysis, confocal slices were imaged at defined zoom (40x objective, 8 zoom), speed, and depth (0.2 µm) and were acquired using a Zeiss confocal LSM 510 or multiphoton microscope. In general, three confocal stacks, including at least 30 plasma cells, were collected. Centrosomes were identified by {gamma}-tubulin immunofluorescence, and regions of interest, containing individual centrosomes, were extracted using Zeiss 510 image analysis software. Images were analyzed with Imaris 3.2.2 software (Bitplane AG, Zurich, Switzerland). Centrosomes were manually counted for at least 80 CD138+ plasma cells per patient; plasma cells with ≥3 centrosomes were considered abnormal. Abnormal centrosomal structure was qualitatively assessed using previously published variables (17). For determination of centrosomal fractions, confocal images were obtained which included the centrosomal region. Regions of interest were defined around the centrosome and the whole cell body, and the intensity within the centrosomal regions of interest was divided by the overall cellular intensity.

q-RT-PCR Assays of Myeloma Bone Marrow Aspirates. Bone marrow mononuclear cell isolation, CD138 selection, and q-RT-PCR were done as described previously to measure RHAMM mRNA levels in multiple myeloma patient bone marrow samples (10). The expression level of each sample was normalized to the sample with the lowest level of RHAMM expression, which was set to an expression level of 1.

Antibodies and Plasmids. Staining used {alpha}-tubulin (clone B-5-1-2) and {gamma}-tubulin (clone GTU-88, Sigma, St. Louis, MO), ß-actin (Sigma), pericentrin (Covance, Richmond, CA), CD138-FITC (Serotec, Raleigh, NC), CD38-PE (BD Biosciences, Palo Alto, CA), and CD45-FITC (Beckman Coulter, Fullerton, CA). The polyclonal RHAMM antibody was produced and characterized as described (8). A second polyclonal RHAMM antiserum was kindly provided by V. Assmann (St. Thomas' Hospital, London, United Kingdom; ref. 4). TPX2 antiserum was kindly provided by O. Gruss (EMBL, Heidelberg, Germany; ref. 28). The mouse monoclonal NuMA antibodies were identified in a monoclonal antibody screen for mitotic chromosome scaffold proteins (29). CENP-F antiserum was kindly provided by G. Chan (University of Alberta, Edmonton, Alberta, Canada; ref. 30). Secondary antibodies were from Molecular Probes (Eugene, OR). GFP-RHAMMFL and pEGFP-C1 (Invitrogen, Carlsbad, CA) plasmids were prepared as described previously (7).

Cell Culture, Transient Transfection, Immunofluorescence, and RNA Interference. RPMI 8226 (a myeloma cell line), Raji (a Burkitt's lymphoma cell line), and HeLa (a cervical adenocarcinoma line) were grown as recommended (American Type Culture Collection, Manassas, VA). Cells were passaged 24 hours before transfection. Suspension cells were transfected by electroporation (270 mV, 960 µF, 47-53 ms), whereas HeLa cells were transfected with LipofectAMINE 2000 (Invitrogen) following the manufacturer's protocols. At defined time points post-transfection, cells were fixed and permeabilized in cold methanol and washed with PBS-0.5% Triton X-100 (Sigma) before immunofluorescence. For double and triple staining experiments, antibodies were added sequentially. Cells were washed thrice in PBS-0.5% Tween before and after incubations. Cells were mounted in 90% glycerol/PBS + DAPI, and images were acquired using a Zeiss confocal LSM 510 or multiphoton microscope. Images were processed using MetaMorph Software (Universal Imaging Corp., Downington, PA) and Photoshop 5.02 software (Adobe Systems, Inc., Ottawa, Ontario, Canada).

We carried out RNA interference as described (31). Small interfering RNA targeting human TPX2 (NM_012112) and human RHAMM (NM_012484) were ordered predesigned using the Cenix Bioscience algorithm (Ambion, Austin, TX). TPX2 targeted sequence was 5'-GGAGAUACACAAAACAUAGtt-3' and RHAMM targeted sequence was 5'-GGUGCUUAUGAUGUUAAAAtt-3'; RHAMM sequence targets all RHAMMFL, RHAMM–exon 4, and RHAMM–exon 13 isoforms. Control RNA, targeting luciferase GL2, was ordered from Dharmacon, Inc. (Lafayette, CO). Oligonucleotides were annealed and transfected with LipofectAMINE 2000 as per manufacturer's suggested protocols. Briefly, 1.5 µL of 20 mmol/L small interfering RNA complex was incubated with 50 µL Opti-MEM (Invitrogen) and 1 µL LipofectAMINE 2000 was incubated with 50 µL Opti-MEM. These solutions were mixed and incubated at room temperature for 20 minutes before being diluted to 2 mL and added to cells. Cells were isolated for immunoblotting or immunofluorescence 24, 30, and 48 hours post-transfection.

Synchronization, Immunoprecipitation, and Quantitation. Raji and HeLa cells were synchronized by double thymidine (2 mmol/L, 14-16 hours) and nocodazole (300 ng/mL, 10-12 hours) block. Unsynchronized populations were released from plates with 1x trypsin. Both mitotic and unsynchronized populations were then washed thrice with PBS andlysed at 5 x 106 to 107 cells/mL in 1% CHAPS plus 10 µg/mL leupeptin, 10 µg/mL antipain, and 1 mmol/L phenylmethylsulfonyl fluoride (all from Sigma). All immunoprecipitation procedures were done at 4°C as described previously (7). For quantitation of coprecipitated proteins, post-immunoprecipitation lysates were collected and analyzed by SDS-PAGE. Protein quantitation used the Odyssey v1.1 IR imaging system (LI-COR) with detection of polyclonal sera using IRDye 800–conjugated anti-rabbit IgG (Rockland, Gilbertsville, PA).

Statistical Methods. Data were analyzed using SAS version 8.2 for Windows (SAS, Inc., Cary, NC). Correlation between centrosomal variables and continuous variables used Pearson's correlation coefficient. Two-group comparisons of centrosome variables used Student's t test. Statistical significance was set at P = 0.05 using two-sided analysis.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Centrosome Abnormalities Characterize Myeloma Cells. Myeloma plasma cells are characterized by numerous and complex chromosomal abnormalities (32). Given that centrosomal abnormalities are often present in chromosomally aberrant malignancies (17), we used multicolor immunofluorescence to examine MMPC from archived bone marrow core biopsies for centrosomal abnormalities. Bone marrow cores were examined from multiple myeloma patients (n = 41), MGUS patients (n = 8), and control patients (n = 4) plasma cells. Plasma cells from the morphologically normal bone marrow of lymphoma patients were used as controls. These plasma cells would be expected to have normal centrosomes. [If somehow cryptic lymphoma cells were mistaken for plasma cells, then this would be expected to lead to an overestimation of control plasma cell centrosome abnormalities (33). Such an error would only bias the results against the conclusions drawn here.] Experimenters were blinded to sample identity/diagnosis at the time of analysis and quantitation. Centrosomes in CD138+ plasma cells were visually assessed for qualitative structural and numerical abnormalities (see Table 1, column 5) as described previously (17). To quantitate structural abnormalities within CD138+ cells, plasma cell centrosomal volumes were determined by three-dimensional rendering of confocal z-stacks labeled with {gamma}-tubulin, a defined centrosomal structural protein (Fig. 1B).


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  		Table 1. Ex vivo analysis of centrosomes in MMPC

 


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  		Figure 1. Ex vivo analysis of the relationship between centrosomal abnormalities and RHAMM expression in multiple myeloma patient plasma cell. A, archived diagnostic multiple myeloma bone marrow cores were immunofluorescently labeled with CD138-FITC to indicate plasma cells and {gamma}-tubulin. CD138+ plasma cell were scored for structural (Cell i) and numerical (Cell ii) centrosomal abnormalities. White arrows, abnormal centrosomes. Bar, 10 µm. B, i, archived diagnostic multiple myeloma bone marrow cores (e.g., B97-62) were immunofluorescently labeled with CD138-FITC to indicate plasma cell and {gamma}-tubulin. Bar, 50 µm. Boxed area (red) in cell ii, confocal slices were collected through the core sections. Individual centrosomes are labeled alphabetically (white arrows). Bar, 10 µm. {gamma}-Tubulin channel was analyzed by Imaris software, and the centrosomes were three-dimensionally rendered. Volumes for represented centrosomes (white arrows) are (C) 0.849, (F) 0.177, (I) 0.926, (J) 1.171, (K) 0.587, and (L) 0.270 µm3 for sample B97-62. C, scatter plot graph for range in centrosomal volumes, centrosomes per cell, and percentage abnormal centrosome calculations for multiple myeloma (n = 42), MGUS (n = 8), and control (n = 4) plasma cell. Bars, mean. D, RHAMM expression within purified multiple myeloma CD138+ plasma cells, or bone marrow mononuclear cell with >45% infiltration, was determined by q-RT-PCR analysis. Relative RHAMM expression was normalized to the lowest patient expresser (patient 1). Columns, mean; bars, SD. Centrosomal volumes were determined within plasma cells from patients with RHAMM expression data. Points, centrosomal volumes; bars, SE. Centrosomal volumes were plotted against RHAMM expression, and a linear (black dash) and logarithmic trend line (red) was added. Linear correlation coefficient (r2) and the P associated with r are given for each relationship. E, purified multiple myeloma patient (n = 5) plasma cells were sorted and cytospun. Slides were methanol fixed, permeabilized, and stained with two centrosomal markers (RHAMM/{gamma}-tubulin or pericentrin/{gamma}-tubulin) or one centrosomal marker and a microtubule marker (RHAMM/{alpha}-tubulin or pericentrin/{alpha}-tubulin). Arrows, centrosomal region. Image intensities were measured within the whole cell and within the centrosomal region to determine centrosomal fractions (data not shown). Bar, 10 µm.

 
Plasma cell from all multiple myeloma samples analyzed showed numerical, structural, and volumetric abnormalities (Fig. 1A). Centrosome abnormalities, including the mean number of centrosomes per cell and the mean total centrosome volume, were highly correlated with one another in multiple myeloma (Pearson's r2 coefficient = 0.27; P < 0.001). On average, multiple myeloma plasma cells showed a significantly greater number of centrosomes per cell than MGUS (P < 0.05) and control plasma cells (P < 0.04). Quantitative volumetric assessment of structural abnormalities revealed a highly significant difference between multiple myeloma and MGUS (P < 0.002) and control plasma cells (P < 0.004). MGUS plasma cells had centrosomal variables intermediate to those of control plasma cells and multiple myeloma plasma cells (Table 1); indeed, MGUS plasma cells had larger centrosomal volumes when compared with those within control plasma cells.

RHAMM is, among its other functions, a centrosome protein (7). Given the presence of centrosome abnormalities in myeloma and the clinical significance of elevated RHAMM expression in this disease (10), we investigated whether elevated expression of RHAMM in multiple myeloma correlates with centrosome abnormalities.

In Myeloma, RHAMM Expression Correlates with Volumetric Centrosome Abnormalities. We used q-RT-PCR to investigate RHAMM expression within multiple myeloma patients (n = 20) with previously investigated centrosome variables. RNA was obtained from multiple myeloma patient CD138+ plasma cell (n = 5), or bone marrow mononuclear cell with ≥40% plasma cell infiltration (n = 15) when purified plasma cells were unavailable, for quantitative analysis. Within this multiple myeloma cohort, RHAMM expression was normalized to the level of the lowest RHAMM expresser (Fig. 1D). As with centrosomal abnormalities, multiple myeloma patients varied considerably in the absolute amount of RHAMM expression (Fig. 1D).

Centrosome volumes were measured in CD138-FITC-labeled multiple myeloma plasma cells from multiple myeloma patients for which the RHAMM q-RT-PCR expression level was known (n = 20). Regression analysis revealed a significant linear (r2 = 0.367; P < 0.005) and logarithmic relationship (r2 = 0.411; P < 0.003) between RHAMM expression and centrosome volumes (Fig. 1D). Centrosome per cell quantitation, however, did not show a significant linear relationship with RHAMM expression (r2 = 0.189; P = 0.104; data not shown).

RHAMM Localizes to Multiple Myeloma Centrosomes to the Same Extent as Other Centrosome Proteins. Next, we investigated the fraction of intracellular RHAMM, which localizes to the centrosome within MMPC. After CD38 or CD138 fluorescence-activated cell sorting separation, multiple myeloma patient (n = 6) plasma cells were stained with either two centrosomal markers (i.e., pericentrin, RHAMM, or {gamma}-tubulin) or one centrosomal marker and {alpha}-tubulin to show microtubule nucleation. Image analysis of confocal slices containing the centrosomal region revealed similar centrosomal fractions for pericentrin, RHAMM, and {gamma}-tubulin (0.10 ± 0.04, 0.08 ± 0.04, and 0.06 ± 0.03, respectively). Thus, although the intracellular fraction of RHAMM that localizes to interphase myeloma centrosomes is low, it is comparable with that of other defined components of the centrosome. As expected, intracellular RHAMM, but not pericentrin or {gamma}-tubulin, also colocalized with microtubules (Fig. 1E).

RHAMM Overexpression In vitro Affects Centrosome Size and Structure. To investigate a potential cause-effect relationship, we next examined whether overexpression of RHAMM in vitro could amplify centrosome volumes in a myeloma cell line. RPMI 8226 cells were transiently transfected with EGFP-C1 empty vector or GFP-RHAMMFL, and centrosomal structure was examined 16 to 20 hours later. Previous analysis indicates that transient transfection results in ~5-fold overexpression of exogenous RHAMM (7). Centrosomal volumes were determined with indirect immunofluorescence targeting {gamma}-tubulin or GFP-RHAMMFL fluorescence (Fig. 2A). {gamma}-Tubulin fluorescence within (a) cells transfected with EGFP-C1 and (b) cells that failed to express GFP-RHAMMFL (internal control) was used as a control for pericentriolar material volume determination (Fig. 2A). The mean centrosomal volumes for negative control samples were 0.541 ± 0.081 and 0.532 ± 0.123 µm3, respectively (Table 2). Overexpression of GFP-RHAMMFL resulted in mean pericentriolar material volumes of 1.039 ± 0.262 µm3 and a trend toward larger {gamma}-tubulin volumes (0.682 ± 0.079 µm3; Table 2). Thus, RHAMM overexpression adds a significant amount of excess pericentriolar material at the centrosome.



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  		Figure 2. RHAMM overexpression affects centrosomal structure and mitotic integrity. A, RPMI 8226 cells were transiently transfected with GFP-RHAMMFL, or EGFP-C1, and stained with {gamma}-tubulin. Confocal z-stacks were collected through individual cells and regions of interest, containing the {gamma}-tubulin signal, were exported to Imaris for three-dimensional rendering. Bar, 10 µm. Inset boxes, 2.5 x 2.5 µm extracted regions around the centrosome. A median (of 2 pixels) filter was applied to extracted regions. Effective zoom is 4x original image. Bar, 1 µm. B, HeLa cells were transiently transfected with GFP-RHAMMFL, or EGFP-C1, and stained with {alpha}-tubulin (data not shown). DNA was visualized with DAPI. Both transfected and nontransfected cells were scored within GFP-RHAMMFL transfected populations. Cells were examined for normal DNA alignment and segregation as well as for symmetrical, organized bipolar spindles (GFP-RHAMMFL and {alpha}-tubulin signals). Individual metaphase cells were scored as normal, disorganized spindles, unaligned chromosomes, monopolar, tripolar, tetrapolar, or multipolar spindles. Cells scored as unaligned chromosomes had phenotypically normal spindles. Bar, 10 µm.

 

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  		Table 2. Dysregulation of centrosomal volumes and mitosis with GFP-RHAMMFL overexpression

 
RHAMM Overexpression Compromises Mitotic Integrity. Structural centrosomal abnormalities, such as those that correlate with RHAMM expression in myeloma (Fig. 1D), have been strongly related to abnormal mitoses in human cancer (16). Thus, we analyzed the effect of RHAMM overexpression on mitotic integrity in vitro.

RHAMM mitotic localization has been documented in multiple different human cell models (34, 4, 7) and the mitotic function of RHAMM is conserved in Xenopus (35). Given the conservation of mitotic localization and function, transiently transfected HeLa cells were examined as they represent a standard in vitro system within which mitoses, and aberrant mitoses, can be analyzed and compared. EGFP-C1 transfected cells and neighboring, nontransfected cells (internal control) were used as negative controls for spindle abnormalities induced by GFP-RHAMMFL overexpression.

Similar to observations in RPMI 8226 (7), GFP-RHAMMFL transfection results in a metaphase block with dramatically fewer transfected cells observed in prophase (3.92% versus 12.1% and 9.8%) and telophase/cytokinesis (13.1% versus 33.5% and 29.1%) compared with EGFP-C1 and nontransfected controls (Table 2). Despite the high transfection efficiency (60-70%), fewer GFP-RHAMMFL transfected cells were mitotic (n = 153) than nontransfected internal control cells (n = 306), suggesting a significant decrease in viable mitotic cells dependent on GFP-RHAMMFL overexpression (Table 2).

To further investigate the effects of RHAMM overexpression on the dysregulation of mitosis, the number of abnormal mitotic cells, at various stages, was quantitated. DNA condensation, alignment, and segregation as well as the structure and number of spindle poles were analyzed to differentiate between normal and abnormal mitotic cells. The criteria for abnormal metaphase and anaphase was similar to that reported for Aurora A–inhibited S2 cells (36). GFP-RHAMMFL overexpression induced multiple abnormalities at various stages, with the most frequent being disorganized/long metaphase spindle fibers (18.3%; data not shown) and monopolar/tripolar/tetrapolar/multipolar metaphase spindles (16.3%; Fig. 2B); the frequency of disorganized spindle phenotypes (0% and 0.5%; data not shown) and mono/multipolar spindles (2.9% and 3.3%) was significantly reduced within nontransfected and EGFP-C1 transfected control populations, respectively (Table 2). Few (n = 28, 18.3%) mitotic, GFP-RHAMMFL overexpressing cells were observed post-metaphase (Table 2). Interestingly, 7 of 28 (25%) transfected cells showed lagging chromosomes, multipolar segregation, or chromosome bridges (Table 2). These post-metaphase phenotypes were present in 3 of 114 (2.6%, 1.0% of total) and 4 of 73 (5.5%, 1.1% of total) of nontransfected and EGFP-C1 transfected controls.

Given the evidence from clinical myeloma samples and in vitro transfection studies that link RHAMM to centrosome and mitotic errors, we sought to elucidate some of the functional aspects of RHAMM at the mitotic spindle that might be operative in producing these errors. The centrosome targeting domain of RHAMM bears significant homology to the BZIP motif of Xklp2, which interacts with dynein and TPX2 to target minus-end microtubules (7, 8). We therefore focused our efforts on the interaction of RHAMM with dynein and TPX2.

RHAMM Functionally Interacts with Dynein in Mitosis. Previous investigation of RHAMM function revealed that a subset of RHAMM colocalized with and coprecipitated dynein and vice versa (7). To extend on these findings, we investigated the dynamics of GFP-RHAMMFL in transiently transfected live HeLa and RPMI 8226 cells at room temperature (22-25°C). In addition to the mitotic spindle association, we observed aggregates of GFP-RHAMMFL that would load to the spindle pole with vectoral movement (Fig. 3A). On average, GFP-RHAMMFL cells contained 13.25 aggregates, with 17% demonstrating movement during the time course. We observed poleward movement of RHAMMFL-GFP aggregates with velocities [2.34 ± 0.39 (HeLa, n = 7) and 1.48 ± 0.28 (8226, n = 5) µm/min; Fig. 3A] similar to those observed for minus-end directed movement of NuMA-GFP by cytoplasmic dynein (1.0 ± 0.3 to 2.6 ± 1.0 m/min; ref. 37). Although singular, larger aggregates were observed within EGFP transfected cells, these aggregates did not transit during the experiment (t = 15 minutes). These observations, in combination with our previous coprecipitation data (7), show that RHAMM associates with the dynein motor complex to localize to the spindle pole within live cells.



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  		Figure 3. RHAMM dynamically redistributes during mitosis and colocalizes with and coimmunoprecipitate TPX2 in a cell cycle–dependent manner. A, HeLa or RPMI 8226 (data not shown) cells were transiently transfected with EGFPC1-RHAMMFL. Live RPMI 8226 were plated on coverslips coated with 100 µg/mL hyaluronan to facilitate imaging. Cells recovered for 8 to 12 hours post-transfection at 37°C, 5% CO2 and were visualized at room temperature and 5% CO2. Forty images were collected at 15-second intervals (time = 10 minutes), and velocities were analyzed on MetaMorph software. Arrow (time = 0 second), a GFP-RHAMM aggregate that is relatively positionally stable. Line (time = 0 second), MetaMorph-determined, minus-end directed path of GFP-RHAMM aggregate. Mean velocities of minus-end movement was 2.34 ± 0.39 µm/min (HeLa) or 1.48 ± 0.28 µm/min (RPMI 8226; data not shown). B, RHAMM and TPX2 colocalize throughout mitosis. HeLa were transiently transfected with GFP-RHAMMFL as described. RHAMM localization was indicated by GFP-RHAMMFL; TPX2 and {alpha}-tubulin localization was determined by indirect immunofluorescence with polyclonal TPX2 antiserum and a monoclonal anti-{alpha}-tubulin antibody followed by goat anti-mouse Alexa 546 and goat anti-rabbit Alexa 633. DAPI staining indicates DNA. White arrows in Prophase, extra centrosomal aggregates. Bottom Prometaphase, RHAMM and TPX2 colocalization within abnormal (multipolar) mitotic cells. Bar, 10 µm. C, HeLa lysates were prepared as described in Materials and Methods. Immunoblot analysis of RHAMM and TPX2 levels indicates equivalent amplification of RHAMM and TPX2 in mitotic (M) compared with unsynchronized (US) lysates. Immunoprecipitation experiments were separately done on 150 µL precleared lysates using a polyclonal RHAMM antibody, the prebleed (PB) immune sera, a polyclonal TPX2 antibody, and a monoclonal NuMA antibody as described. One fifth of the immunoprecipitation volume (25 µL) of precleared lysate (Loading Control) and post-immunoprecipitation fractions were analyzed to determine relative quantity and efficiency of the precipitations. Resulting immunoprecipitates were blotted with a RHAMM polyclonal serum or a TPX2 polyclonal serum. Blots showing RHAMM and TPX2 detection of prebleed and RHAMM precipitation lanes were done in repetitive experiments. Detection of protein bands used enhanced chemiluminescence detection. D, post-immunoprecipitation fractions for the mitotic lysates indicate the relative amounts of RHAMM or TPX2 coprecipitated by various antisera. Quantitative detection used the Odyssey v1.1 IR imaging system with detection of polyclonal sera using IRDye 800–conjugated anti-rabbit IgG (1:20,000) to determine protein band concentrations. Quantitation of protein bands with the Odyssey systems subtracts background fluorescence intensity around the region of interest (in this case, the boxed regions outlining the protein bands). Top row, bands without concentration determination; bottom row, raw concentration measurements. Concentrations of the prebleed and NuMA bands were averaged to give a mean total fraction. Thus, the fraction remaining was determined by dividing the band concentration of each lane by the mean total fraction. E, HeLa and Raji (data not shown) cells were synchronized by double thymidine block followed by nocodazole treatment. Mitotic HeLa cells were shaken off the plate and examined for DNA, message, and protein content. Flow cytometry profile of DAPI-stained HeLa populations confirms >90% synchronization. M1, G0-G1 populations; M2, G2-M populations; M3, S populations. F, unsynchronized and mitotic HeLa and Raji (data not shown) populations were lysed in CHAPS, and the soluble fraction was analyzed by quantitative immunoblot analysis. Quantitative detection used the Odyssey v1.1 IR imaging system with detection of polyclonal sera using IRDye 800–conjugated anti-rabbit IgG. Note that detection of RHAMM antiserum used 1:5,000 dilution, whereas TPX2 detection used 1:20,000. ß-actin intensities were determined and used as a quantitative control for lane loading; M-phase lysates consistently contained more protein per cell volume lysed. G, unsynchronized and mitotic HeLa and Raji (data not shown) populations were reverse transcribed as per manufacturer's (Applied Biosystems, Foster City, CA) recommendation for q-RT-PCR analysis. RHAMM expression in mitotic populations were normalized to the unsynchronized populations.

 
RHAMM Colocalizes with and Coimmunoprecipitates TPX2 and Vice Versa in a Cell Cycle–Dependent Manner. We next investigated whether RHAMM colocalizes with TPX2 during mitosis. During interphase, GFP-RHAMMFL colocalizes with microtubules and centrosomes, whereas TPX2 is nuclear (Fig. 3B). At prophase, however, both RHAMM and TPX2 redistribute to the separating centrosomes (Fig. 3B). At prometaphase and throughout metaphase, both proteins localize to the poles, and along the arms, of phenotypically normal as well as multipolar, mitotic spindles (Fig. 3B). Both proteins localize to the spindle midzone during anaphase and concentrate at the midzone during telophase (data not shown).

To further investigate a putative RHAMM-TPX2 association, HeLa cells were synchronized through double thymidine/nocodazole block to provide mitotic extracts. Mitotic and unsynchronized extracts were separately immunoprecipitated with prebleed serum and sera targeting RHAMM, TPX2, and NuMA. NuMA/dynein complexes play an integral role in spindle assembly and focusing (37) and thus NuMA served as an excellent control for the specificity of a putative RHAMM-TPX2 interaction. Lysis of equivalent numbers of cells from mitotic and unsynchronized populations resulted in amplification of both RHAMM and TPX2 levels in the mitotic extracts (Fig. 3C). These results are consistent with previous reports demonstrating cell cycle regulation of TPX2 (26) and RHAMM (ref. 38; see Fig. 3F and G). The high efficiency of immunoprecipitation was confirmed by examination of post-immunoprecipitation lysates, with 70% to 100% of mitotic TPX2 and RHAMM being precipitated (Fig. 3C). During mitosis, RHAMM antibodies coprecipitated a significant amount of TPX2 and vice versa. This reciprocal coprecipitation was not obtained within unsynchronized lysates. To quantitate the level of coprecipitation, IR detection of protein was used to determine the levels of RHAMM and TPX2 remaining in the post-immunoprecipitation lysates (Fig. 3D). Unlike protein detection based on enzymatic amplification (enhanced chemiluminescence, Amersham Biosciences, Sunnyvale, CA), IR labeling results in linear detection of protein and, as a consequence, results in differential detection of RHAMM and TPX2 bands. Thus, the level of mitotic RHAMM seems greater with enhanced chemiluminescence detection. Neither prebleed nor NuMA precipitation resulted in a significant loss of RHAMM or TPX2 protein levels in the post-immunoprecipitation fractions. The concentrations of the bands resulting from prebleed and NuMA precipitations were averaged to give an expected post-precipitation fraction. The fraction remaining was determined by dividing the band concentration of each lane by the expected post-precipitation fraction. Immunoprecipitation with RHAMM antibodies resulted in a depletion of 36% and 100% of the cellular TPX2 and RHAMM in mitotic lysates, respectively. Within mitotic lysates, TPX2 precipitation resulted in a 70% and 56% loss of TPX2 and RHAMM, respectively (Fig. 3D). These results show that a significant population of RHAMM is associated with TPX2 within HeLa mitotic cells and vice versa.

RHAMM, Like TPX2, Is Regulated during the Cell Cycle. As mentioned, RHAMM and TPX2 are subject to cell cycle regulation (26, 38) although the level of RHAMM amplification during mitosis has not been reported. HeLa, and Raji (data not shown), lines were synchronized during G2-M with a double thymidine/nocodazole block. The efficiency of these treatments was confirmed by flow cytometry analysis of DAPI staining within the synchronized populations (Fig. 3E). Cell lysates were prepared through equivalent CHAPS lysis of HeLa (5 x 106/mL) and Raji (20 x 106/mL) synchronized populations. Quantitative immunoblot analysis was done by normalizing the RHAMM or TPX2 level to the corresponding ß-actin levels. RHAMM protein levels, like TPX2, are cell cycle regulated (Fig. 3F) with 1.5 to 2.0 times greater expression in G2-M versus unsynchronized Raji and HeLa lysates. q-RT-PCR revealed a 1.43 to 1.75 times amplification of RHAMM message in G2-M Raji and HeLa populations (Fig. 3G).

Inhibition of RHAMM or TPX2 Function Results in Similarly Aberrant Mitoses. As significant amounts of RHAMM and TPX2 interact during mitosis, we hypothesized that RNA inhibition of RHAMM may induce similar phenotypes to that of TPX2 inhibition. RNA inhibition resulted in sustained loss of >95% of cellular RHAMM and TPX2 protein 24 to 48 hours post-transfection (Fig. 4A). Cells were lysed and examined by immunoblot 24 and 48 hours post-transfection and compared with control-treated cells (GL2). Treatments were examined in triplicate. Inhibited and control cells were then fixed at defined time points and examined for mitotic index using the marker CENP-F and {alpha}-tubulin. CENP-F expression, and localization, varies during the cell cycle, demonstrating increasing nuclear staining during G2 and accumulation on kinetochores early in G2-M transition (39). Mitotic index was scored for an average of 493 cells per treatment at 30 and 48 hours post-transfection.



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  		Figure 4. RNA inhibition of RHAMM and TPX2 reveal a common role in the maintenance of mitotic integrity. We carried out RNA interference as described (31). A, cells were treated with test or control duplexed RNA/LipofectAMINE 2000 mixes overnight in Opti-MEM (Life Technologies). At 24 and 48 hours, CHAPS soluble fractions were analyzed by immunoblot. Immunoblotting confirmed >95% inhibition of RHAMM and TPX2 24 and 48 hours post-transfection. Blots were detected with enhanced chemiluminescence (shown) and LI-COR detection. Equivalent loading was confirmed with ß-actin. B, coverslips were recovered at appropriate time points following transfection. Cells were stained with CENP-F, {alpha}-tubulin, and DAPI to quantitate mitotic index. At the 30-hour time point, inhibited cells were scored for stage, mitotic spindle structure, and chromosome condensation/alignment. Bars, SD of mitotic index calculations (~500 cells per treatment scored). For analysis of mitotic defects, RNA inhibition was confirmed in individual cells by staining for the appropriate target protein. Individual inhibited cells were assessed for DNA alignment and spindle structure to determine mitotic stage. TPX2 inhibition resulted in a significant increase in mitotic index (P = 0.017), whereas RHAMM inhibition resulted in a significant increase in G2 cells (P = 0.007) when compared by a Student's t test with control inhibited cells at the 30-hour time point. At the 48-hour time point, both RHAMM and TPX2 inhibition resulted in significant increases in metaphase cells (P < 0.05) and multipolar architecture (P < 0.03). All graphical data are percentages of all cells examined. C, control RNA inhibition did not affect normal localization or intensity of RHAMM and TPX2 staining at all mitotic stages. Bar, 10 µm. D, at the 30-hour time point, RHAMM inhibition induced phenotypically normal spindles with a proportion of metaphase cells (28.2%) exhibiting chromosome missegregation defects as shown by DAPI (white to facilitate viewing). TPX2 intensity and localization was not affected by RHAMM inhibition. At the 30-hour time point, TPX2 inhibition induced an increase in mitotic cells with prometaphase-like phenotypes (separated microtubule asters, condensed chromosomes but little microtubule connections between the two asters). RHAMM localization to microtubule asters was reduced within TPX2-inhibited cells. Nuclear CENP-F intensity and localization initiates and increases throughout G2, loading onto pre-kinetochore at G2-M transition. Control inhibition resulted in a G2 index of ~20.5%, whereas RHAMM inhibition resulted in a significantly increased G2 index (41.6%, P < 0.01). Bar, 10 µm. E, at the 48-hour time point, RHAMM inhibition resulted in a significant increase in metaphase cells. A large proportion of these metaphase cells (27.1%) contained mutlipolar spindle architecture. TPX2 localization to multipolar spindles was not affected by RHAMM inhibition. At the 48-hour time point, TPX2 inhibition also resulted in a significant increase in metaphase cells. Like RHAMM inhibition, these metaphase cells frequently (27.0%) displayed a multipolar phenotype. RHAMM localization to poles was dependent on TPX2. Bar, 10 µm.

 
At 30 hours, inhibition of TPX2 resulted in an increased mitotic index (7.0% versus 3.3%; P < 0.02) and the suppression of spindle formation (Fig. 4C). As described by Gruss et al. (26), prometaphase-like phenotypes, characterized by chromosome condensation, aster separation, and decreased microtubule polymerization, were observed in 82.1% of TPX2-inhibited mitotic cells (Fig. 4D). Within these cells, RHAMM fluorescence was not significantly localized to the separated microtubule asters, suggesting a role for TPX2 in the spindle pole targeting of RHAMM (Fig. 4D). Very few mitotic cells (2.4%) were observed post-metaphase, which strongly supports the hypothesis that TPX2 is important in the initiation of spindle assembly (Fig. 4D). However, as TPX2-inhibited cells clearly proceed through mitosis, albeit at later time points and with significantly aberrant spindles (see Fig. 4E), we conclude that it is not essential for mitotic assembly. We confirm previous work (27) and report that at later time points TPX2 inhibition induces a mitotic block with ~30.9% of TPX2-inhibited metaphase cells containing multipolar spindle phenotypes. Within this population of cells, we detected reduced amounts of TPX2 at the spindle poles. In the presence of reduced TPX2, RHAMM localized to the poles (Fig. 4E). However, a proportion of TPX2-inhibited cells lacked polymerized spindles similar to that described above; within this population, RHAMM was not localized to microtubule asters.

At 30 hours, RHAMM inhibition resulted in a significant increase in G2 cells (41.6% versus 20.5%; P < 0.01) characterized by CENP-F-positive nuclei (Fig. 4B and D). Although bipolar spindles formed by the 30-hour time point, approximately one third of the metaphase cells had chromosomal missegregation defects (see Fig. 4D). The increased G2 population suggests that RHAMM inhibition results in a G2-M stall before the initiation of prophase. At the 48-hour time point, RHAMM inhibition resulted in an accumulation of mitotic cells in metaphase (see Fig. 4B and E). A large proportion of these metaphase cells (36.8%) contained multipolar architecture, confirming our previous microinjection observations (7). RHAMM inhibition also resulted in a population of mitotic cells, designated abnormal metaphase, with small spindles or no spindles (similar to that described for TPX2 inhibition). Within the RHAMM-inhibited multipolar populations, TPX2 localized to the mitotic spindle poles (Fig. 4E). At 48 hours, both RHAMM and TPX2 inhibition resulted in dramatic increases in apoptotic cells and few G1-S cells, demarcated by CENP-F-negative nuclei (data not shown). These results support a role for RHAMM and TPX2 in the maintenance of mitotic integrity, consistent with the conclusions drawn previously for TPX2 (27). Temporal examination of RHAMM and TPX2 inhibition reveal that these proteins perform distinct roles early in mitotic assembly; inhibition of RHAMM delays mitosis at the G2-M boundary, whereas TPX2 inhibition delays spindle assembly after nuclear envelope breakdown. These differences in early mitotic function are to be expected between proteins that localize to the centrosome and nucleus during interphase. However, the functions of these proteins converge during metaphase. In the absence of RHAMM (or TPX2), mitotic spindles stall and fragment during metaphase; this phenomenon implicates both RHAMM and TPX2 as structural components of the spindle that cross-link microtubules and maintain spindle integrity as various microtubule-dependent motors exert force on spindle microtubules.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Elevated expression of centrosomal proteins, including pericentrin and aurA, have been associated with centrosomal abnormalities in vivo, and exogenous overexpression of these gene products can induce centrosomal and chromosomal abnormalities (17, 40). Here, we investigate the relationship among RHAMM, centrosomal abnormalities, and mitotic stability. Elevated RHAMM expression characterizes the malignant multiple myeloma cell (9) and correlates with poor prognosis, cytogenetic abnormalities (10), and centrosomal abnormalities within multiple myeloma plasma cell ex vivo. Elevated RHAMM significantly correlates with centrosomal structural abnormalities as quantitatively determined by centrosomal volume measurements but not numerical abnormalities. These data are consistent with RHAMM contributing to pericentriolar material structure but not affecting centriolar replication. Cells with pericentriolar excess are strongly associated with abnormal mitoses (16). Accordingly, exogenous GFP-RHAMMFL localizes to centrosomes, increasing their size and the amount of {gamma} -tubulin present, and induces aberrant mitotic division. Live cell imaging confirms previous coprecipitation data and suggests that RHAMM targets spindle poles through an interaction with dynein.

RHAMM coprecipitates a significant amount of cellular TPX2 in a cell cycle–dependent manner. RNA inhibition experiments show that at early time points TPX2 plays an important, but nonessential, role in spindle assembly, whereas RHAMM functions in the progression through G2. Interestingly, addition of a soluble RHAMM variant, lacking the NH2-terminal microtubule binding domain, also dysregulates G2-M progression (41), whereas overexpression of a RHAMM COOH deletion variant inhibits mitotic progression (7). After spindle assembly, RHAMM and TPX2 converge to play essential roles in maintaining spindle integrity. These data are consistent with a model in which a subset of intracellular RHAMM contributes to centrosomal structure during interphase and functions in early G2-M progression events. During spindle assembly, RHAMM interacts with a large fraction of the cellular pool of TPX2, which may facilitate an interaction between RHAMM and the dynein motor complex. This ternary RHAMM-TPX2-dynein complex participates in the maintenance of spindle integrity. Depletion of either TPX2 or RHAMM results in an imbalance of motor forces and subsequent spindle fragmentation. Conversely, overexpression of RHAMM results in an opposite imbalance of force cumulating in disorganized or multipolar spindles and an inability to appropriately align and segregate DNA. Similarly, TPX2 overexpression in Xenopus extracts results in unbalanced monopolar spindles (42). Although multipolar phenotypes are likely catastrophic, disorganized or fragmented spindles may induce more subtle losses of genetic material (see Fig. 4D), or impede mitotic progression, and potentially result in aneuploid progeny. The progression of such defects through mitosis is likely dependent on simultaneous disruption of the p53-dependent cell cycle checkpoint (43). A direct link among RHAMM overexpression, oncogenesis, and apoptosis may be mediated through TPX2 and aurA.

Like RHAMM, aurA localizes to interphase centrosomes, duplicating prophase centrosomes, and the mitotic spindle pole (7, 44); aurA localization and activation at spindle poles is also dependent on the action of TPX2 (28). Here, we describe a cell cycle–dependent interaction between significant mitotic fractions of RHAMM and TPX2, suggesting that RHAMM may affect TPX2-mediated aurA activation. AurA is an essential determinant of G2-M progression that regulates, and is regulated by, the activities of p53 (44), ras-GTPase-activating protein (45), BRCA1 (46), and apoptotic pathways (47). Overexpression of RHAMM, through an interaction/sequestration of TPX2, may disrupt these pathways and induce/permit aberrant, or abortive, mitoses with consequent centrosome amplification and chromosomal instability. Alternatively, overexpression of RHAMM, or splice variants (i.e., RHAMM–exon4), may augment spindle forces leading to spindle fragmentation and chromosomal segregation defects.

Depending on reagents, methodologies, cell models, and culture conditions, intracellular RHAMM has multiple documented subcellular localizations (i.e., interactions with cytoskeletal components, centrosomes, mitochondria, nuclei, and podosomes) and functions, including participation in hyaluronan-mediated motility, signaling events, and cytoskeletal interactions (34, 4, 7). However, multiple groups, with multiple different reagents, have documented RHAMM at the mitotic spindle within mammalian and Xenopus systems (4, 34, 7, 35). These observations, along with the elevated expression of RHAMM in proliferative tissues like the testis (48) and its cell cycle–dependent regulation, imply a conserved role for this protein during cell division. Interestingly, evidence is mounting for the accumulation of intracellular hyaluronan during mitosis (49), and RHAMM-hyaluronan interactions participate in the accumulation of hyaluronan within the nucleus during motility (50). Thus, the possibility exists for intercompartmental cross-talk involving RHAMM-hyaluronan interactions outside and within the cell during division. However, the relevance of intracellular hyaluronan accumulation, and its effect(s) on RHAMM or other B(X)7B proteins, is currently undefined.


   Acknowledgments
 
Grant support: Canadian Institutes of Health Research (L.M. Pilarski, T. Reiman, and A.R. Belch); National Cancer Institute grant CA80963 (L.M. Pilarski and A.R. Belch); Canada Research in Biomedical Nanotechnology Chairs Program (L.M. Pilarski); and Natural Sciences and Engineering Research Council of Canada studentship, Alberta Heritage Foundation for Medical Research studentship, and Department of Oncology Ph.D. endowed studentship (C.A. Maxwell).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

We thank Laith Dabbagh and Sam Johnson for their expert assistance.

Received 4/22/04. Revised 10/12/04. Accepted 11/22/04.


   References
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
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