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Originally published In Press as doi:10.1074/jbc.M310934200 on January 12, 2004

J. Biol. Chem., Vol. 279, Issue 14, 14039-14048, April 2, 2004
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N Terminus Is Essential for Tropomyosin Functions

N-TERMINAL MODIFICATION DISRUPTS STRESS FIBER ORGANIZATION AND ABOLISHES ANTI-ONCOGENIC EFFECTS OF TROPOMYOSIN-1*

Shantaram Bharadwaj{ddagger}, Sarah Hitchcock-DeGregori§, Andrew Thorburn¶, and G. L. Prasad, Recipient of Career Development Award DAMD-98-1-8162) from the United States Army Breast Cancer Research Program{ddagger}||

From the Departments of {ddagger}General Surgery and Cancer Biology, Wake Forest University School of Medicine, Winston-Salem, North Carolina 27157 and the §Department of Neuroscience and Cell Biology, Robert Wood Johnson Medical School, Piscataway, New Jersey 08854

Received for publication, October 3, 2003 , and in revised form, January 3, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Down-regulation of several key actin-binding proteins, such as {alpha}-actinin, vinculin, gelsolin, and tropomyosins (TMs), is considered to contribute to the disorganized cytoskeleton present in many neoplastic cells. TMs stabilize actin filaments against the gel severing actions of proteins such as cofilin. Among multiple TMs expressed in non-muscle cells, tropomyosin-1 (TM1) isoform induces stress fibers and functions as a suppressor of malignant transformation. However, the molecular mechanisms of TM1-mediated cytoskeletal effects and tumor suppression remain poorly understood. We have hypothesized that the ability of TM1 to stabilize microfilaments is crucial for tumor suppression. In this study, by employing a variant TM1, which contains an N-terminal hemagglutinin epitope tag, we demonstrate that the N terminus is a key determinant of tropomyosin-1 function. Unlike the wild type TM1, the modified protein fails to restore stress fibers and inhibit anchorage-independent growth in transformed cells. Furthermore, the N-terminal modification of TM1 disorganizes the cytoskeleton and delays cytokinesis in normal cells, abolishes binding to F-actin, and disrupts the dimeric associations in vivo. The functionally defective TM1 allows the association of cofilin to stress fibers and disorganizes the microfilaments, whereas wild type TM1 appears to restrict the binding of cofilin to stress fibers. TM1-induced cytoskeletal reorganization appears to be mediated through preventing cofilin interaction with microfilaments. Our studies provide in vivo functional evidence that the N terminus is a critical determinant of TM1 functions, which in turn determines the organization of stress fibers.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
One of the most common and yet prominent features of neoplastic cells is the presence of disorganized actin microfilaments (1, 2). A functionally defective cytoskeleton, arising from the disorganized microfilament architecture, has been shown to be responsible for the loss of normal cellular morphology and cell polarity; altered intracellular transport, cell motility, and cell adhesion; and defective cytokinesis. Suppression of several key actin-binding proteins, including tropomyosins (TMs),1 occurs in many neoplastic cells, and this contributes to the assembly of disorganized cytoskeleton (reviewed in Refs. 3 and 4). Down-regulation of TMs in malignantly transformed cells has been known for about 2 decades and is widely reported (5–7), and yet the role of TMs in neoplastic transformation of cells remains incompletely understood.

TMs are a family of actin-binding proteins that stabilize microfilaments from the gel severing actions of proteins such as gelsolin and cofilin (1, 8). Multiple TMs are generated by alternative splicing with a high degree of tissue specificity. For example, fibroblasts express five different, closely related TMs that may be categorized into high and low Mr species containing 284 and 248 amino acids, respectively. TMs are key regulatory proteins of actin cytoskeleton in that they regulate almost all aspects of actin polymerization (9–13). Although the function of TMs is better elucidated in skeletal and cardiac muscles, given their diverse and tissue-specific expression patterns, the importance of the existence of multiple TMs in nonmuscle cell physiology is poorly understood (8). It has been suggested that TM isoforms perform distinct functions rather than being simply redundant (1, 8). Work from this and several other laboratories shows that the high Mr TMs are consistently down-regulated in many malignantly transformed cells (5–7, 14–16). This suggests a role for TMs in the maintenance of normal growth and cytoskeletal organization, and that expression of TM1 may be incompatible with neoplastic growth.

In support of this hypothesis, we have recently shown that TM isoform-1 (TM1) expression is widely and profoundly down-regulated in primary breast tumors (17). Furthermore, TM1 restores the microfilament organization in transformed cells, and suppresses malignant growth (16, 18–20). Tumor suppression by TM1 is isoform-specific; for example, unlike TM1, closely related TMs such as TM2 failed to suppress transformed growth of highly malignant v-ki-ras-transformed NIH3T3 (DT) cells or MCF-7 human breast carcinoma cells (17, 19). Although these studies suggested TM1 is a class II tumor suppressor (17, 20, 21), the molecular basis of cytoskeletal organization and tumor suppression by TM1 remains unknown.

TM1 lacks distinct catalytic activity or binding partners that readily explain the isoform-specific anti-oncogenic effects. Structurally, all TMs are predominantly {alpha}-helical proteins in which hydrophilic and hydrophobic amino acids occupy defined positions in a repeating heptapeptide (22). TM1, like other TMs, is a cytosolic structural protein that binds to actin stoichiometrically in micromolar range. For example, both TM1 and TM2 bind to actin at 1:7 ratio, although some differences in binding properties exist (23). Nevertheless, it is intriguing that the biological effects of TM1 expression in tumor cells are remarkably different from those of other TM isoforms. We have considered that reorganization of microfilaments is a critical component of tumor suppression by TM1. The studies presented here demonstrate that the N terminus of TM1 is essential for TM1 functions and imply that TM1 is a key modulator of stress fibers.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture and Antibodies—Culture conditions and media for NIH3T3, NIH3T3/TM1, and DT/TM1 cells have been described previously 2(4). Doubly transformed (DT) cells are NIH3T3 cells transformed with two copies of the v-ki-ras oncogene (7). An epitope-tagged TM1 was constructed by cloning TM1 in-frame in pCGN vector to add hemagglutinin (HA) epitope at the N terminus. The variant TM1 thus produced would have an N-terminal extension, ASSYPYDVPDYASLGGPSR, but contains identical wild type TM1 sequence from the "R" onward. DT cells were cotransfected with the recombinant plasmid and pCMVneo to generate single cell-derived clones by standard transfection methods. N-[1-(2,3-Dioleoyloxy)propyl]-N,N,N-trimethylammonium methyl sulfate salts (Roche Applied Science) or LipofectAMINE (Invitrogen) were used for transfections. In experiments involving transient transfections, cells were routinely processed 48 h after transfection, unless otherwise indicated.

A TM polyclonal antibody, generated in this laboratory, that detects multiple TMs was described previously (16). TM311 mouse monoclonal antibody that recognizes a common epitope found in all high Mr TMs (25) was obtained from Sigma. HA (12CA5) mouse monoclonal antibody (Roche Applied Science) and antibodies against {alpha}-tubulin (Sigma), phosphocofilin (Upstate Biotechnology, Inc.), actin, and cofilin (Cytoskeleton, Inc., Denver, CO) were purchased.

Monolayer Growth and Morphology—Ten thousand cells were plated and counted, and cell numbers were plotted against time of culture. Morphology of subconfluent monolayer cultures was recorded by fixing and staining with a HEMA 3 kit obtained from Fisher. The samples were photographed at x40 magnification.

Soft Agar Assays—Anchorage-independent experiments were performed in soft agar as described previously. One thousand cells were mixed in 0.36% agar, plated on a 0.8% agar base, and cultured for 10–14 days (18). Cells were stained overnight with 0.05% nitro blue tetrazolium in PBS, and colonies were counted.

Immunofluorescence—Cells were cultured in chamber slides (Nunc), fixed with 3.7% paraformaldehyde, and extracted with 0.5% Triton X-100 for 5 min (26). The samples were incubated with an appropriate primary antibody, followed by a second antibody conjugated to a fluorochrome, and finally with Texas Red-conjugated phalloidin. Slides were finally rinsed in water and mounted using Antifade kit (Molecular Probes). Images were recorded using a Zeiss LSM 510 confocal microscope and imported into Adobe Photoshop.

Immunoblotting and Immunoprecipitations—Cells were lysed in a buffer containing Nonidet P-40, sodium deoxycholate, and protease inhibitors and clarified at 14,000 x g (26). Supernatants containing 50–100 µg of proteins were separated on 13% SDS-polyacrylamide gels and immunoblotted. For immunoprecipitations, 200 µg of protein lysate, precleared with protein G for 1 h, was incubated with the primary antibody (27). The immune complexes were washed with immunoprecipitation buffer and subjected to SDS-PAGE. To quantify the protein expression, the exposed membranes were scanned, and the band intensities were calculated using the magic wand tool of the Adobe Photoshop (version 6.0).

Metabolic Labeling of Cells—Subconfluent cultures were pre-incubated with labeling medium (Dulbecco's modified Eagle's medium containing 10% dialyzed fetal bovine serum) for 3 h. Metabolic labeling was carried out in the labeling medium containing [35S]methionine (50 µCi/ml) for 6 h (24). Cells were washed with PBS and extracted in Nonidet P-40/deoxycholate lysis buffer. For immunoprecipitations and cross-linking experiments, lysates equivalent to 4 x 106 cpm were used.

F-actin Quantitation in Cells—F-actin content in cells was measured using a protocol described by Zigmond and co-workers (28) with minor modifications. Cells (1.5 x 105) were plated 12–24 h prior to experimentation. Monolayer cultures were washed with PBS, fixed, and stained in PBS containing 3.7% paraformaldehyde, 0.5% Triton X-100, and 0.2 µM TRITC phalloidin for 3 h at room temperature with constant rocking in the dark. Unincorporated TRITC phalloidin was removed by four PBS washes. Cell monolayer was extracted by using 1 ml of methanol, and the cell suspension was transferred to microcentrifuge tubes and incubated for 48 h with constant rocking at 4 °C. Cell debris was removed by centrifugation and fluorescence (540ex/575em) was read in an Aminco Bowman luminescence fluorimeter using the methanol solvent as blank. To determine background fluorescence, 2 µM of unlabeled phalloidin was added along with TRITC-phalloidin. To determine total actin content, cell lysates were probed with anti-actin antibody in immunoblots and expressed as a ratio with endogenous tubulin.

Cell Cycle Analysis—To estimate the fraction of cells in each phase of cell cycle, asynchronously growing subconfluent cultures were trypsinized and stained with 50 µg/ml propidium iodide in PBS containing 0.06% Nonidet P-40 and 30 µg/ml RNase A. Cell cycle analyses were carried out using a BD FACS Star Plus flow cytometer. Flow cytometry of serum-starved (24 h) or stimulated cells was performed. For serum stimulation, serum-starved cells were cultured in regular medium for 28 h.

HA-TM1 Purification—HA-TM1 was subcloned in pET3a (Novagen) and expressed in BL21 (DE3) pLysS bacteria. Cultures were grown to A600 0.4–0.6 and induced with 0.4 mM isopropyl-1-thio-{beta}-D-galactopyranoside for 3 h. Cells were harvested and resuspended for sonication in lysis buffer containing 20 mM Tris (pH 7.5), 100 mM NaCl, 2 mM EDTA, 1x protease inhibitor mixture (Roche Applied Science), 1 mg/liter DNase, 1 mg/liter RNase A, 1 mM 4-(2-aminoethyl)benzenesulfonyl fluoride, 5 mM EDTA, 5 mM EGTA, 5 mM PMSF, 5 mM benzamidine, and 1 mg of potato carboxypeptidase inhibitor (Calbiochem) and immediately subjected to ammonium sulfate precipitation to achieve ~50% saturation. The protein pellet was resuspended and extensively dialyzed against a buffer containing 20 mM Tris (pH 7.5), 1 mM PMSF, and 1mM benzamidine. The salt concentration of the dialysate was adjusted to 50 mM and subjected to DEAE-cellulose chromatography. HA-TM1 was eluted with 0.3 M NaCl containing buffer (20 mM Tris (pH 7.5), 5 mM EDTA and 2.5 mM PMSF), reprecipitated with ammonium sulfate (100% saturation to concentrate the eluate), and dialyzed. The protein sample was further purified by affinity chromatography on HA epitope affinity column (Roche Applied Science) as per the manufacturer's instructions. HA-TM1 was eluted with 1 mg/ml HA epitope peptide (Roche Applied Science) and dialyzed against the binding assay buffer (10 mM Tris (pH 7.5), 150 mM NaCl, 2 mM MgCl2, 0.2 mM EGTA).

In Vitro Actin Binding Studies—Actin binding assays were performed as described previously (29) with modifications (30). HA-TM1 and wild type TM2 were cosedimented at 20 °C with chicken pectoral muscle F-actin (5 µM) in a buffer containing 150 mM NaCl, 10 mM Tris-HCl (pH 7.5), 2 mM MgCl2, and 0.5 mM dithiothreitol. The amounts of bound and free tropomyosin in the supernatants and pellets were quantitated by densitometry of SDS-polyacrylamide gels stained in Coomassie Blue using a Molecular Dynamics model 300A computing densitometer (Amersham Biosciences). To separate HA-TM1 from actin, the gels also contained 6 M urea. The free tropomyosin in the supernatants was calculated from standard curves for wild type tropomyosin. The curve for TM2 was fit using the Hill equation using SigmaPlot (SPSS Science, Chicago) that reported a Kapp.

Cross-linking Studies—Homodimers of TM1 were stabilized by cross-linking with 5,5'-dithiobis-2-nitrobenzoic acid (24, 31) (DTNB) (Sigma), a sulfhydryl cross-linker, as described previously (24). Cell lysates were incubated with DTNB and were either subjected to immunoblotting (for unlabeled samples) or immunoprecipitation followed by SDS-PAGE and fluorography (for 35S-labeled samples). Dimers are detectable when 2-mercaptoethanol is omitted in the gel sample buffer.

Statistical Analyses—Data are presented as mean ± S.D. from at least three independent determinations. p values were calculated by Student's two-tailed t test (32) using the software provided in Microsoft Excel (2002 edition).


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
In vitro studies have indicated that the N- and C-terminal ends of TMs are critical for binding to actin. Several studies have shown that some TMs, including TM1, require the acetylated N terminus for optimal binding to actin. N-terminal extensions, depending on the length and sequence, can alter TM functions (33–35). To elucidate the mechanism of TM1-mediated tumor suppression, we have modified the N-terminal end of TM1 molecule by introducing a hemagglutinin (HA) epitope. This epitope tag contains three prolines and would not be predicted to be {alpha}-helical. This tagged protein, referred to as HA-TM1, contains a 19-residue N-terminal extension, which otherwise is identical to the wild type protein, and retains all the actin binding domains of TM1. We tested the ability of HA-TM1 to regulate cytoskeletal organization and growth phenotype.

N-terminal Modification Abolishes TM1-mediated Cytoskeletal Reorganization and Tumor Suppression—We have used DT (NIH3T3 cells transformed by v-Ki-ras) cells as a model and transfected them with HA-TM1. Stable single cell clones, designated as DT/HA-TM1, were isolated. DT cells express TM1 at 50% levels compared with normal NIH3T3 fibroblasts, and TM2 and TM3 at essentially undetectable levels (18). Expression of HA-TM1 was detected by Northern blot (not shown) and immunoblot methods (Fig. 1A). The variant protein, because of the N-terminal extension, migrates slower than wild type TM1 on SDS-PAGE. The expression of endogenous TM1 in parental DT and DT/HA-TM1 cells is determined by the ratio of TM1: {alpha}-tubulin. In DT cells, the relative expression of TM1 was 0.7 ± 0.12, and in DT/HA-TM1 cells, the endogenous TM1 was expressed at 0.75 ± 0.08, indicating that the transfected HA-TM1 did not alter the levels of the endogenous wild type protein (p < 1).



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  		FIG. 1.
Effect of expression of HA-TM1 on DT cell morphology and microfilament organization. A, TM profiles in the cell lines used in this study. Cell lysates were immunoblotted with either TM antibody or with HA antibody. B, morphology of cells grown in monolayer, as observed by light microscopy (x40 magnification). C, immunofluorescence microscopy. DT cells expressing either HA-TM1 or TM1 were stained with a TM polyclonal antibody that recognizes all TM isoforms and Texas Red-conjugated phalloidin to visualize F-actin. Bar, 10 µm.

 
Morphologically DT cells are spindle-shaped, lack stress fibers, and are not contact-inhibited. Because TM1 induces reorganization of the cytoskeleton to restore stress fibers with attendant cell spreading, we examined whether HA-TM1 alters the cell morphology. Morphologically, DT/HA-TM1 cells resembled parental DT and empty vector transfected cells. DT/HA-TM1 cells displayed spindle-shaped morphology, were not subject to contact inhibition and formed multiple foci (Fig. 1B). Confocal microscopy revealed that HA-TM1 expression, unlike that of the wild type TM1 protein (18), did not induce the formation of stress fibers (Fig. 1C).

Because TM1 induces cytoskeletal reorganization with the assembly of stress fibers, it is likely that enhanced TM1 levels may increase the levels of F-actin. Therefore, we examined whether TM1 up-regulates F-actin levels and whether the inability of the variant protein to reorganize cytoskeleton is reflected in lower F-actin content. Overnight cultures of NIH3T3 cells, DT cells, DT/TM1, and DT/HA-TM1 cells were fixed, and the F-actin content was determined using TRITC-conjugated phalloidin. The fluorescent intensities were normalized to 1.5 x 105 cells (Table I). Consistent with the degree of microfilament organization, NIH3T3 cells contained most F-actin. DT and DT/HA-TM1 cells contained comparable amounts of F-actin, but exhibited significantly lower F-actin than NIH3T3 cells, as measured by fluorescence intensity (p < 0.02). Enhanced expression of TM1 resulted in the reemergence of microfilaments, which reflected in increased F-actin content. F-actin content was significantly higher in DT/TM1 cells (1.97 ± 0.32) compared with DT cells (1.01 ± 0.24) (p < 0.03). However, the total actin content in DT-derived cells was unchanged, when quantitated by immunoblotting and expressed relative to tubulin (Table I).


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  		TABLE I
TM1 enhances the cellular F-actin content F-actin content was determined by measuring the fluorescence of TRITC-labeled phalloidin, as described under "Experimental Procedures." Means ± S.D. are derived from three independent experiments. Relative fluorescence was normalized to 1.5 x 105 cells. Total actin and tubulin content were measured in 50 µg of protein from the indicated cell lines by immunoblotting. The ratio of actin to tubulin is taken as a measure of total actin content. The p values reported in the text are calculated by two-tailed paired t test.

 
Next we determined whether HA-TM1 altered the growth properties of DT cells. Variant TM1 did not affect either the monolayer growth or anchorage-independent growth rates of DT cells (Fig. 2, A and B). In contrast, wild type TM1 significantly decreased monolayer growth compared with HA-TM1 (p < 0.005), comparable with the NIH3T3 cells. Consistent with its inability to alter the cytoskeletal organization and morphology, HA-TM1 expression did not affect the anchorage-independent growth of DT cells. DT/HA-TM1 cells grew rapidly and formed colonies in soft agar as efficiently as the parental DT cells or vector control cells. In contrast, DT/TM1 cells failed to grow under anchorage-independent growth conditions (18). Thus, the N-terminal modification of TM1 abolishes the ability of TM1 to induce cytoskeletal reorganization and inhibits its anti-oncogenic properties.



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  		FIG. 2.
N-terminal modification abolishes anti-oncogenic effects of TM1. Growth of cells in monolayer (A) or under anchorage-independent conditions (B) was measured. A, the following cell lines were used: DT/TM1 ({blacksquare}), NIH3T3 ({blacktriangleup}), DT/HA-TM1 ({diamondsuit}), and DT (•). The p values were calculated using two-tailed t test assuming unequal variance. Photomicrographs of anchorage-independent cultures (B) are shown. The histogram shows the efficiency of colony formation in DT, two independent clones of DT/HA-TM1 and DT/TM1 cells. The error bars indicate mean ± S.D. from triplicate samples.

 
HA-TM1 Disrupts Microfilaments in Normal Fibroblasts— Next, we investigated whether the modification of the N terminus of TM1 would interfere with the ability of TM1 to associate with existing normal microfilament structures. NIH3T3 cells were transiently transfected with HA-TM1 (Fig. 3A). HA-TM1 incorporated into microfilaments and colocalized with F-actin. The HA-TM1 was distributed uniformly throughout the cytoskeleton, a finding consistent with a previous report (27). However, examination of the microfilament architecture after extended periods (48–72 h) revealed severe disruptions in microfilaments (Fig. 3B). Cells expressing HA-TM1 generally lacked well defined linear microfilaments that traverse the cell, which are typical of NIH3T3 cells. Instead, the transfected cells displayed wavy and disorganized microfilaments containing the variant protein and F-actin filaments, and aberrant cellular morphology when compared with the untransfected cells. These findings suggest that HA-TM1 associates with microfilaments and subsequently perturbs the cytoarchitecture (Fig. 3B).



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  		FIG. 3.
Microfilament organization in HA-TM1-expressing cells. Immunofluorescence was performed to detect the expression of HA-TM1 (anti-HA antibody) or TMs (TM antibody) and F-actin (phalloidin). NIH3T3 cells transiently transfected with HA-TM1 were stained at 24 (A) or 48–72 h (B). Stable cell lines of NIH3T3/TM1 cells (C) and NIH3T3/HA-TM1 (D) cells are also depicted. Bar, 10 µm.

 
Transient transfection typically produces an overabundance of the gene product, which could potentially result in artifacts. In this case, enhanced expression of a TM protein could perturb intracellular TM pools, leading to aberrant cytoskeleton, an effect independent of the N-terminal modification of TM1. To rule out the effects of overabundance of the transfected protein, we have examined the microfilament organization in NIH3T3 cell lines that were stably transduced with TM1 (NIH3T3/TM1 cells) (24). Enhanced expression of TM1 did not disorganize microfilaments, as did the variant protein (Fig. 3C). TM1 colocalized with linear, well defined stress fibers, and the normal cellular morphology was not compromised in NIH3T3/TM1 cells. These results indicate that enhanced TM1 expression per se does not induce disorganized cytoskeleton. Because TMs dimerize in "head to tail" fashion and associate with actin, modification of the N terminus of TM1 could have interfered with either dimerization or binding to actin, or both.

To investigate further the effects of the N-terminal modification of TM1 molecule, we have stably transfected NIH3T3 cells with HA-TM1 (Fig. 3D). Immunofluorescence experiments show that microfilaments are often "pushed" to a side, and a substantial amount of variant TM1 occupies the perinuclear area. Variant TM1 is also colocalized with phalloidin-positive microfilaments, indicating that during dynamic reorganization of stress fibers, the HA-TM1 associates with filaments. The variant protein, because of its abundance, may compete with the endogenous TM1, although the F-actin binding and dimerization properties of TM1 and variant TM1 significantly differ (see below). However, it is possible that HA-TM1 and endogenous TM1 coexist in the stress fibers.

Light microscopic observation of NIH3T3/HA-TM1 cells revealed the presence of high number of binucleated cells, indicating defects in cytokinesis in the transfected cells (Fig. 4). Asynchronously growing populations of NIH3T3/HA-TM1 cells contained as high as 11.91% (43 of 361 cells) of binucleated cells compared with <=1% (5 of 550 cells) found with the unmodified cells. Flow cytometry revealed a significantly higher number of cells with >4 N DNA content (Table II). NIH3T3, NIH3T3/TM1, and NIH3T3/HA-TM1 cells were analyzed for cell cycle progression under normal, serum-starved, and serum-stimulated conditions. Although the NIH3T3 and NIH3T3/TM1 cells did not differ in the number of hypertetraploid cells (2.9 ± 0.2 and 3.16 ± 0.03, respectively), NIH3T3/HA-TM1 cells contained 10.3 ± 1.38% cells with >4 N DNA content. Thus, the number of hypertetraploid cells in NIH3T3/HA-TM1 was significantly higher compared with parental and wild type TM1-expressing cells (p < 0.01). The number of hypertetraploid cells obtained by flow analysis is comparable with the number of binucleated cells obtained by visual counting (above). Serum starvation resulted in the accumulation of cells in G0-G1 phase in all three cell types. Although NIH3T3 and NIH3T3/TM1 cells contained a smaller percentage of cells in G2-M phase in starved samples (7.96 ± 0.31% and 10.12 ± 0.36%, respectively), the percentage of cells in G2-M phase in NIH3T3/HA-TM1 was much higher (26.52 ± 1.4%; p < 0.001 compared with NIH3T3 and NIH3T3/TM1 cells). Significantly, the number of cells with >4 N DNA content in NIH3T3/HA-TM1 cells dramatically decreased when the progression of cell cycle was inhibited by deprivation of serum (1.96 ± 0.46%). Serum stimulation, however, led to an increase in the hypertetraploid populations in NIH3T3/HA-TM1 cells (7.4 ± 0.35%) but not in NIH3T3 or NIH3T3/TM1 (0.98 ± 0.37 and 2.24 ± 0.22%, respectively; p < 0.001 compared with NIH3T3/HA-TM1 cells). Collectively, these results suggest that the cytokinesis was slower in NIH3T3/HATM1 cells and that perturbing the intracellular TM1 pool interferes with normal cytokinesis.



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  		FIG. 4.
HA-TM1 increases hypertetraploid cells. A representative binucleated, stable NIH3T3/HA-TM1 cell stained with anti-HA antibody and phalloidin is depicted. The numbers of binucleated cells were counted and the percentage is given. In control NIH3T3 cells the number of binucleated cells is <=1%.

 


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  		TABLE II
N-terminal modification of TM1 induces hypertetraploid The percent fraction of cells in each phase of the cell cycle was analyzed from either actively growing subconfluent cells or from serum-starved cultures for 24 h. Starved cultures were stimulated for 28 h. Accumulation of hypertetraploid cells is significantly higher in HA-TM1 cells compared with those expressing wild type TM1 or unmodified NIH3T3 cells. Data are shown as means ± S.D. from triplicate samples of a representative experiment. The p values reported in the text are calculated by two-tailed t test, assuming unequal variance.

 
The NIH3T3 cell lines selected for stable expression of the HA-TM1 extinguish the expression of the transfected TM1 upon a few passages, indicating that sustained expression of HA-TM1 is not compatible with cell growth. Whereas expression of HA-TM1 induces cytoskeletal disorganization and delays cytokinesis, it does not promote anchorage-independent growth in NIH3T3 cells (data not shown).

Variant TM1 Does Not Bind to F-actin—TM1 is a major actin-binding protein, and the functions of TM1, including suppression of transformed growth, may be dependent upon its ability to bind to and stabilize actin microfilaments. Therefore, we have examined HA-TM1 binding to actin in vitro. HA-TM1 was expressed in bacteria and purified, and actin-binding properties of the isolated protein were determined by cosedimentation. As a positive control for actin binding, we used unacetylated, recombinant (Escherichia coli-expressed) rat TM2. Unacetylated rat TM2 bound well to actin, with a binding constant in the micromolar range (23, 30, 33), similar to that of acetylated TM1. Unacetylated TM1 binds F-actin with lower affinity (23). Fig. 5 shows that TM2 bound to F-actin in a concentration-dependent and saturable manner with a Kapp of 2 x 106 M-1, consistent with published work (30). In contrast, HA-TM1 bound poorly to F-actin, too weakly to obtain a binding constant. Although the effect of other N-terminal extensions on TM1 function has not been investigated, it is well established that introduction of various N-terminal extensions on striated muscle {alpha}-tropomyosin overcomes the requirement for N-terminal acetylation for actin binding (29, 34, 35). In addition, striated muscle {alpha}-tropomyosin with N-terminal fusions, including HA, can incorporate into actin-containing structures in living cells (36, 37). Whereas the mechanism by which the HA epitope adversely influences TM1-actin affinity is unclear, the results are consistent with the inability of HA-TM1 to induce microfilaments and to suppress transformed growth, as well as with its interference with cytokinesis.



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  		FIG. 5.
N-terminal modification decreases TM1 binding to F-actin. Actin binding of bacterially purified HA-TM1 ({circ}) is measured by cosedimentation, as described under "Experimental Procedures." Wild type TM2 (•) is used as a control, which bound to actin with an affinity of Kapp 2 x 106 M-1. The TM/actin density ratio (an arbitrary number) versus free TM is plotted. The value at saturation for wild type TM2 represents stoichiometric saturation (1 TM:7 actins).

 
Dimerization and Interactions between Wild Type and Variant TM1 Proteins—TMs bind to actin as parallel, in register dimers (38, 39). Previous studies from this laboratory indicate that homodimers of TM1 form a stable component in cytoskeletal compartment in DT/TM1 cells (24). We examined whether actin binding of the variant TM1 is influenced by the ability of TM1 to associate as dimers, and whether dimerization is important in TM1-induced cytoskeletal reorganization.

First, we tested whether HA-TM1 forms homodimers by cross-linking the unique cysteine residue with a sulfhydryl cross-linker, DTNB, to stabilize the dimers in the lysates of metabolically labeled DT/HA-TM1 cells (Fig. 6A). Cross-linked samples were immunoprecipitated with either TM antiserum or monoclonal anti-HA antibody and analyzed by SDS-PAGE, either in the presence or absence of 2-mercaptoethanol in gel sample buffer by SDS-PAGE. In DT and DT/TM1 cells, homodimers of TM1 were readily evident when samples were analyzed using a buffer lacking 2-mercaptoethanol. The HA-TM1 protein expressed in DT/HA-TM1 cells, however, remained as monomer. However, by cross-linking using unlabeled cells, which measures steady state interactions, dimers of HA-TM1 were detected by both TM and HA antibodies in immunoblotting (Fig. 6B). These results indicate that nascently synthesized HA-TM1 either compartmentalizes differently or folds improperly preventing dimeric associations. Furthermore, HA-TM1 isolated from bacteria (same preparation used for binding assays, see above) also cross-linked into dimers (data not shown), indicating that dimerization alone may not ensure proper actin binding.



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  		FIG. 6.
Dimerization and interactions of HA-TM1. A, metabolically labeled cell lysates were cross-linked with DTNB, immunoprecipitated, and analyzed without or with 2-mercaptoethanol (EtSH) in gel sample buffer. TM1 dimers, monomeric TM1, and HA-TM1 are identified. B, unlabeled cell lysates (steady state) were cross-linked and immunoblotted with TM antibody. Dimeric and monomeric TMs are marked. C, immunoprecipitation (-ip) of TM1 and HA-TM1 in metabolically labeled cells. Although TM antibody immunoprecipitates TM1 and HA-TM1, the epitope tag antibody does not coimmunoprecipitate wild type TM1, indicating a lack of heterodimer formation. D, HA-TM1 interacts weakly with TM1 at steady state. Cell lysates from NIH3T3/HA-TM1 and DT/HA-TM1 were immunoprecipitated with TM or HA antibodies and immunoblotted, as indicated. Cell lysates were also run as controls.

 
Next we investigated whether the wild type and variant TM1 proteins interact with each other. Coimmunoprecipitation experiments were performed using metabolically labeled (Fig. 6C) and unlabeled cell lysates (Fig. 6D) to detect interactions at steady state. As expected, both proteins were immunoprecipitated with TM antibody. In labeled DT/HA-TM1 cells, however, the tagged protein did not coimmunoprecipitate with the endogenous protein as judged by immunoprecipitation with anti-HA antibody, suggesting a TM1:HA-TM1 heterodimer is not formed (Fig. 6C). Immunoprecipitation of unlabeled DT/HA-TM1 and NIH3T3/HA-TM1 cell proteins with anti-HA antibody resulted in coimmunoprecipitation of a modest amount of endogenous TM1, indicating a small amount of heterodimer formation (Fig. 6D).

Interestingly, during our efforts to isolate HA-TM1 from DT/HA-TM1 cells, we found that as the purification progressed, the enrichment of HA-TM1 on HA affinity columns also yielded the endogenous TM1, resulting in isolation of HA-TM1 and TM1 complexes (data not shown). This finding suggests that although HA-TM1 is capable of binding to TM1, such interactions do not occur efficiently in cells (Fig. 6, C and D).

TM1, but Not Variant TM1, Alters Cofilin Distribution— Stress fiber organization is controlled by Rho kinase, which regulates phosphorylation of myosin light chain kinase and LIM kinase. Whereas increased phosphorylation of myosin light chain kinase promotes contractility leading to microfilament reorganization, activated LIM kinase phosphorylates cofilin at Ser-3 and inhibits its severing action (reviewed in Ref. 40). Previous studies from this laboratory have shown that the Rho kinase pathway is essential for TM1-induced microfilament reorganization in ras-transformed cells (26). Down-regulation of LIM kinase, which appears to result in a significant increase in activated cofilin, has been implicated as a key mechanism in cytoskeletal disruption in ras-, and src-transformed cells (41, 42). Tropomyosins protect microfilaments from the severing and depolymerizing actions of cofilin (43, 44). Therefore, we considered whether TM1-induced cytoskeletal reorganization involves inhibition of cofilin-mediated microfilament depolymerization.

We measured activation status of cofilin in NIH3T3, DT, DT/TM1, and DT/HA-TM1 cells using phosphocofilin-specific antibodies. Normal NIH3T3 cells contained consistently the highest phosphocofilin to cofilin ratio, which indicates the least activity (Fig. 7A). The ratio of phosphocofilin:cofilin in NIH3T3 cells is taken as 100%. The ratio of phosphocofilin to total cofilin is significantly lower in DT (57.3 ± 11.6%, p < 0.05) and DT/HA-TM1 (68.9 ± 13.7%, p < 0.03) cells compared with NIH3T3 cells (100%) in five different experiments. However, TM1 expression in DT cells modestly elevated phosphocofilin content. In DT/TM1 cells phosphocofilin to cofilin ratio was found to be at 79.43 ± 15.49% when compared with NIH3T3 cells (p < 0.02) but does not appear to be significantly different from DT/HA-TM1 cells (p < 0.1). These results suggest that down-regulation of cofilin activity alone may not completely account for TM1-induced cytoskeletal organization.



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  		FIG. 7.
TM1, but not modified TM1, alters cofilin distribution. A, cellular phosphocofilin content is down-regulated in DT and DT/HA-TM1 cells. Cell lysates were immunoblotted with an antibody that recognizes phosphocofilin or total cofilin. The blot was also probed with a TM polyclonal antibody and anti-{alpha}-tubulin antibody. The ratio of phosphocofilin to total cofilin was calculated. The results (mean values) of five independent experiments, normalized to NIH3T3 as 100%, are presented as the phosphocofilin/cofilin ratio. The p values are calculated, based on paired two-tailed t test. B, TM1 and F-actin do not colocalize with cofilin. Indicated cell lines were stained for TM (using TM311 antibody), cofilin and F-actin. C, HA-TM1 and cofilin colocalize. NIH3T3 and DT cells were transiently transfected with HA-TM1 and stained to detect HA-TM1, cofilin, and F-actin. At earlier points of transfection (top panel), HA-TM1 and cofilin colocalize in transfected cells and appear to be excluded from microfilaments. Upon continued expression of the variant protein, stress fibers are displaced to the periphery of the cell, whereas cofilin and HA-TM1 colocalize to the cell body (middle panel). DT cells were also transfected with HA-TM1 and observed at 72 h (bottom panel). Cofilin and HA-TM1 were distributed throughout the cell body in transfected and untransfected DT cells. Bar, 10 µm.

 
Next, we have examined whether the subcellular distribution of cofilin is altered in TM1-induced cytoskeletal reorganization. In NIH3T3 cells, cofilin is distributed throughout the cytoplasm, as well as in the nucleus (Fig. 7B). Cofilin does not appear to associate with either F-actin or TMs present in stress fibers. In DT/HA-TM1 cells, however, cofilin and TMs are distributed in the cytoplasm; similar staining was found in DT cells (data not shown). In contrast, in DT/TM1 cells, which contain well developed microfilaments, TMs and cofilin do not colocalize, and the staining pattern resembles that of NIH3T3 cells.

Because high Mr TMs, including TM1, bind and stabilize microfilaments against the action of cofilin, we investigated whether HA-TM1-induced microfilament disruption is mediated by cofilin. NIH3T3 cells were transfected with HA-TM1 to test whether cofilin and variant TM1 colocalize. At earlier time points (24–48 h) of transfection, we could detect cofilin association with HA-TM1-positive aberrant microfilaments (Fig. 7C, top panel). However, the residual normal microfilaments lacked detectable staining of cofilin and HA-TM1. As the transfection time progressed, both of these proteins were localized to the perinuclear area, which lacked defined microfilament structures. In such cells stress fibers were displaced to the periphery and were stained positive for phalloidin alone. Cofilin and HA-TM1 were not detectable in those stress fibers (Fig. 7C, middle panel). It is likely that the linear filaments protected by the endogenous TM1 may exclude cofilin binding to the stress fibers. Transfection of DT cells with HA-TM1, however, did not result in the formation of microfilaments, and cofilin colocalized with the HA-TM1 throughout the cytoplasm (Fig. 7C, bottom panel). These results suggest that modified TM1, which interacts poorly with F-actin, allows cofilin binding to microfilaments and remodels the cytoskeleton.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
The important findings of this investigation are as follows. 1) The integrity of the N terminus of TM1 is critical for TM1-induced microfilament reorganization and tumor suppression. 2) Modification of the N terminus decreases TM1 binding to actin which allows cofilin-mediated cytoskeletal disorganization. 3) TM1 is required for normal cytokinesis. Suppression of TM1 in normal cells is difficult due to its abundance, and to date no mutants of TM1 have been described. Hence, the variant TM1 is a useful tool to study TMs, as it functions as a dominant negative mutant.

In this study we have focused on structural aspects of TM1 that mediate cytoskeletal reorganization and tumor suppression. It is intriguing that although TMs bind to F-actin in the micromolar range, and TM1 has no unique structural domains, TM1 exerts remarkably specific effects on cell growth compared with other TMs (17, 19, 45, 46). Subtle differences in TM-actin interactions modulate a range of cellular events that depend on cytoskeletal organization (1). Many in vitro binding studies, however, have shown that the N and C termini are critical determinants of TM binding to actin (47, 48). Some TMs, for instance, striated muscle {alpha}-tropomyosin and yeast TM, require N-terminal acetylation, which may stabilize the N terminus (49) for actin binding (23, 35, 50) and for the function of yeast TM in vivo (51, 52). The requirement for striated muscle {alpha}-tropomyosin and yeast TM for N-acetylation may be overcome by extending the N terminus by addition of a variety of peptide sequences (29, 34, 35, 37, 50). However, this is not the case with HA-TM1.

Other efforts to obtain insight into cellular TM functions have also examined the contribution of the N and C termini of TMs, utilizing chimeric TMs. Chimeras of non-muscle and muscle type TMs (TM5/3), consisting of TM5 and TM3, have been shown to bind F-actin avidly and cause accumulation of multinucleated cells resulting from delayed cytokinesis (48). Most interesting, cells expressing the chimera retained normal microfilament architecture. More recent studies using transgenic expression of chimeras of {alpha} and {beta} TMs have identified that the C terminal portion of TM is an important determinant of cardiac function (53).

Because TMs bind cooperatively in a head to tail fashion, modification of N terminus is sufficient to disrupt TM1 association with actin, even though the rest of the coding sequence and actin binding domains are identical to the wild type protein. The abundantly produced variant TM1, notwithstanding its lower affinity to actin, competes with the endogenous protein, associates with microfilaments, and subsequently disrupts microfilament architecture. Alternatively, HA-TM1 may compete for caldesmon (54), and thus could impact TM1-actin interactions, which also could disrupt the stability of the cytoskeleton.

Interaction of Wild Type and HA-TM1—Homodimers of TM1 are incorporated into the cytoskeletal compartment. The dimeric TM1 is hypothesized to be important for cytoskeletal reorganization and tumor suppression (24). It is interesting that although HA-TM1 retains the ability to dimerize and associate with endogenous TM1 in vitro, it fails to bind to F-actin in the cosedimentation experiments. Coimmunoprecipitation and cross-linking studies (Fig. 6) show that HA-TM1 homodimerizes and can interact in vitro with the endogenous TM1, a finding consistent with earlier studies (27). In vivo, however, HA-TM1 does not associate with the endogenous TM1, nor does the nascently synthesized protein dimerize. Several possibilities may explain this discrepancy.

First, N-terminal modification may alter subcellular localization of TM1. HA-TM1 segregates into a distinct subcellular location from microfilaments in NIH3T3 cells (Figs. 3D and 7C). Second, altered conformation and/or the inability to be recognized for certain post-translational modifications could account for the failure of the variant TM1 to be sorted with the endogenous TM1.

TM1 Regulates Cytokinesis—Because microfilaments provide the necessary force for cytokinesis (55), and the stability of actin microfilaments is controlled by TMs, alterations in TM-actin interactions could impact cell division. Expression of HA-TM1 remarkably increases the number of binucleated cells, indicating aberrant cytokinesis (Fig. 4 and Table II). Similar to the results reported here, a previous study showed that chimeric TMs exhibit higher binding affinity to F-actin and induce defective cytokinesis. These data suggest that the exact matching of the N and C termini is essential for normal functioning of TMs (48). Further work showed that the hTM5/3 chimeras induce altered motile behavior during the cytokinesis (56). These workers suggested the stronger binding of the chimeras with actin (57) would differentially regulate the contractile ring than would the wild type TMs. The defective cytokinesis could be the consequence of a generation of much higher force for separating daughter cells through interaction between the chimeras and actin than that produced by the wild type proteins (56). Because HA-TM1 interacts poorly with actin, HA-TM1 binding could yield lower than necessary force and hence delay the cytokinesis. This is supported by the fact that blockade of the cell cycle through serum starvation results in the completion of cytokinesis in HA-TM1 cells with a decrease in hypertetraploid populations. Serum starvation of NIH3T3/HA-TM1 cells, however, produces a higher percentage of cells in G2-M phase, compared with the parental and wild type TM1-expressing cells, indicating a slower cytokinesis.

Other researchers also report that high Mr TMs localize to the contractile ring during cell division (58). Collectively, these results suggest that both high and low Mr TMs are required for the normal completion of cytokinesis.

Role of Cofilin in Variant TM1 Induced Cytoskeletal Changes— TMs protect actin filaments from the gel severing and depolymerization actions of ADF/cofilin and gelsolin (12, 43, 59). Investigations into TM1-induced microfilament organization suggested that total gelsolin levels are not altered in TM1-expressing DT cells, suggesting that gelsolin may not be involved (26). Cofilin and TMs bind to actin in a mutually exclusive manner, and cofilin increases rate of depolymerization from the pointed end of actin filament and severs actin filaments (reviewed in Ref. 13). Recent studies have suggested that some TM isoforms may bind to cofilin (60) and inhibit actin nucleation and branching (11). Furthermore, genetic studies in Caenorhabditis elegans show that CeTM inhibits ADF/cofilin-mediated thin filament dynamics (43). LIM kinases, one of the effectors of Rho kinase signaling pathways, phosphorylate and inactivate cofilin (61–63). Our previous work (26) shows that Rho kinase signaling pathways are required to maintain the TM1-induced cytoskeleton, which indicates a role for cofilin in TM1-induced cytoskeletal dynamics.

We find that ras transformation decreases the phosphocofilin content (Fig. 7A), indicating that enhanced activation of cofilin may reshape the deregulation of cytoskeletal organization into a more dynamic and motile cytoskeleton. These results are in agreement with the published reports that show that neoplastic transformation uncouples Rho signaling pathways, enhancing cofilin activity (41, 42). Restoration of TM1 expression modestly elevates the phosphocofilin content in DT cells, indicating that additional mechanisms such as the altered intracellular compartmentalization may limit cofilin activity in TM1-induced cytoskeleton in ras-transformed cells. Support for this possibility comes from the immunofluorescence experiments that show that cofilin does not colocalize with F-actin in DT/TM1 cells or NIH3T3 cells, which contain well defined stress fibers. Functionally defective TMs, such as HA-TM1, however, appear to be unable to protect microfilaments, as evidenced by colocalization of cofilin in aberrant microfilaments. The variant TM1 and cofilin colocalize and are absent in normal stress fibers. Thus, TM1 is a key protein in stabilizing stress fibers against the action of cofilin-mediated remodeling. Similarly, TMs are suggested to limit cofilin to the leading edge of the cells, and where localized, dynamic remodeling of actin filaments occurs (44).

Alternate mechanisms of cytoskeletal reorganization by TM1 in ras-transformed cells, involving post-translational modifications, cannot be ruled out. Extracellular signal-regulated kinase-mediated phosphorylation of TM1 is suggested to be a signal for the assembly of stress fibers in endothelial cells (64). Because extracellular signal-regulated kinase signaling is constitutively activated in DT cells and the variant TM1 expression is unable to rescue stress fiber assembly, further work is needed to clarify the role of intracellular signaling in TM1-mediated cytoskeletal reorganization in transformed cells. Although we have shown that TM1 reorganizes microfilaments in oncogene-transformed cells (18, 20) and breast carcinoma cells (16, 17), other investigators have found that TM1 is unable to reorganize microfilaments in ras-transformed RIE cells (65) and neuroblastoma cells (66), suggesting a requirement for additional and as yet unknown factors in TM1-induced cytoskeletal reorganization.

In summary, we have identified that N-terminal integrity is a key regulator of TM1 functions. TM1-mediated cytoskeletal reorganization and tumor suppression may be dependent on TM1-F-actin interactions as well as on restricting the access of gel-severing proteins to actin filaments. Furthermore, TM1-induced cytoskeletal reorganization may involve inhibiting the gel severing activity of cofilin through modulation of phosphorylation status and preventing its association with microfilaments. Our results also suggest that TM1 is pivotal in maintaining the stress fibers.


   FOOTNOTES
 
* This work was supported in part by the Research and Development funds from the Department of General Surgery, a Cancer Center Pilot grant, and National Institutes of Health Grant GM 63257 (to S. H-D.). 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. Back

|| To whom correspondence should be addressed. Tel.: 336-716-2788; Fax: 336-716-2528; E-mail: gprasad{at}wfubmc.edu.

1 The abbreviations used are: TMs, tropomyosins; DT, doubly transformed; HA, hemagglutinin; PBS, phosphate-buffered saline; TRITC, tetramethylrhodamine isothiocyanate; PMSF, phenylmethylsulfonyl fluoride; DTNB, 5,5'-dithiobis-2-nitrobenzoic acid. Back


   ACKNOWLEDGMENTS
 
We thank Dr. Ethan Lange for the advice on statistical analysis. We thank Ken Grant for help with microscopy and Abhishek Singh for help with the actin binding assays.



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