Animal cells divide into two daughter cells by the formation of an actomyosin-based contractile ring through a process called cytokinesis. Although many of the structural elements of cytokinesis have been identified, little is known about the signaling pathways and molecular mechanisms underlying this process. Here we show that the human ECT2 is involved in the regulation of cytokinesis. ECT2 catalyzes guanine nucleotide exchange on the small GTPases, RhoA, Rac1, and Cdc42. ECT2 is phosphorylated during G2 and M phases, and phosphorylation is required for its exchange activity. Unlike other known guanine nucleotide exchange factors for Rho GTPases, ECT2 exhibits nuclear localization in interphase, spreads throughout the cytoplasm in prometaphase, and is condensed in the midbody during cytokinesis. Expression of an ECT2 derivative, containing the NH2-terminal domain required for the midbody localization but lacking the COOH-terminal catalytic domain, strongly inhibits cytokinesis. Moreover, microinjection of affinity-purified anti-ECT2 antibody into interphase cells also inhibits cytokinesis. These results suggest that ECT2 is an important link between the cell cycle machinery and Rho signaling pathways involved in the regulation of cell division.

Cellular division into two daughter cells occurs through nuclear division and cytokinesis. In cytokinesis, formation of an actomyosin-based contractile ring separates the cytoplasm to two daughter cells. A small GTPase RhoA appears to be localized in the cleavage furrow during cytokinesis (Takaishi et al. 1995). In lower eukaryotes, such as sand dollar, Dictyostelium, and Xenopus, involvement of Rho GTPases in cell division has been reported (Kishi et al. 1993; Mabuchi et al. 1993; Larochelle, 1996; Drechsel et al. 1997). In Saccharomyces cerevisiae, cell division is achieved by budding, which is controlled by Cdc42 (Chant and Pringle 1991). In contrast, in mammalian cells, where Rho GTPases also regulate cell surface cytoskeletal organization, the role of Rho GTPases in cell division is not well-documented.

The Rho family of small GTPases, represented by RhoA, Rac1, and Cdc42, act as molecular switches of diverse biological functions involving cytoplasmic actin remodeling (Hall 1998). The GTP-bound form of Rho proteins is active, whereas the GDP-bound form is inactive. Activation of the Rho proteins is promoted by guanine nucleotide exchange factors (GEFs), which catalyze the replacement of bound guanosine diphosphate (GDP) by GTP. The GTP-bound form of Rho proteins can specifically interact with their effectors or targets and transmit signals to downstream molecules. Rho proteins are inactivated through the hydrolysis of bound GTP to GDP by intrinsic GTPase activity, assisted by GTPase activating proteins (GAPs). In fibroblasts, RhoA, Rac1, and Cdc42 regulate formation of actin stress fibers, membrane ruffling, and filopodia, respectively (Hall 1998). The upstream stimuli that regulate these GTPases are largely unknown, although lysophosphatidic acid, PDGF, and Ras have been shown to regulate the cytoskeletal changes controlled by these GTPases (Hall 1998).

We previously isolated the ect2 oncogene in a search for mitogenic signal transducers in epithelial cells, where a murine keratinocyte expression cDNA library was introduced into fibroblasts to induce foci of morphologically transformed cells (Miki et al. 1991). Removal of the NH2-terminal half of Ect2 strongly activates the transforming activity of ect2, and injection of ect2 transfectants into nude mice efficiently induces tumor formation. The central domain of Ect2 has sequence similarity with the dbl oncogene (Miki et al. 1993), which has been shown to catalyze guanine nucleotide exchange on the Rho family of small GTPases (Hart et al. 1991). Ect2 associates with a subset of the Rho family proteins: RhoA, RhoC, and Rac1 (Miki et al. 1993).

In this study, we show that human ECT2 catalyzes guanine nucleotide exchange on Rho proteins. ECT2 is phosphorylated in a G2/M phase-specific manner, and phosphorylation is required for the exchange activity of ECT2. In interphase cells, ECT2 is mainly localized in the nucleus. However, in mitotic cells, ECT2 is localized predominantly in the midzone, where the formation of cleavage furrow starts. We found that the inhibition of ECT2 by expression of a dominant negative mutant or microinjection of anti-ECT2 antibody specifically blocks the completion of cytokinesis, resulting in multinucleated cells.

Materials and Methods

Guanine Nucleotide Exchange Assays

The open reading frame of human ECT2 was introduced into the mammalian expression vector pCEV32F3 (Lorenzi, et al. 1999) to express a FLAG–ECT2 fusion protein. COS-7 cells were plated in 100-mm dishes and transfected with 10 μg of plasmid DNA with Lipofectamine (GIBCO BRL). Transfected cells were cultured for 48 h, harvested, and lysed in 1 ml of cold lysis buffer (25 mM Hepes, pH 7.5, 0.3 M NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 1% Triton X-100, 0.5% sodium deoxycholate, 20 μg/ml each of leupeptin and aprotinin, and 100 μg/ml PMSF). FLAG–ECT2 fusion proteins were immunoprecipitated from the lysates with 9 μg/ml of anti-FLAG mAb (Sigma Chemical Co.) and protein G–Sepharose beads (Amersham Pharmacia Biotech). Guanine nucleotide exchange assays were performed essentially as described (Horii et al. 1994) using these immunoprecipitates. In brief, 3 μg of GDP-loaded recombinant GTPases were incubated with 5 μM [35S]GTPγS (0.25 mCi mmol−1) and 10 μl of protein G beads suspension in 190 μl of exchange buffer (20 mM Tris-HCl, pH 7.5, 10 mM MgCl2, 0.5 mM DTT, 100 mM NaCl, 0.5 mg/ml BSA). At the indicated times, 30 μl of the reaction was removed and passed through nitrocellulose filters. Filters were washed and then counted in a liquid scintillation counter. For phosphatase treatment, immunoprecipitates were incubated with recombinant VHR protein (Ishibashi et al. 1992) or λ phosphatase (New England Biolabs) for 30 min at 30°C, and then used for exchange assays.

Preparation of Anti-ECT2 Antibodies

GST–Ect2 fusion protein was expressed in Escherichia coli and used for immunizing rabbits. The NH2-terminal half (ECT2-N; amino acids 1–421) or Dbl homology domain (DH; amino acids 414–639) of human ECT2 was expressed as fusion proteins with thioredoxine and oligohistidine using the pET-32 vector (Novagen). Anti-ECT2 antibodies were prepared by passing antiserum through affinity columns coupled with the corresponding human ECT2 proteins using AminoLink Plus Immobilization Kit (Pierce). Anti–ECT2-DH recognized a single ECT2 protein of ∼100 kD. Anti–ECT2-N also recognized the endogenous ECT2 protein, although some additional bands were weakly detected.

Analysis of ECT2 Modification during Cell Cycle Progression

HeLa cells were grown in DMEM (GIBCO BRL) supplemented with 10% FCS in 7% CO2 at 37°C. Cells were synchronized at the G1/S boundary by a thymidine-aphidicolin double block (Golsteyn et al. 1995). In brief, cells were incubated with 2 mM thymidine for 14 h, released from arrest, and then arrested at G1/S again with aphidicolin (1 μg/ml) (Sigma Chemical Co.). Cells were then placed under normal growth conditions (time 0). After 6 h, nocodazole (final concentration 100 ng/ml) was added to arrest the cells at prometaphase. Mitotic cells were collected by mechanical shake-off and replated in normal growth medium. Following trypsinization, samples were analyzed by flow cytometry using FACS® II instrument (Becton Dickinson). For phosphatase treatment, G1 or M phase arrested cells were lysed in TNE buffer (10 mM Tris-HCl, pH 7.8, 1% NP-40, 0.15 M NaCl, 1 mM EDTA pH 7.0, 10 μg/ml aprotinin) and incubated with 1 μg/ml of recombinant VHR protein for 30 min at 30°C. Samples were resolved by 6% SDS-PAGE, transferred to Immobilon-P membranes (Millipore), and analyzed by immunoblotting using affinity-purified anti-ECT2 antibodies.

Subcellular Localization of ECT2

To determine the localization of endogenous ECT2, HeLa cells were grown on coverslips, fixed for 10 min with 3.7% formaldehyde in PBS, permeabilized for 10 min with 0.2% Triton X-100 in PBS, and then incubated with 1% BSA in PBS for 10 min. For costaining of endogenous ECT2 and tubulin, coverslips were incubated with affinity-purified anti–ECT2-DH antibody and anti–β-tubulin (TUB2.1; Sigma Chemical Co.) mAbs for 1 h at 4°C. After washing, samples were incubated with FITC-conjugated anti–rabbit IgG and Cy3-conjugated anti–mouse IgG (Jackson ImmunoResearch, Inc.) for 1 h at room temperature. To express green fluorescent protein (GFP)-fused ECT2 proteins in U2OS cells, ECT2-N, ECT2-C, and ECT2-F were introduced into pEGFP-C1 (Clontech). After transfection, cells were fixed and proteins visualized by green fluorescence using a Zeiss Axiovert microscope equipped with a Photometrics digital camera and IPLab software (Signal Analytics Co.).

Microinjection

HeLa cells were grown on poly-l-lysine–coated glass coverslips for 18–24 h until the cultures reached ∼50% confluency. The coverslips were transferred into fresh medium containing 10 mM Hepes, pH 7.6, before injection. Antibodies were dialyzed and concentrated to 1 mg/ml using a Centricon-30 concentrator (Amicon). Rabbit IgG (Sigma Chemical Co.) was used as a control. Microinjection into the cytoplasm was performed using Eppendorf semiautomatic microinjection apparatus. Immediately after injection, coverslips were washed with PBS and incubated in growth medium for the indicated time period.

Results

ECT2 Is a GEF for RhoA, Rac1, and Cdc42

To examine the function of ECT2 as a Rho GEF, we expressed ECT2 as a FLAG epitope-tagged protein in COS cells immunoprecipitated with anti-FLAG mAb, and used immunoprecipitated protein for exchange assays. Full-length ECT2 efficiently stimulated nucleotide exchange on three representative members of the Rho GTPases, RhoA, Rac1, and Cdc42 in vitro, whereas similar immunoprecipitates from vector transfectants showed no significant activity (Fig. 1 b, left panel). In contrast to the activity on Rho GTPases, ECT2 did not exhibit significant exchange activity on two Ras family GTPases, H-Ras and Rap1A (data not shown).

ECT2 Is Phosphorylated Specifically in G2 and M Phases of the Cell Cycle

The predicted ECT2 protein also contains a novel domain homologous to yeast S phase cyclin Clb6 (Schwob and Nasmyth 1993; Stuart and Wittenberg 1998), and two tandem repeats of the recently identified BRCA1 COOH-terminal repeat (BRCT) motif (Fig. 1 a) that is widespread throughout checkpoint and repair proteins (Saka et al. 1994; Bork et al. 1997; Zhang et al. 1998). Since these structural similarities suggest the possibility that ECT2 is involved in cell cycle control, we examined in detail the expression pattern of ECT2 during the cell cycle. HeLa cells were synchronized at the G1/S boundary, and the expression of ECT2 protein was examined by immunoblotting after the release from G1 arrest. Although we did not observe significant changes in the overall expression level of ECT2 protein during progression of the cell cycle, a shift of the mobility of ECT2 was detected by anti-ECT2 antibody (Fig. 1 c, left panel). A slowly migrating protein appeared at 9 h as cells entered G2 phase. At 14 h, as cells entered M phase, the slowly migrating band predominated. The fraction of slowly migrating protein gradually decreased as cells entered G1 phase (Fig. 1 c). To determine whether the shift of mobility of ECT2 was due to protein phosphorylation, we treated M phase–arrested cells with the tyrosine/serine/threonine phosphatase VHR. No mobility shift was detected in M phase cells after VHR treatment (Fig. 1 c, left panel). Therefore, ECT2 appeared to be phosphorylated specifically at G2 phase, and this state was manifested through M phase.

Phosphorylation Is Required for the Exchange Activity of ECT2

To further examine the effect of phosphorylation on the exchange activity of ECT2, we treated FLAG–ECT2 immunoprecipitates with various concentrations of VHR. As shown in the right panel of Fig. 1 b, the exchange activity of ECT2 was inhibited by VHR in a dose-dependent manner, and the incubation for 1 h with 0.5 μg/ml of VHR decreased the activity of FLAG–ECT2 to background levels. Inhibition of exchange activity was also observed by treatment of FLAG–ECT2 with λ phosphatase, and this inhibitory effect was strongly prevented by a phosphatase inhibitor vanadate (data not shown). These results suggest that phosphorylation is required for the exchange activity of ECT2 in vitro.

ECT2 Colocalizes with the Mitotic Spindle and Midbody in M Phase

Next, we examined the subcellular localization of endogenous ECT2 during cell cycle progression by immunofluorescent analysis of HeLa cells. Interestingly, in interphase cells ECT2 was located predominantly in the nucleus, where no expression of Rho family proteins has been reported (Fig. 2 a). After nuclear membrane breakdown in prometaphase, ECT2 spread into the cytoplasm (Fig. 2 b). During metaphase, ECT2 accumulated in the regions where the mitotic spindle is present (Fig. 2 c). Costaining of ECT2 and tubulin generated the yellow colocalization signal. As cells entered anaphase, ECT2 appeared to concentrate in the central region of the spindle (Fig. 2 d). In late anaphase and telophase, ECT2 was mainly located in the midzone, where the cleavage furrow is formed (Fig. 2 e). During cytokinesis, ECT2 accumulated at the midbody, a region in the middle of the bridge that connects the two daughter cells (Fig. 2 f). When the cells exited mitosis and the nuclear envelope reassembled, ECT2 translocated to the nucleus again (data not shown). To locate the region of ECT2 responsible for midbody localization, we transfected cells with plasmids containing the NH2-terminal half (ECT2-N), the COOH-terminal half (ECT2-C), or full-length (ECT2-F) ECT2 as GFP fusion proteins. GFP proteins were visualized 72 h after transfection (Fig. 3 A). Like endogenous ECT2, GFP-tagged ECT2-F was detected in the midbody of dividing cells. Although ECT2-N localized in the cytoplasm of interphase cells (data not shown), it accumulated in the midbody of dividing cells. In contrast, ECT2-C was detected in the cytoplasm of mitotic cells. These observations indicate that the NH2-terminal half of ECT2 is required for midbody localization. The nuclear localization of ECT2-F in interphase cells may be attributed to nuclear localization signals located in the central region of ECT2 (Fig. 1 a).

Expression of ECT2 NH2-Terminal Domain Inhibits Cytokinesis

The dynamic change of subcellular localization and G2/M-specific modification of ECT2 suggest a possible role of ECT2 in cell division. If ECT2 controls cell division through the regulation of Rho GTPases, expression of ECT2 mutants containing the domain that is required for localization, but lacking the catalytic domain, may induce a dominant negative phenotype. Therefore, we closely examined the morphology of cells expressing ECT2-N. Surprisingly, ∼60% of cells expressing ECT2-N became multinucleated, with most cells containing two nuclei 72 h after transfection (Fig. 3 B). Division of ECT2-N–expressing cells appeared to proceed to a stage just before the separation of two daughter cells with single nuclei (Fig. 3 A, panels c and e), suggesting that a very last step of cytokinesis was inhibited by ECT2-N expression. In contrast, cells expressing ECT2-F, ECT2-C, or GFP expression vector alone did not exhibit such phenotypes. Since most of the cells expressing ECT2-N underwent normal nuclear division (Fig. 3 A, panels d and f), it is unlikely that these multinucleated cells were generated as a consequence of aberrant nuclear division. These results suggest that ECT2-N can function as a dominant negative mutant to inhibit cytokinesis.

Microinjection of Anti-ECT2 Antibodies Specifically Inhibits Cytokinesis

To further examine the involvement of ECT2 in cytokinesis, we inhibited ECT2 function by microinjection of affinity-purified anti-ECT2 antibodies into asynchronously growing HeLa cells (Fig. 4). Whereas cells injected with control IgG divided normally, ∼60% of cells injected with anti–ECT2-DH, which recognizes the catalytic domain of ECT2, became larger with double nuclei 24 h after injection. Since 25% of the injected cells had not divided at this stage, ∼80% of the cells were multinucleated upon completion of mitosis. After 48 h, 29% of anti–ECT2-DH–injected cells contained three or four nuclei. The presence of cells with more than two nuclei may indicate inhibition of multiple rounds of cytokinesis with anti–ECT2-DH. In contrast, very few control IgG-injected cells were multinucleated. To rule out the possibility that anti-ECT2 cross-reacted with other molecules that regulate cytokinesis, we prepared a second affinity-purified antibody that specifically recognizes the NH2-terminal domain of ECT2. Microinjection of this antibody (anti–ECT2-N) also strongly inhibited cytokinesis (Fig. 3 A, panels g–i; Fig. 3 B). These results strongly suggest that ECT2 plays a critical role in cytokinesis.

Discussion

The results from this study provide details about the biochemical and biological functions of the ECT2 protooncogene product. We detected a mobility shift of ECT2 at G2/M phases. Treatment of the ECT2 immunoprecipitates with VHR dual specificity phosphatase resulted in shift back of the band to the normal size. This strongly suggests that ECT2 is modified by phosphorylation. We found that dephosphorylation of ECT2 by phosphatases reduced the exchange activity in vitro in a dose-dependent manner. Therefore, ECT2 appears to be activated by phosphorylation, which occurs specifically in M phase. Since we could not detect phosphorylated ECT2 with antiphosphotyrosine antibodies (our unpublished results), ECT2 may be phosphorylated by serine/threonine kinases. Candidate protein kinases that can activate ECT2 may include Cdc2-regulated kinases and Cdc2 itself. The predicted amino acid sequence of ECT2 contains two consensus Cdc2 phosphorylation sites (amino acids 327–330 and 814–817). Further studies on ECT2 phosphorylation, including determination of phosphorylation sites and generation of phosphorylation-deficient mutants, will clarify the activation mechanisms of ECT2.

Rho proteins were found to regulate actin polymerization in studies on cytoskeletal organization of fibroblasts (Hall 1998). During cytokinesis in animal cells, an actin-based contractile ring divides the cell into two daughter cells. We found that ECT2 catalyzes nucleotide exchange on Rho proteins and this activity appears to be dependent on G2/M phase-specific phosphorylation. ECT2 was released to the cytoplasm after the nuclear membrane breakdown and then accumulated in the midbody during cytokinesis. Since ECT2-N localized in the midbody, but ECT2-C did not, ECT2-N appears to determine the midbody localization of ECT2. We found that expression of ECT2-N strongly inhibited cytokinesis. Because ECT2-N localizes in the midbody but cannot catalyze nucleotide exchange, it appears to function as a dominant negative mutant. Moreover, microinjection of affinity-purified anti-ECT2 antibody also inhibited cytokinesis. Therefore, ECT2 might play an important role in cytokinesis. The nuclear localization of ECT2 may suggest another role of ECT2 in mitosis. This possibility is under investigation. However, the inhibition of cytokinesis with ECT2-N or anti-ECT2 might not be attributed to the inhibition of mitosis, because ECT2-N, which lacks nuclear localization signals, was expressed in the cytoplasm and anti-ECT2 was injected into the cytoplasm.

The presence of cell cycle regulator-related domains and G2/M phase-specific phosphorylation of ECT2 suggest the regulation of ECT2 activity by cell cycle machinery. Therefore, ECT2 might be an important molecule linking cell cycle machinery to Rho signaling pathways involved in the regulation of cytokinesis. In Xenopus, multiple Rho proteins are involved in cytokinesis: whereas local activation of Rho is important for proper constriction of the contractile furrow, Cdc42 plays a role in furrow ingression (Drechsel et al. 1997). In Drosophila, cytokinesis is developmentally controlled during embryogenesis and is omitted during the initial nuclear division cycles. It starts in somatic cells with mitosis 14, but is blocked with mutations in the pebble gene (Lehner 1992). Recently, it was reported that pebble encodes an ECT2-related protein, which can interact with Drosophila Rho1 GTPase (Prokopenko et al. 1999), suggesting that ECT2 is also involved in cytokinesis in Drosophila.

In mammalian cells, Rho GTPases also regulate cortical actin remodeling (Hall 1998). Inhibition of RhoA in normal rat kidney cells revealed that Rho regulates the completion of cytokinesis possibly through modulating the mechanical strength of the cortex (O'Connell et al. 1999). Although the role of Rho in division of mammalian cells is not well-documented, recently it was reported that two Rho effectors, Rho-associated kinase and Citron kinase, play a role in cytokinesis (Goto et al. 1998; Madaule et al. 1998). Further studies on ECT2, Rho GTPases, and their effectors should clarify the signal transduction pathways involved in the control of cell division.

Acknowledgments

We thank Dr. D. Lowy for support; Dr. D.P. Bottaro (National Cancer Institute, Bethesda, MD) for VHR phosphatase; V. Kapoor for technical assistance; Dr. G. Vande Woude for valuable suggestions; and Drs. D.P. Bottaro and S.O. See for critical reading of the manuscript.

T. Tatsumoto was supported by a Japan Society of the Promotion of Science fellowship for Japanese Biomedical and Behavioral Researchers at the National Institutes of Health.

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T. Tatsumoto and X. Xie contributed equally to this paper.

Abbreviations used in this paper: BRCT, BRCA1 COOH-terminal repeat; DAPI, 4′,6-diamidino-2′-phenylindole; DH, Dbl homology domain; ECT2-C, ECT2 COOH-terminal half; ECT2-F, full-length ECT2; ECT2-N, ECT2 NH2-terminal half; GDP, guanosine diphosphate; GEF, guanine nucleotide exchange factor; GFP, green fluorescent protein.