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Cell, Tumor, and Stem Cell Biology |
1 Endocrinology and Metabolism and 2 Biostatistics, University of Virginia Health System, Charlottesville, Virginia; 3 Department of Surgery, Kyungpook National University Hospital, Daegu, South Korea; 4 Neuroscience Research Institute, University of California, Santa Barbara, California; 5 Core Endocrine Laboratory, Pennsylvania State University College of Medicine, Hershey, Pennsylvania; and 6 Natural Products Research Institute, College of Pharmacy, Seoul National University, Seoul, South Korea
Requests for reprints: Richard J. Santen, Department of Internal Medicine, University of Virginia, P.O. Box 801416 Charlottesville, VA 22908. Phone: 434-924-2207; Fax: 434-924-1284; E-mail: rjs5y{at}virginia.edu.
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Abstract |
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Introduction |
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Women who have developed secondary resistance to one hormonal therapy may often respond to a second- or third-line hormonal therapy (1). However, nearly all women ultimately become completely resistant and require alternative treatment approaches, which generally involve cytotoxic chemotherapy (1). Because of the side effects and toxicity associated with chemotherapy, therapy targeted to more specific intracellular signaling events is a major focus of current research. The crosstalk between estrogen receptors (ER) and growth factor receptor pathways in secondarily resistant ER + breast cancer cells has suggested the use of inhibitors of growth factor pathways rather than chemotherapy at this stage of the disease (8). Both preclinical and early-phase clinical trials are testing various combinations of inhibitors and particularly those targeting the EGF-R, HER-2, and mammalian target of rapamycin (mTOR) pathways. Potential drawbacks are that these agents can produce significant toxicity in the heart and other organs, and that treatment might not be focused on targets that are up-regulated in the individual patient. Based on these concepts, a relatively nontoxic agent that would act upon multiple targets but exert minimal toxicity would be a welcome addition to the therapeutic armamentarium.
During the course of our studies, we identified unexpected properties of a drug, called tetramethoxystilbene (TMS), a synthetic derivative of the herbal product, rhapontigenin (9). Rhapontigenin, a naturally occurring stilbene, is a potent mechanism-based inactivator of cytochrome P450 1A1(10). Trans-stilbene compounds have been chemically modified with the goal of creating anticancer agents with a higher potency of inhibition against cytochrome P450 1B1, an enzyme that catalyzes the formation of potentially genotoxic estradiol metabolites (9–12). Naturally occurring stilbenes have dihydroxyl groups on their phenyl ring. Because substitution of dihydroxyl for dimethoxy groups was thought to increase the lipophilicity and binding to the active sites of the P450 family 1 enzymes, TMS was synthesized such that dimethoxy groups were placed on the 3 and 5 positions of the phenyl ring. TMS, a methoxy derivative of 2,4,3',5'-tetrahydroxystilbene, showed potent and selective inhibition of P450 1B1 (9–11). The original rationale of our studies was to evaluate the effect of TMS on the conversion of estradiol to 4-OH-estradiol and ultimately on tumor formation (12). However, we observed that TMS was highly effective in reducing cell number in breast cancer cells in tissue culture. In an attempt to uncover the mechanisms responsible for this effect, we discovered "off-target" actions including a marked inhibition of microtubule polymerization, substantial induction of apoptosis, and effects on several growth factor signaling molecules.
This manuscript describes the effects of TMS on microtubule polymerization, cell cycle dynamics, growth factor pathways, and apoptosis. We found that this agent induces apoptosis in 80% to 90% of cells within a 48-h period and inhibits breast cancer cells with secondary hormonal resistance more effectively in vivo and in vitro than hormone-dependent cells. Based on its lack of apparent toxicity in intact animals, its antitumor effects in vitro and in vivo, and its potent actions on cell signaling pathways, we believe that this agent could potentially serve as an important therapeutic agent for the treatment of women who develop secondary resistance to hormonal therapy.
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Materials and Methods |
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. Cell culture medium (IMEM) was from Biosource International, Inc. Fetal bovine serum (FBS), glutamine, and trypsin were from Invitrogen. z-VAD was purchased from Calbiochem and was dissolved in dimethylsulfoxide (Sigma Chemical Co.). Dextran-coated charcoal-stripped FBS (DCC-FBS) was prepared as previously described (2). Cell culture conditions. Wild-type MCF-7 cells were grown in IMEM containing 5% FBS. Tamoxifen-resistant (TAM-R) MCF-7 cells were treated with tamoxifen (10–7 M) continuously for more than 1 year with a change in medium every 2 to 3 days (13). ICI 182,780 (ICI)–resistant MCF-7 cells were treated with ICI (ICI 10–8 M) continuously over a period of more than 1 year. Long-term estradiol-deprived (LTED) cells were developed as previously described (14) and maintained routinely in phenol red–free IMEM supplemented with 5% DCC-FBS.
Assessment of cell growth. Cells were plated in six-well plates at a density of 60,000 cells per well in their culture media. Two days later, the cells were treated as described in figure legends for 5 days with media change on day 3. Nuclei were prepared by sequential addition of 1 mL HEPES-MgCl2 solution (0.01 mol/L HEPES and 1.5 mmol/L MgCl2) and 0.1 mL ZAP solution [0.13 mol/L ethylhexadecyldimethylammonium bromide in 3% glacial acetic acid(v/v)] and counted using a model Z1 Coulter counter (Coulter Corp).
Cell proliferation. To specifically assess cell proliferation, 3H-thymidine incorporation into DNA was assessed by methods previously described (15).
Plasma estradiol assay. Estradiol was measured with a previously described RIA method involving purification with column chromatography and use of an iodinated estradiol trace (Diagnostic Products Corp.). Assay sensitivity is 5 pg/mL, and interassay precision averages 11% at a concentration of 32 pg/mL (16).
Immunoblotting. Cells were grown to 80% confluence in 60-mm dishes and lysed with buffer containing 20 mmol/L Tris (pH, 7.5), 150 mmol/L NaCl, 1 mmol/L EDTA, 1 mmol/L EGTA, 1 mmol/L sodium orthovanadate, 2.5 mmol/L sodium PPi, 1% Triton X-100, 1 mmol/L ß-glycerophosphate, 1 µg/mL leupeptin and aprotinin, 1 mmol/L phenylmethylsulfonyl fluoride. Cells were then pulse sonicated for 30 s at room temperature and centrifuged at 14,000 rpm for 10 min. Cell lysates were stored at –80°C. The total protein concentration was determined using a standard Bradford assay reagent (Bio-Rad). Western blots were carried out using primary antibodies dissolved in PBS containing 5% bovine serum albumin (BSA); secondary antibody conjugated with horseradish peroxidase (1:2,000); and SuperSignal West Pico Chemiluminescent Substrate (Pierce) to identify bands on X-ray film.
Apoptotic cell death detection by ELISA. To quantitate histone-associated DNA fragments (mono- and oligonucleosomes) in vitro in the cytoplasm of MCF-7 cells undergoing early apoptosis, 
8 x 104 MCF-7 cells were plated per well into 12-well plates. After 2 days, the cells were treated with either DMSO, 3 µmol/L TMS, or 10 nmol/L 17ß-estradiol + 3 µmol/L TMS. After 24 h, floating cells and adherent cells were collected for analysis. A cell death detection ELISAPLUS apoptosis kit (Roche) was used as described by the manufacturer. Experiments were done in triplicate, and P values were determined using SigmaPlot (Systat Software Inc.).
Measurement of DNA degradation via terminal nucleotidyl transferase–mediated nick end labeling assay. Apoptotic MCF-7 cells were detected by terminal nucleotidyl transferase–mediated nick end labeling (TUNEL) using the APO-BRDU Complete Flow Cytometry Analysis Kit (Phoenix Flow Systems, Inc.) according to the manufacturer's protocol. Controls included positive and negative cells provided in the kit following the manufacturer's protocol.
Cell cycle distribution. MCF-7 cells were grown in IMEM + 5% FBS. Cells were treated with either DMSO or TMS for 12, 24, 36, or 48 h. Both floating and adherent cells were collected and filtered through a 40-µm cell strainer (BD Falcon) to remove clumps. Cells were counted, and 1 x 106 cells were hypotonically lysed in 1 mL of DNA staining solution [3.4 mmol/L Tris (pH, 7.6), 0.075 mmol/L propidium iodide (PI), 0.1% NP40, and 700 units/L RNase A] for 10 min on ice. Samples were then analyzed on a FACSCalibur flow cytometer (Becton Dickinson), and cell cycle distribution and percentage of cells within the sub-G1 peak were determined using Modfit software (DNA Modelling System, Verity Software House, Inc.).
Annexin V assay. MCF-7 cells were treated with either DMSO or 3 µmol/L TMS for 12, 24, 36, or 48 h. Floating cells were collected by centrifugation and added to adherent cells, which were gently lifted off the tissue culture plate with Accutase (Innovative Cell Technologies, Inc.). A total of 20,000 MCF-7 cells were labeled with Annexin V–Alexa 488 antibody and PI using an apoptosis detection kit (Invitrogen) and analyzed on a FACScalibur flow cytometer (Becton Dickinson). The distribution of apoptosis was determined using Cell QuestCell software (Becton Dickinson), and cells that were Annexin V (–) and PI (–) were considered viable cells. Cells that were Annexin V (+) and PI (–) were considered early-stage apoptotic cells. Cells that were Annexin V (+) and PI (+) were considered late-stage apoptotic cells. Cells that were PI (+) and Annexin V (–) were considered necrotic.
Immunofluorescence microscopy. MCF-7 cells were grown for 24 h on coverslips pretreated with poly-lysine to enhance cell flattening and adherence and then incubated in the presence or absence of TMS (0.1, 1, 5, 10, and 50 µmol/L) for 20 h. Control cells were incubated with vehicle alone. Cells were fixed in 10% formalin (20 min, 25°C) and permeabilized in methanol (–20°, 10 min). Nonspecific antibody staining was blocked with 20% normal goat serum in PBS, and cells were incubated with DM1a anti–-tubulin antibody (Sigma Chemicals; 1:1,000 dilution) followed by goat anti-mouse FITC-conjugated secondary antibody (Sigma Chemicals; 1:500 dilution) for 1 h at 37°C to visualize microtubules. Nuclei were stained with 4',6-diamidino-2-phenylindole (0.2 µg/mL in PBS; Sigma Chemicals) for 1 min. Coverslips were mounted with Prolong Antifade (Molecular Probes) and viewed and photographed with a 60x objective using an ORCA II digital camera driven by Metamorph software (Universal Imaging) on a Nikon Eclipse E800 fluorescence microscope.
Purification of tubulin and microtubule polymerization. Microtubule protein preparations consisting of tubulin and microtubule-associated proteins were isolated from bovine brain by three cycles of polymerization and depolymerization. Tubulin was purified from the microtubule protein by phosphocellulose chromatography, drop-frozen in liquid nitrogen, and stored at –70°C (17). On the day of use, tubulin was thawed on ice and centrifuged (17,000 x g, 20 min, 4°C) to remove aggregated or denatured tubulin. Protein concentration was determined by Bradford assay using BSA as standard. For polymerization, tubulin (2.75 mg/mL) was mixed with a range of concentrations of TMS in 100 mmol/L PIPES, 1 mmol/L MgCl2, 1 mmol/L EGTA, and 1 mmol/L GTP. Microtubule seeds were prepared with the same solution (without TMS) plus 10% glycerol and 10% DMSO by incubating at 30°C for 30 min and shearing the assembled microtubules six times through a 25G 7/8 in needle. Microtubule polymerization was initiated by adding seeds to the tubulin solution and was monitored at 30°C by light scattering at 350 nm using a Beckman DU 640 spectrophotometer.
Mouse xenograft model. Tumor fragments (30–40 mg) of TAM-R MCF-7 human mammary tumors were implanted s.c. as xenografts in oophorectomized nude mice (Charles River Labs) aged 4 to 5 weeks at the start of the experiment. Animals containing continuously passaged tumors in tamoxifen-treated animals were obtained as a gift from V.C. Jordan (Fox Chase Cancer Center, Philadelphia, PA; ref. 18). Estradiol was delivered to produce plasma levels of 280 pg/mL with an "estradiol clamp," silastic capsule method previously described (19). Treatment with s.c. silastic capsules containing 30 mg TMS was started when tumors reached a measurable size (300 mm3), about 6 months after inoculation. Tumors were measured at least weekly with calipers, and volumes were calculated using the formula 4/3
x r12 x r2 (r1 < r2) during 8 weeks. All animal experiments were conducted under Federal and Institutional guidelines and approved by the University of Virginia Animal Care and Use Committee.
Statistical analysis. Statistical analysis was done by using the R statistical package.7 A mixed-effect linear model with two fixed effects (day and drug) and a random effect (mouse) was fitted, and the statistical significance of the drug effect was examined with the significance level 0.05. In particular, log10 transformation was used for MCF tumor volume data to produce a linear relationship between tumor volumes and days.
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Results |
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TMS inhibited the wild-type MCF-7 and MDA-MB-468 cells by 50% at a concentration of 1 µmol/L with greater suppression at higher doses (Fig. 1A
). The benign MCF10A cells were inhibited by 50% at a similar concentration, but with a plateau at higher doses. All three secondarily resistant cells lines were inhibited by 50% at TMS concentrations of 0.1 to 0.3 µmol/L (Fig. 1B). With increasing doses, the cells exhibited a 70% to 90% reduction in cell number in a dose response fashion with increasing concentration of TMS until a plateau was reached at 0.3 to 1 µmol/L (see Fig. 1B).
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We then questioned whether the TMS effects might represent generalized toxicity to the breast cancer cells with resulting increase in necrotic cell death, as evidenced by trypan blue uptake at 24 h of exposure to TMS. Surprisingly, a large fraction of the secondarily resistant cells detached from the culture plates (see Supplementary Fig. S1) in response to TMS. The parental MCF-7 cells seemed less sensitive to this effect of TMS, as shown above for inhibition of cell growth (Fig. 1A). Although detached, these cells were still able to exclude trypan blue, a finding that excluded necrotic cell death and provided evidence of continued viability (Fig. 1D).
Effects of TMS on tubulin polymer mass and mitosis. The detachment of viable cells at 24 h suggested that TMS might exert specific effects on cellular structures that mediate adherence to culture plate surfaces. Resveratrol, a stilbene with a structure similar to TMS, had been reported to interfere with microtubule function (21). Accordingly, we examined the effect of TMS on microtubule polymerization. A microtubule polymerization assay was used to study this phenomenon. Concentrations of TMS ranging from 0.3 to 3.0 µmol/L inhibited microtubule polymerization in a time and dose response fashion, with a maximum inhibition of 50% of vehicle control (Fig. 2A ).
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TMS induces G2-M cell cycle arrest. Blockade of microtubule polymerization can result in arrest of the cell cycle at the S or G2-M phase depending on the specific microtubule polymerization agent used (23). We carried out fluorescence-activated cell sorting analysis using PI to stain DNA and examined MCF-7 cells that had been treated for 12, 24, 36, and 48 h in the absence or presence of 3 µmol/L TMS. We found that at 36 h, a large percentage of the TMS-treated cells were arrested in the G2-M phase of the cell cycle when compared with the control DMSO-treated cells (Fig. 3A , compare G1 with G2-M peaks). The increased G2-M peak is consistent with what has been reported for the microtubule inhibitor nocodazole (24). In addition, after 48 h treatment with 3 µmol/L TMS, we observed a sub-G1 peak that is apparent in Fig. 3A (bottom right), suggesting the possibility of apoptosis. Modfit was used to quantitate the sub-G1 peak, and an increase from 2% at 12 h to a maximum of 20% at 36 and 48 h (Fig. 3B) was observed. The sub-G1 peak was negligible in the DMSO-treated control cells.
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TMS induces apoptosis in breast cancer cells. The presence of viable (i.e., trypan blue excluding) floating cells at 24 h and the sub-G1 peak on flow cytometry suggested that TMS triggers apoptosis. To distinguish apoptosis from necrosis at later time points, we examined Annexin V and PI staining followed by flow cytometry. We found that at 12 h, only 6% of the TMS-treated cells were apoptotic (see Supplementary Fig. S3, compare DMSO-treated to TMS, sum of early and late apoptosis). At 24 h, apoptosis increased to 12%, and at 36 and 48 h, apoptosis increased to 40%. During each time point, the maximum percent of necrotic cells in the TMS-treated groups was 0.73% compared with 0.88% in the DMSO-treated cells (Supplementary Fig. S3, compare top left quadrant DMSO to TMS).
As further evidence of apoptosis, we carried out an anti-histone/DNA monoclonal antibody ELISA assay, which examines double-strand DNA breaks occurring at an early stage. Three cell lines, parental, LTED, and Tam-R, were treated with either ethanol, 3 µmol/L TMS, or 3 µmol/L TMS + 10 nmol/L E2 for 24 h and assayed by ELISA. We found that addition of 3 µmol/L TMS resulted in an increase in the level of dsDNA and nucleosomes present in the cytoplasm (Fig. 4A–C compare lanes 1 to 2). Because it is known that the microtubule active agent, paclitaxel, is inhibited in the presence of estradiol, we also tested whether the addition of estradiol would inhibit the action of TMS in these cell lines (25, 26). When 10 nmol/L E2 was added in the presence of TMS, there was no decrease in the level of apoptosis for the parental or LTED cells (Fig. 4A and B, compare lanes 2 with 3) and only a slight decrease in the TAM-R cells (Fig. 4C, compare lanes 2 with 3). Most, but not all, types of programmed cell death involve activation of caspases (27). For this reason, we examined the effects of the pan-caspase inhibitor, z-VAD, and showed blockade of TMS-elicited apoptosis, a finding providing strong evidence for the presence of caspase-induced apoptosis (Fig. 4A–C, compare lanes 2 to 4).
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Most types of apoptosis involve the cleavage of poly(ADP-ribose) polymerase (PARP) as molecular signature of apoptosis (28). We therefore examined the effect of TMS on the cleavage of this protein and the ability of the pan-caspase inhibitor z-VAD to block these effects. TMS caused an increase in the p85 cleavage product of PARP, an effect blocked by z-VAD (see Supplementary Fig. S4). As an indication of the mechanism of apoptosis, the c-jun-NH2-kinase (JNK) inhibitor (SP600125) also blocked PARP cleavage, but the p38 inhibitor (SB 203580) did not.
TMS inhibits multiple signaling pathways. Disruption of microtubules is known to trigger a sequence of downstream signaling events resulting in apoptosis (29). Accordingly, we next sought to identify the signal transduction pathways involved in regulating cell survival and cell death using Western blot techniques. Activated JNK and p-38 increased substantially with initial increases at concentrations of TMS of 0.3 and 3 µmol/L, respectively. As evidence of specificity of its effects, phospho-MAPK and total MAPK levels did not change with TMS. However, we did detect reduced phosphorylation of Akt, FAK, and 4-E-BP-1(T-70) at concentrations above 1 µmol/L (Fig. 5A ).
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Effects of TMS on a breast xenograft model. We implanted tumor fragments (30–40 mg) of TAM-R MCF-7 human mammary tumor cells into both flanks of athymic mice. Six months after implantation of tumors, the xenografts reached measurable size (300 mm3). The group (n = 5) treated with vehicle alone grew substantially over the 56-day period of observation. Two months of continuous treatment with TMS, using 30 mg s.c. implants (n = 4, reduced tumor volume by 53% when compared with control (P < 0.02). In the parental MCF-7 cells xenograft model, the 30-mg TMS implant (n = 16) reduced tumor volume by 41.7% when compared with control (n = 17; Fig. 6A
), but this effect did not reach statistical significance. Animals receiving the 30-mg implant of TMS for 2 months tolerated this procedure well and exhibited no apparent systemic toxicity. As evidence of this, there were no changes in total body weight, uterine weight (Fig. 6B) ovarian weight, and plasma estradiol levels and no changes in liver and renal histology (see Supplementary Figs. S6 and S7).
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Discussion |
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The effect of TMS on apoptosis in vitro was especially dramatic with 80–90% of cells undergoing programmed cell death by 48 h of exposure. The magnitude of this process required verification that cell death did not merely represent necrosis, and that apoptosis was indeed present. Accordingly, our study used multiple methods to provide definitive evidence of programmed cell death and the lack of necrotic toxicity. These included the use of an ELISA apoptosis assay, flow cytometry coupled with the BrdUrd-based TUNEL and Annexin V methods, demonstration of a sub-G1 peak on flow cytometry, blockade of apoptosis with the pan-caspase inhibitor z-VAD, examination of the formation of the 89-kDa PARP cleavage product and its blockade with z-VAD, and exclusion of necrotic cell death with a trypan blue viability test. To further rule out necrotic cell death, we used the Annexin V flow cytometry–based assay and showed that a maximum of only 0.73% of cells were necrotic under all conditions examined. Taken together, these various methods provided compelling evidence of the presence of apoptosis and absence of significant necrotic cell death.
One mechanism whereby TMS induces apoptosis seems to be related to its effects on microtubule polymerization. Agents such as colchicines and nocodazole block microtubule polymerization in a manner similar to TMS and also trigger apoptosis (30). Exactly how apoptosis occurs is not completely understood, but a marked up-regulation of p-38 kinase and JNK occurs in association with this phenomenon. A suggested mechanism is that microtubule inhibitors release MLK-2 from binding to specific sites on tubulin and free it to act as an upstream kinase (MAP KKK) in the JNK kinase cascade sequence (31). JNK itself is known to mediate apoptosis under certain circumstances. Our data showing blockade of PARP cleavage with a JNK inhibitor support a role for JNK in the apoptotic process. These observations, taken in concert, would suggest that blockade of microtubule polymerization might be a primary effect of TMS and result in the secondary activation of JNK and p38 kinase. However, additional primary effects of TMS on other targets such as FAK, Akt, and mTOR might also be possible (32, 33). Further experiments are necessary to precisely determine which effects of TMS are primary and which are secondary to more upstream events.
Treatment of neoplastic diseases with other microtubule-interacting agents such as the taxanes and Vinca alkaloids provide effective therapy but are associated with substantial toxicity (30, 34). Our data with TMS revealed no apparent systemic toxicity at concentrations which effectively blocked growth of secondarily resistant breast cancer xenografts in vivo. Parameters used to assess systemic toxicity included total body weight, plasma estradiol levels, uterine and ovarian weight, and histologic changes in the liver or kidney (Supplementary Fig. S7). More detailed studies will be required to assess toxicity in greater depth and particularly neurotoxicity.
An important consideration is the relative efficacy and toxicity of TMS compared with another class of microtubule active agents, the taxanes, because both compounds exert similar actions. TMS inhibits phosphorylation of FAK, Akt, and mTOR and stimulates JNK and p38 MAPK activity. Paclitaxel also inhibits Akt and increases JNK and p38 MAPK activity (35, 36). However, paclitaxel acts differently than TMS in that it blocks microtubule depolymerization and causes stabilization of these structures. Comparative studies on antitumor efficacy, toxicity, and organ specificity will be required to fully address potential differences between these agents. An important consideration, however, is the observation that the herbal parent of TMS, rhapontigenin, has been used in oriental medicine for more than three millennia without untoward toxicity. While speculative, this would suggest that TMS, a derivative of rhapontigenin, might be relatively nontoxic as well.
Our current studies are intriguing but still contain weaknesses that we plan to address in the near future. For example, we have not as yet conducted rigorous experiments to determine if wild-type breast cancer cells of several derivations (i.e., ZR-75-1, T 47D, cloned MCF-7, or others) also seem to be less sensitive to TMS than their secondarily resistant counterparts. This could be important for concluding that TMS might be an ideal agent for treating hormone-resistant disease. However, because hormone therapy is effective initially in receptor-positive breast cancer, it is unlikely that TMS would be chosen to replace this modality of therapy. Thus, it is not essential to our conclusions that hormone-resistant cells be more sensitive to TMS than parental cells. Another issue we considered is that we did not examine DNA laddering, which is usually required for demonstration of apoptosis when using a new antiapoptotic agent. However, this technique requires the activation of caspase 3, a factor lacking in MCF-7 cells (37). Finally, it will be important in future studies to carefully examine whether TMS induces apoptosis via mitochondrial or death receptor pathways and to dissect out the precise mechanisms by which apoptosis occurs.
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Acknowledgments |
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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.
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Footnotes |
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H. Park and S.E. Aiyar are co–first authors.
Received 1/15/07. Revised 2/28/07. Accepted 4/18/07.
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References |
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out of the nucleus and enhances its interaction with epidermal growth factor receptor in MCF-7 breast cancer cells. Cancer Res 2007;67:1352–60. and vascular endothelial growth factor A through phosphatidylinositol 3-kinase/Akt/FRAP pathway in OVCAR-3 and A2780–70 human ovarian carcinoma cells. Toxicol Appl Pharmacol 2004;196:124–35.[CrossRef][Medline]
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), 0.3 µmol/L (
), 1 µmol/L (
), 3 µmol/L (
). Microtubule polymerization was inhibited in a TMS concentration–dependent manner. TMS (3 µmol/L) inhibited polymer mass by 50%. B, effects of TMS on microtubule organization in cells. Immunofluorescence images of microtubules (green) and DNA (blue) in fixed cells in the presence or absence of TMS. a and b, untreated control cells. The remaining photos illustrate TMS-treated cells: c, 1 µmol/L TMS; d, 10 µmol/L TMS; e, 50 µmol/L TMS; f, 10 µmol/L TMS. In control mitotic cells (a), spindles were bipolar, and all chromosomes were congressed to a compact metaphase plate. However, in the presence of 1 µmol/L TMS (c), mitotic cells contained multiple asters of very short microtubules and chromosomes were disorganized. Interphase control cells (b) and interphase cells incubated with 1 µmol/L (d) or 10 µmol/L TMS (e) exhibited microtubules, but the cells were less well spread in the presence of TMS, and the overall microtubule polymer was reduced. Microtubules were depolymerized, and the cytoplasm was severely reduced after incubation with 50 µmol/L TMS (e). TMS incubation induced frequent multinucleated cells (d, top cell, and f). Bar, 5 µm.
