G-protein–coupled estrogen receptor 1 (GPER1) has been reported to play a significant role in mediating the rapid estrogen actions in a wide range of normal and cancer cells. G-1 was initially developed as a selective agonist for GPER. However, the molecular mechanisms underlying the actions of G-1 are unknown, and recent studies report inconsistent effects of G-1 on the growth of breast cancer cells. By employing high-resolution laser scanning confocal microscopy and time-lapse imaging technology, as well as biochemical analyses, in the current study, we provide convincing in vitro and in vivo evidence that G-1 is able to suppress the growth of breast cancer cells independent of the expression status of GPERs and classic estrogen receptors. Interestingly, we found that triple-negative breast cancer cells (TNBC) are very sensitive to G-1 treatment. We found that G-1 arrested the cell cycle in the prophase of mitosis, leading to caspase activation and apoptosis of breast cancer cells. Our mechanistic studies indicated that G-1, similar to colchicine and 2-methoxyestradiol, binds to colchicine binding site on tubulin, inhibiting tubulin polymerization and subsequent assembly of normal mitotic spindle apparatus during breast cancer cell mitosis. Therefore, G-1 is a novel microtubule-targeting agent and could be a promising anti-microtubule drug for breast cancer treatment, especially for TNBC treatment. Mol Cancer Ther; 16(6); 1080–91. ©2017 AACR.

Estrogens play critical roles in the regulation of many physiological and pathological processes, including the development of various types of cancer (1). It has long been thought that the actions of estrogen are mediated by estrogen receptor α (ERα) and estrogen receptor β (ERβ). Recent evidence indicates that G-protein–coupled estrogen receptor 1 (GPER, also called GPR30) is able to mediate the rapid actions of estrogen in numerous tissues and cells (2). G-1 {1-[4-(6-bromobenzo[1,3]-dioxol-5yl)-3a,4,5,9b-tetrahydro-3H-cyclopenta[c]quinolin-8-yl]-ethan one} was developed as a selective agonist of GPER to distinguish the estrogen effects mediated by GPER from those mediated by classic nuclear estrogen receptors (ERα and ERβ; ref. 3). The original report showed that G-1 was able to bind to GPER with high-affinity and did not bind to classic nuclear ERs and other 25 examined G-protein coupled receptors (3). Since its first report in 2006, G-1 has been wildly used to investigate the functions of GPER in various tissues and organs (4).

Current research results considering effects of G-1 on cancer cell proliferation and survival are highly controversial. Magiollini and colleagues (5, 6) reported that G-1 stimulates the growth of ovarian and endometrial cancer cells. Supporting these results, several studies indicated that G-1 promotes proliferation of endometrial cancer and seminoma cells (7, 8). However, accumulating evidence indicated that G-1 is a potent tumor growth inhibitor. Recent studies demonstrated that G-1 suppresses proliferation and induces apoptosis in numerous types of cancer cells, including: ovarian cancer (9–11), adrenocortical carcinoma (12), lung cancer (13), cervical cancer (14), and prostate cancer (15).

Breast cancer, which accounts for 23% of the total diagnosed cancer cases and 14% of the total cancer-related deaths among females, is the most common type of cancer in women worldwide (16). Prolonged exposure to estrogens represents a major risk factor for the progression of breast cancer. In breast cancer cells, 17β-estradiol (E2) triggers stimulatory effects by binding to the nuclear ERs that regulate the expression of genes, which contribute to cancer cell proliferation, migration, and survival (17). Because GPER is a membrane ER and G-1 is an agonist of GPER, it is implicated that G-1 promotes survival and growth of breast cancer cells. However, research results considering effects of G-1 on breast cancer cell growth are also very inconsistent. Several reports show that G-1 (from 10 nmol/L to 1 μmol/L) simulates the proliferation of breast cancer cells (6, 18, 19). Other studies show that G-1 potently suppresses breast cancer cell growth at micromolar levels (20–23).

Although previous reports show conflicting outcomes regarding the effects of G-1 on the growth and survival of cancer cells, the majority of these studies postulate that the actions of G-1 are mediated by activation of GPER (5–8, 14, 15, 18–22, 24). However, it is surprising that none of these studies actually provide direct evidence that G-1, via binding to GPER, activates G-proteins. Therefore, the controversial results may be attributed to the fact that the molecular mechanism by which G-1 regulates cancer cell growth is still unclear. The aim of this study is to investigate effects of G-1 on the growth of breast cancer cells and to define the molecular mechanisms underlying the actions of G-1. Our studies show that G-1 is able to arrest breast cancer cell cycle in the prophase of mitosis and induces cancer cell death independent of the expression status of ERs. Moreover, our mechanistic studies demonstrate that G-1 binds to the colchicine-binding site of tubulin protein to block microtubule assembly, leading to failure of spindle formation, cell-cycle arrest, and apoptosis.

Chemicals and reagents

G-1 and G-15 were purchased from Tocris Bioscience. Antibodies against human cleaved caspase-3, caspase-3, cleaved caspase-7, caspase-7, cleaved PARP, PARP, cyclin B1, phosphorylated histone H3 (S10), phosphorylated CDC2 (Y15), phosphorylated CDC25C (T48), and phosphorylated CDC25C (S216) were from Cell Signaling Technology Inc.; β-actin antibodies were from Sigma; Alexa-conjugated second antibodies were from Molecular Probes, Inc.; horseradish peroxidase–conjugated second antibodies were from Jackson Immunoresearch Laboratories Inc.; the Vybrant MTT Assay Kit was from Invitrogen; The Caspase-Glo 3/7 assay Kit was purchased from Promega; The microtubule sedimentation assay products were purchased from Cytoskeleton, Inc. All other molecular-grade chemicals were purchased from Sigma, Thermo Fisher Scientific, or United States Biochemical.

Cell lines and culture

Breast cancer cell lines MCF-7, SK-BR-3, BT-549, Hs 578T, HCC 1937, and MDA-MB-231 were purchased from the ATCC, authenticated by the ATCC using STR polymorphism analysis, and used within 6 months after receipt. The MCF-7 cell line expresses ERs, progesterone receptors (PR), and GPERs. The SK-BR-3 cell line expresses GPER but not classical ERs. The MDA-MB-231 cell line lacks of ERs, PRs, and HER2. This triple-negative breast cancer (TNBC) cell line does not express GPER (or very low expression of GPER if any). Hela-GFP cells were from Dr. Jixin Dong. Cells were cultured in DMEM medium with 10% FBS and 1% penicillin–streptomycin (10,000 U/mL, Gibco) in a 37°C, humidified cell culture incubator supplied with 5% CO2. Cells were treated in the fresh phenol red-free DMEM supplemented with or without steroids-free FBS for indicated time points.

Cell proliferation assay

The effect of G-1 on breast cancer cell proliferation was analyzed as described before (10). Briefly, half confluent cells were cultured with DMSO (control) or different concentrations of G-1 for indicated time. Cell morphology was pictured before trypsin addition with an Olympus IX71 inverted microscope equipped with a DP73 digital camera. Cells were then trypsinized and the live cell number was counted by using a Countess Automated Cell Counter.

Anchorage-independent cell growth assay

Anchorage-independent growth of breast cancer cells were analyzed with a method established in our laboratory as described previously (25).

Xenograft mouse model

MDA-MB-231 cells (5 × 106 cells in 100-μL PBS for each mouse) were injected subcutaneously into the right dorsal flank of 6-week-old female athymic nude mice. The animal handling and all experimental procedures were approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Nebraska Medical Center. Two weeks after injection, small xenografts were formed in all mice. We randomly distributed 12 mice in two groups with 6 mice in each group. One group mice were injected with G-1 (5 mg/kg) in sesame oil every day for 2 weeks, whereas in the other group, mice were injected with only sesame oil at the same time. The tumor size was measured (shortest diameter and longest diameter) weekly and the tumor volume was estimated as following: volume = (shortest diameter)2 × longest diameter × 3.14/6. Mice were sacrificed in the fourth week and the xenografts were harvested, weighted, and processed for preparation of frozen sections.

Fluorescent IHC and immunocytochemistry

Expression of Ki67 in tumor xenografts was determined by fluorescent IHC on frozen sections to examine the effect of G-1 on the proliferation of breast cancer cells in vivo (26). Fluorescent ICC was also used to determine tubulin distribution and phosphorylated histone H3. Images were captured with a Zeiss 710 Meta Confocal Laser Scanning Microscope and analyzed with the Zeiss Zen 2010 software.

Flow cytometry to detect cell apoptosis and cell cycle

For cell apoptosis analysis, cell staining, and fixation were performed using an Annexin V-FITC Apoptosis Kit as described previously (27). Cell apoptosis and cell cycle were analyzed using flow cytometry as described previously (28).

Western blot analysis

Western blot analysis was performed to determine protein expression and activation using methods established in our laboratory (29).

Caspase-3/7 activity assay and MTT assay

Caspase-3/7 activity was determined as described previously (10). MTT assay was performed using the Vybrant MTT Assay Kit with a protocol described previously (30).

In vitro tubulin polymerization assay

G-1 (20 μmol/L), paclitaxel (20 μmol/L) or DMSO vehicle was mixed with X-rhodamine–labeled tubulin in G-PEM buffer (with glycerol, Cytoskeleton Inc.) and incubated at 37°C for 20 minutes for polymerization. Microtubules was monitored using a Zeiss 710 Meta Confocal Laser Scanning Microscope (Carl Zeiss Microscopy, LLC).

Microtubule sedimentation assay

Purified bovine tubulin was incubated at 37°C for 20 minutes with or without 20 μmol/L G-1. Taxol (20 μmol/L) was used as a microtubule polymerization control. The polymerized microtubule filaments were precipitated by centrifugation at 100,000 × g for 30 minutes at room temperature. The pellet and supernatant was fractioned with a SDS-PAGE, transferred to nitrocellulose membranes, and stained with SYPRO Ruby protein blot stain. The images were captured and quantified with a UVP gel documentation system (UVP LLC).

Time-lapse video microscopy for microtubule dynamics and cell division

Hela cells expressing GFP-labeled tubulin were seeded in Nunc Lab-Tek II Chambered Cover glass (Thermo Fisher Scientific) and imaged in a live-cell imaging system using Zeiss 710 Meta Confocal Laser Scanning Microscope with 63× oil objective. Time-lapse image series were acquired at 3-minute intervals for microtubule dynamics and 5-minute intervals for cell division experiments.

Colchicine-binding scintillation proximity assay

The ability of G-1 binding to the colchicine-binding site in tubulin was examined using a CytoDYNAMIX screen 15 assay kit (Cytoskeleton Inc.) in accordance with the manufacturer's instruction and previous description. Biotin-labeled tubulin (0.5 μg) in 10 μL of reaction buffer was mixed with [3H]colchicine (0.08 μmol/L, PerkinElmer) and the test compounds (positive control colchicine, negative control vinblastine, G-1, fluorescent G-1, or 2-ME) in a 96-well plate (final volume: 100 μL). After incubating for 2 hours at 37°C with gentle shaking, streptavidin-labeled yttrium SPA beads (80 μg in 20 μL reaction buffer, PerkinElmer) were added to each well and incubated for 30 minutes at 4°C. The radioactivity was determined using Packard TopCount Microplate Scintillation Counter (Packard Instrument).

Statistical analysis

All experiments were repeated at least three times unless otherwise mentioned. The statistical analyses were performed by using GraphPad Prism software (GraphPad Software, Inc.) and quantitative data were analyzed using Student t test and one-way ANOVA with Tukey post-test. A P value of <0.05 was considered to be significant.

G-1 inhibits breast cancer cell growth in an ER-independent manner

MCF7 (ERα+, ERβ+, and GPER+), SK-BR-3 (ERα, ERβ, and GPER+), and MDA-MB-231 (ERα, ERβ+, and GPER− or Low) breast cancer cell lines were used as cellular models to evaluate the effect of G-1 on the growth of breast cancer cells. Regardless of the expression status of ERs, breast cancer cells treated with 2 μmol/L G-1 detached from the culture plates within several hours and eventually died (Supplementary Fig. S1A). Quantitative studies indicated that G-1 inhibited breast cancer cell growth in a concentration- and time-dependent manner (Fig. 1A and B; Supplementary Fig. S1B and S1C). Lower concentrations of G-1 (1–10 nmol/L) had no obvious effect on the growth of breast cancer cells in either FBS-supplemented or FBS-free media. However, 100 nmol/L of G-1 significantly inhibited growth of breast cancer lines in FBS-free media (Supplementary Fig. S1B). When concentrations approached micromolar levels, G-1 consistently suppressed breast cancer cell proliferation and viability (Fig. 1A and B; Supplementary Fig. S1B and S1C). G15, a previously reported selective GPER antagonist (31), was not able to block the suppressive effects of G-1 on breast cancer cell proliferation (Supplementary Fig. S1B and S1C). Interestingly, the proliferation of MCF7, but not SK-BR-3 and MDA-MB-231 cells, was slightly stimulated by G15 (Supplementary Fig. S1B and S1C). This slight increase in cell proliferation may be attributed to the nonspecific binding of G15 to ERα in MCF cells, an event that has been reported to weakly activate ERE in this cell line (32).

Figure 1.

G-1 inhibits the proliferation of breast cancer cells independent of the expression status of GPER and classic ERs. A, G-1 inhibits the proliferation of breast cancer cells in a concentration-dependent manner. Cells were treated with increasing concentrations of G-1 for 48 hours. Cell numbers were counted using an automatic cell counter and presented as percentages of that of the control group (0.1% DMSO treated). Each bar represents mean ± SEM of four repeats. Bars with different letters are significantly different from each other (P < 0.05). B, G-1 inhibits the proliferation of breast cancer cells in a time-dependent manner. Cells were treated with 2 μmol/L G-1 for different time. Cell numbers were presented as percentages of that of control group (0 hours group). Each bar represents mean ± SEM of four repeats. Bars with different letters are significantly different from each other (P < 0.05). C, Representative images showing colony formation of breast cancer cells in the presence of different concentrations of G-1 for 7 days. Quantitative data show the colony numbers. Colonies were stained with MTT; scale bar, 200 μm. CTRL: 0.1% DMSO treatment. ***, P < 0.001 compared with control. D, Representative images showing the nude mice bearing breast cancer xenograft treated with or without G-1 for 2 weeks (left), and the harvested tumors (right). Red circles indicate xenograft location and size. CTRL: sesame oil control treatment daily for 2 weeks; G-1: 5 mg/kg G-1 treatment every other day for 2 weeks. E, Tumor xenograft growth curves in athymic nude mice treated with or without G-1. The xenograft volume was measured weekly. Student's t test was used to compare the tumor volumes between control and G-1–treated groups. Each point represents mean ± SEM (n = 6); *, P < 0.05. F, The weight of tumor xenografts from nude mice treated with or without G-1 for 2 weeks. Each bar represents mean ± SEM (n = 6). **, P < 0.01 compared with control. G, Representative images showing expression of Ki67 in control and G-1–treated tumor xenografts determined by fluorescent IHC. Ki67 was labeled in green; actin was stained with rhodamine-phalloidin (red); nuclei were stained with DAPI (blue); scale bar, 20 μm. The number of Ki67-positive cells and total cells in control and G-1–treated groups were also counted under a fluorescent microscope, and the ratio of Ki67-positive cells was calculated. Each bar represents mean ± SEM (n = 6), ***, P < 0.001. H, The body weight of athymic nude mice with or without G-1 treatment. Each bar represents mean ± SEM (n = 6). Student t test was used to do statistical analysis. No statistical significance on the body weight was observed between control and G-1–treated groups.

Figure 1.

G-1 inhibits the proliferation of breast cancer cells independent of the expression status of GPER and classic ERs. A, G-1 inhibits the proliferation of breast cancer cells in a concentration-dependent manner. Cells were treated with increasing concentrations of G-1 for 48 hours. Cell numbers were counted using an automatic cell counter and presented as percentages of that of the control group (0.1% DMSO treated). Each bar represents mean ± SEM of four repeats. Bars with different letters are significantly different from each other (P < 0.05). B, G-1 inhibits the proliferation of breast cancer cells in a time-dependent manner. Cells were treated with 2 μmol/L G-1 for different time. Cell numbers were presented as percentages of that of control group (0 hours group). Each bar represents mean ± SEM of four repeats. Bars with different letters are significantly different from each other (P < 0.05). C, Representative images showing colony formation of breast cancer cells in the presence of different concentrations of G-1 for 7 days. Quantitative data show the colony numbers. Colonies were stained with MTT; scale bar, 200 μm. CTRL: 0.1% DMSO treatment. ***, P < 0.001 compared with control. D, Representative images showing the nude mice bearing breast cancer xenograft treated with or without G-1 for 2 weeks (left), and the harvested tumors (right). Red circles indicate xenograft location and size. CTRL: sesame oil control treatment daily for 2 weeks; G-1: 5 mg/kg G-1 treatment every other day for 2 weeks. E, Tumor xenograft growth curves in athymic nude mice treated with or without G-1. The xenograft volume was measured weekly. Student's t test was used to compare the tumor volumes between control and G-1–treated groups. Each point represents mean ± SEM (n = 6); *, P < 0.05. F, The weight of tumor xenografts from nude mice treated with or without G-1 for 2 weeks. Each bar represents mean ± SEM (n = 6). **, P < 0.01 compared with control. G, Representative images showing expression of Ki67 in control and G-1–treated tumor xenografts determined by fluorescent IHC. Ki67 was labeled in green; actin was stained with rhodamine-phalloidin (red); nuclei were stained with DAPI (blue); scale bar, 20 μm. The number of Ki67-positive cells and total cells in control and G-1–treated groups were also counted under a fluorescent microscope, and the ratio of Ki67-positive cells was calculated. Each bar represents mean ± SEM (n = 6), ***, P < 0.001. H, The body weight of athymic nude mice with or without G-1 treatment. Each bar represents mean ± SEM (n = 6). Student t test was used to do statistical analysis. No statistical significance on the body weight was observed between control and G-1–treated groups.

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MDA-MB-231 cells are TNBC (ERα, PgR, and HER2) cells with very low or no GPER protein expression (33). Interestingly, these cells were very sensitive to G-1 treatment. MDA-MB-231 cells started to detach from the culture plate 2 to 4 hours after G-1 addition and the cell number counts significantly decreased within 6 hours (Fig. 1A and B). Nanomolar concentrations of G-1 (100–500 nmol/L) were able to significantly suppress growth of MDA-MB-231 cells (Fig. 1A, Supplementary Fig. S1B and S1C). Soft agar colony formation assays showed that G-1 suppressed anchorage-independent growth of breast cancer cells. Treatment of MDA-MB-231 cells with 2 μmol/L G-1 nearly abolished colony formation and cell growth in soft agar. MDA-MB-231 cells treated with 10 μmol/L G-1 did not form any colonies (Fig. 1C). We used three additional cell lines to confirm the suppressive effects of G-1 on TNBC cell growth. MTT results showed that micromolar G-1 consistently suppressed the proliferation of BT-549, Hs 578T, and HCC1937 TNBC cell lines in both FBS-supplemented and FBS-free media (Supplementary Fig. S2). Consistent with our previous findings, G15 was not able to block the suppressive effects of G-1 in these cells (Supplementary Fig. S2). Interestingly, we found that TNBC cell lines are very sensitive to G-1 treatment. We observed that 100 nmol/L G-1 was able to suppressed proliferation of BT-549, Hs 578T, and HCC1937 TNBC cell lines after treatment for 4 days (Supplementary Fig. S2).

G-1 inhibits growth of breast cancer cells in vivo

The MDA-MB-231 breast cancer xenograft mouse model was used to examine the effect of G-1 on the growth of breast cancer cells in vivo. Tumor-carrying athymic nude mice were injected with vehicle (control) or 5.0 mg/kg G-1 daily for 2 weeks. Compared with the vehicle treated control group, G-1 treatment significantly reduced tumor volume and tumor weight (Fig. 1D–F). IHC analysis indicated that G-1 treatment significantly reduced the percentage of Ki-67–positive cells in breast cancer tissue, suggesting that G-1 is able to suppress the proliferation of breast cancer cells in vivo (Fig. 1G). Importantly, compared with the control groups, G-1 treatment for 2 weeks had no significant effect on mouse body weight, indicating that G-1 was well tolerated in intact nude mice (Fig. 1H).

G-1 induces caspase-dependent apoptosis of breast cancer cells

G-1 treatment resulted in the detachment of breast cancer cells from culture plates, suggesting that G-1 may induce breast cancer cell apoptosis. Annexin V-FITC/propidium iodide (PI) double staining showed that G-1 (2 μmol/L, 72 hours) significantly increased the number of Annexin V–positive cells (apoptotic cells) in all three breast cancer cell lines, regardless of their expression status of ERs (Supplementary Fig. S3A). Caspase assays showed that G-1 treatment also significantly increased caspase-3/7 activities in all three breast cancer cell lines (Supplementary Fig. S3B). The Western blot analyses further demonstrated that treatment of breast cancer cell lines with 1 μmol/L G-1 for 24 hours induced cleavage of caspases 3 and 7 (please note that MCF7 cells do not express caspase-3; ref. 34), and PARP, all of which are markers for apoptosis (Supplementary Fig. S3C). Together, these results indicated that G-1 is able to induce caspase activation and cell apoptosis in breast cancer cells, regardless their expression status of classic and membrane ERs.

G-1 blocks cell cycle in the prophase of mitosis in breast cancer cells

Treatment of breast cancer cells with G-1 for 24 hours decreased the percentage of cells in G1 phase, but increased the portion of cells in G2–M phase of the cell cycle in all three breast cancer cell lines (Supplementary Fig. S4). Flow cytometry also showed that G-1 treatment induced a significant increase in the portion of cells in the sub-G1 peak in all three breast cancer cell lines (Supplementary Fig. S4), supporting the observation that G-1 induced apoptosis in breast cancer cells. Importantly, these results indicate that G-1 arrests breast cancer cells in the G2–M phase and induces apoptosis of breast cancer cells in an ER-independent manner.

Active mitosis-promoting factors (MPF), consisting of CDC2 and cyclin B, drive the G2–M transition during cell division. Western blot analyses showed that G-1 treatments decreased phosphorylation of CDC2 at Y15 (inactive form), suppressed phosphorylation of CDC25C at S216 (inactive form), but increased expression of cyclin B and promoted phosphorylation of CDC25C (T48, active form, activate MPFs) in all three examined breast cancer cell lines (Supplementary Fig. S5). These data strongly suggest that G-1 treatment does not block G2–M transition. Taken together, these results suggest that G-1 arrested cell cycle in the mitosis, but not G2 phase.

The phosphorylation of histone H3 at serine 10 (S10) is considered to be closely related to chromosome condensation and is present during mitosis. Western blot analyses showed that phosphorylation of histone H3 (S10) was increased in all three breast cancer cell lines after G-1 treatments (Supplementary Fig. S5). Fluorescent ICC analyses showed that in MCF7 and SK-BR-3 cells, G-1 significantly increased the number of cells in mitosis [phosphorylated histone H3 (S10)-positive cells] within 16 hours (Supplementary Fig. S6A and S6B). Interestingly, in MDA-MB-231 cells, which had very low or no GPER expression, G-1 significantly increased the portion of mitotic cells within 8 hours (Supplementary Fig. S6A and S6B). These results further confirm the notion that G-1 is able to arrest cell cycle in the prophase of mitosis in a GPER-independent manner.

G-1 targets tubulin to interrupt mitotic spindle formation in breast cancer cells

Laser scanning confocal microscopy and fluorescent immunocytochemistry were used to further explore the mechanism by which G-1 blocks the cell cycle of breast cancer cells. Compared with control cells that went through all stages of mitotic phase (prophase, metaphase, anaphase, telophase, and cytokinesis), G-1–treated cells only entered the prophase of mitosis, which was characterized by phosphorylation of histone H3 on serine 10, chromosome condensation, and the formation of microtubule asters (Fig. 2A–C). However, normal spindles were never formed (Fig. 2A and B), suggesting that microtubule assembly was disrupted by G-1 treatment. Those cells arrested in prophase of mitosis eventually formed apoptotic bodies (Fig. 2B). These results suggest that G-1 is a potential novel microtubule-targeting agent.

Figure 2.

G-1 arrests cell-cycle progression of breast cancer cells in the prophase of mitosis. A, Representative images showing morphology of MDA-MB-231 cells incubated for 8 hours in the absence (CTRL) or presence of G-1 (2 μmol/L) for 8 hours. Microtubules were stained with β-tubulin antibody (red), and the nuclei were stained with DAPI (blue). Phosphorylated histone H3 (S10) was used as cell-cycle progression marker (green). Cells in different stages of cell cycle were indicated by white arrows; scale bar, 20 μm. B, Representative high-resolution confocal microscopy images showing detailed morphology of microtubules of MDA-MB-231 breast cancer cells incubated in the absence (top) or presence (bottom) of G-1 (2 μmol/L) for 8 hours; scale bar, 20 μm. Microtubules were stained with β-tubulin antibody (red), and the nuclei were stained with DAPI (blue). Phosphorylated histone H3 (S10) was used as cell-cycle progression marker (green). I, interphase; P, prophase; M, metaphase; A, anaphase; T&C, telophase and cytokinesis. C, A representative high-resolution image showing the microtubule asters formed in MDA-MB-231 cells after G-1 treatment for 8 hours. Microtubules were stained with β-tubulin antibody (red), and the nuclei were stained with DAPI (blue). Phosphorylated histone H3 (S10) was used as cell-cycle progression marker (green). White arrowheads point to spindle-like microtubule asters; scale bar, 5 μm.

Figure 2.

G-1 arrests cell-cycle progression of breast cancer cells in the prophase of mitosis. A, Representative images showing morphology of MDA-MB-231 cells incubated for 8 hours in the absence (CTRL) or presence of G-1 (2 μmol/L) for 8 hours. Microtubules were stained with β-tubulin antibody (red), and the nuclei were stained with DAPI (blue). Phosphorylated histone H3 (S10) was used as cell-cycle progression marker (green). Cells in different stages of cell cycle were indicated by white arrows; scale bar, 20 μm. B, Representative high-resolution confocal microscopy images showing detailed morphology of microtubules of MDA-MB-231 breast cancer cells incubated in the absence (top) or presence (bottom) of G-1 (2 μmol/L) for 8 hours; scale bar, 20 μm. Microtubules were stained with β-tubulin antibody (red), and the nuclei were stained with DAPI (blue). Phosphorylated histone H3 (S10) was used as cell-cycle progression marker (green). I, interphase; P, prophase; M, metaphase; A, anaphase; T&C, telophase and cytokinesis. C, A representative high-resolution image showing the microtubule asters formed in MDA-MB-231 cells after G-1 treatment for 8 hours. Microtubules were stained with β-tubulin antibody (red), and the nuclei were stained with DAPI (blue). Phosphorylated histone H3 (S10) was used as cell-cycle progression marker (green). White arrowheads point to spindle-like microtubule asters; scale bar, 5 μm.

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We then compared the actions of G-1 and known microtubule-targeting agents on the growth of breast cancer cells. We found that G-1 induced detachment of MDA-MB-231 and MCF-7 cells from the culture plates and increased the phospho-Histone H3–positive cells in a way similar to those known microtubule-targeting agents such as paclitaxel, 2-methoxyestradiol (2-ME), colchicine, and vinblastine (Supplementary Fig. S7), further suggesting that G-1 is a novel microtubule–targeting agent.

G-1 binds to the colchicine-binding site of tubulin to interrupt microtubule assembly

Paclitaxel interrupts tubulin dynamics by binding to the inner surface of microtubules to promote microtubule lengthening, whereas colchicine, 2-ME, and nocodazole bind to tubulin dimers to inhibit tubulin polymerization (35). Treatment of MDA-MB-231 and MCF-7 cells with paclitaxel induced massive assembly of microtubule bundles, whereas treatment of these cells with nocodazole induced disappearance of microtubule filaments (Fig. 3). Similar to nocodazole, G-1 treatment also induced the disruption of microtubule network in both MDA-MB-231 and MCF7 cells (Fig. 3), suggesting that G-1 is a suppressor of microtubule assembly. It seems that G-1 disrupts microtubule assembly in a concentration-dependent manner. Low concentrations of G-1 (<100 nmol/L) had no obvious effect on microtubule structure based on our microscopic analysis. However, treatment of MDA-MB-231 cells with 1 μmol/L G-1 for 2 hours partially depolymerized microtubules. Higher concentrations of G-1 (10 μmol/L) disrupted almost all microtubules in cells within 2 hours (Supplementary Fig. S8). Consistent with this observation in live cells, using in vitro microtubule assembly assays, we found that paclitaxel was able to promote the assembly of microtubules, whereas G-1 greatly suppressed microtubule assembly (Fig. 4A). In fact, addition of G-1 almost completely blocked the assembly of microtubules (Fig. 4A). We also performed in vitro microtubule sedimentation assays to explore the effects of G-1 on microtubule dynamics. We found that addition of G-1 in the reaction system decreased the amount of microtubules in the pellet while increased the amount of tubulin in the supernatant when compared with the control group (Fig. 4B). Paclitaxel was used as a tubulin polymerization–positive control. Paclitaxel elevated microtubules in the precipitate and reduced tubulin in the supernatant, indicating that the system worked well (Fig. 4B). Together, these results indicate that G-1, similar to 2-ME and colchicine, disrupts microtubule assembly.

Figure 3.

G-1 disrupts microtubule assembly in vivo. Representative high-resolution images showing microtubule structures (red) of MDA-MB-231 and MCF7 cells incubated in the presence or absence of vehicle (0.1% DMSO), paclitaxel (10 μmol/L), nocodazole (10 μmol/L), or G-1 (10 μmol/L) for 2 hours. Representative high-resolution image (focusing on a single cell) in each group was presented in the middle to show the detail of the microtubule structure. The low magnification image with multiple cells in each group was also presented (on the according side); Scale bar, 10 μm. Paclitaxel stabilizes microtubule, whereas G-1, similar to nocodazole, disrupts microtubule formation.

Figure 3.

G-1 disrupts microtubule assembly in vivo. Representative high-resolution images showing microtubule structures (red) of MDA-MB-231 and MCF7 cells incubated in the presence or absence of vehicle (0.1% DMSO), paclitaxel (10 μmol/L), nocodazole (10 μmol/L), or G-1 (10 μmol/L) for 2 hours. Representative high-resolution image (focusing on a single cell) in each group was presented in the middle to show the detail of the microtubule structure. The low magnification image with multiple cells in each group was also presented (on the according side); Scale bar, 10 μm. Paclitaxel stabilizes microtubule, whereas G-1, similar to nocodazole, disrupts microtubule formation.

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Figure 4.

G-1 disrupts microtubule assembly in vitro. A, High-resolution laser scanning confocal images showing effect of G-1 on microtubule assembly in vitro. Paclitaxel was used as a positive control. Addition of DMSO (≤0.05%, CTRL) had no effect on microtubule assembly, which is indicated by appearance of many microtubules formed by polymerization of X-rhodamine–labeled tubulin in the in vitro reaction system. Addition of paclitaxel greatly increased microtubule assembly, whereas addition of G-1 to the reaction drastically disrupted microtubule assembly. B, Effects of G-1 on microtubule assembly evaluated through a microtubule sedimentation assay. The left showed the representative gels loaded with pellet or supernatant after incubation with paclitaxel, control (0.05% DMSO), or G-1. The graphs show quantitative results. Each bar represents mean ± SEM (n = 3). *, P < 0.05; **, P < 0.01; ***, P < 0.001 compared with control. Taxol: with 20 μmol/L paclitaxel; CTRL: with DMSO vehicle treatment; G-1: with G-1 treatment.

Figure 4.

G-1 disrupts microtubule assembly in vitro. A, High-resolution laser scanning confocal images showing effect of G-1 on microtubule assembly in vitro. Paclitaxel was used as a positive control. Addition of DMSO (≤0.05%, CTRL) had no effect on microtubule assembly, which is indicated by appearance of many microtubules formed by polymerization of X-rhodamine–labeled tubulin in the in vitro reaction system. Addition of paclitaxel greatly increased microtubule assembly, whereas addition of G-1 to the reaction drastically disrupted microtubule assembly. B, Effects of G-1 on microtubule assembly evaluated through a microtubule sedimentation assay. The left showed the representative gels loaded with pellet or supernatant after incubation with paclitaxel, control (0.05% DMSO), or G-1. The graphs show quantitative results. Each bar represents mean ± SEM (n = 3). *, P < 0.05; **, P < 0.01; ***, P < 0.001 compared with control. Taxol: with 20 μmol/L paclitaxel; CTRL: with DMSO vehicle treatment; G-1: with G-1 treatment.

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We next used live-cell imaging and Hela cells expressing GFP-labeled tubulin to confirm our inference that G-1 is able to interrupt the dynamics of microtubules in cancer cell to suppress microtubule assembly. Live-cell imaging revealed that microtubules were highly dynamic and frequently grew and shrank at a rapid yet constant rate in the DMSO vehicle–treated control cells (Fig. 5A). However, in the G-1–treated Hela cells, the number of microtubules was significantly reduced. The remaining curved microtubules finally depolymerized and became invisible under microscope (Fig. 5A). Using the high-resolution laser scanning confocal microscopy and a live-cell imaging system, we further observed that in the dividing cells, G-1 treatment interrupted the dynamics of microtubules, leading to the disruption of microtubule assembly and subsequent failure of mitotic spindle formation (Fig. 5B). The elongated prophase of mitosis finally induced cell apoptosis, which was evidenced by the appearance of apoptotic bodies sooner after they failed to form normal mitotic spindles (Fig. 5B).

Figure 5.

Screenshots of cell division videos support the notion that G-1 targets microtubules to suppress cell growth. Hela cells with GFP-labeled α-tubulin were incubated in the culture chamber. Cell division and microtubule dynamics were monitored with a high-resolution laser scanning microscope and a live cell imaging system. A, Screenshots from the time-lapse video showing the dynamics of microtubules in cells treated with vehicle (0.1% DMSO, top) or G-1 (bottom). White arrows point to the intact and depolymerized microtubules. B, Representative screenshots from time-lapse videos showing the microtubule dynamics and morphology cells with or without G-1 treatment. Red arrows indicate the time point when DMSO or G-1 was added to the culture system. This video clearly shows that G-1 treatment interrupts microtubule assembly, leading to the failure of spindle formation and subsequent activation of cell death pathways, which is indicated by the formation of apoptotic bodies.

Figure 5.

Screenshots of cell division videos support the notion that G-1 targets microtubules to suppress cell growth. Hela cells with GFP-labeled α-tubulin were incubated in the culture chamber. Cell division and microtubule dynamics were monitored with a high-resolution laser scanning microscope and a live cell imaging system. A, Screenshots from the time-lapse video showing the dynamics of microtubules in cells treated with vehicle (0.1% DMSO, top) or G-1 (bottom). White arrows point to the intact and depolymerized microtubules. B, Representative screenshots from time-lapse videos showing the microtubule dynamics and morphology cells with or without G-1 treatment. Red arrows indicate the time point when DMSO or G-1 was added to the culture system. This video clearly shows that G-1 treatment interrupts microtubule assembly, leading to the failure of spindle formation and subsequent activation of cell death pathways, which is indicated by the formation of apoptotic bodies.

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Because the action of G-1 is similar to 2-ME and nocodazole, which bind to colchicine binding sites of β-tubulin to exert their tumor-suppressing action (35, 36), we used a binding competition assay to examine whether G-1 also exerts its role by binding to the colchicine-binding sites of β-tubulin to interrupt microtubule assembly. As expected, vinblastine did not compete for the H3-colchicine–binding site (Fig. 6). However, G-1, similar to colchicine and 2-ME, effectively competed for the binding of H3-colchicine to β-tubulin in a concentration-dependent manner (Fig. 6). These results indicate that G-1 binds to colchicine-binding site in β-tubulin to block tubulin polymerization and microtubule assembly in breast cancer cells.

Figure 6.

G-1 binds to the colchicine-binding site in β-tubulin. The ability of G-1 to bind to the colchicine-binding site in tubulin was evaluated with a colchicine site competitive assay. Incubation with colchicine, G-1, fluorescein-labeled G-1, and 2-ME–suppressed H3-Colchicine binding in a concentration dependent manner. Incubation with vinblastine, which was used as a negative control, had no competition on H3-Colchicine binding to β-tubulin. Each point represents mean ± SEM (n = 4). The competition analysis was repeated for three times. FG-1: fluorescein-labelled G-1. 2-ME; 2-methoxyestradiol.

Figure 6.

G-1 binds to the colchicine-binding site in β-tubulin. The ability of G-1 to bind to the colchicine-binding site in tubulin was evaluated with a colchicine site competitive assay. Incubation with colchicine, G-1, fluorescein-labeled G-1, and 2-ME–suppressed H3-Colchicine binding in a concentration dependent manner. Incubation with vinblastine, which was used as a negative control, had no competition on H3-Colchicine binding to β-tubulin. Each point represents mean ± SEM (n = 4). The competition analysis was repeated for three times. FG-1: fluorescein-labelled G-1. 2-ME; 2-methoxyestradiol.

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It is well known that prolonged exposure to estrogens represents a major risk factor for the progression of breast cancer (17). ERα antagonists like tamoxifen and raloxifene are currently used as first-line pharmacologic interventions in ERα-positive breast cancer to inhibit the mitogenic stimulation of estrogens (37). However, more than 20% of ERα-positive patients do not respond to treatment, and in most patients tamoxifen produces agonist effects after treatment for a few years (38). Recent studies indicated that tamoxifen and fulvestrant activate GPER, which has been shown to interact with the EGFR signaling pathway to drive proliferative genes and pathways to promote breast cancer cell growth (2, 39). Consistently, several studies did show that activation of GPER by estrogen or tamoxifen promotes breast cancer cell growth (40–43). Developed as a high-affinity nonsteroidal GPER agonist (3), G-1 is expected to specifically activate GPER to stimulate the growth of breast cancer cells. In favor of this speculation, Magiollini and colleagues (6, 18) showed that G-1 stimulates proliferation of breast cancer cells through the GPER/EGFR/ERK signal transduction pathways. Moreover, Scaling and colleagues (19) reported that G-1 increases the mitotic index in breast MCF10A cells and promotes proliferation of cells in human normal breast and breast cancer explants. More recent observations, however, demonstrated that G-1 arrests the cell cycle at the G2–M phase, downregulates cyclin B, and induces mitochondrial-related apoptosis in SK-BR-3 and MDA-MB-231 cells (23, 44, 45). The disparate effects of G-1 on the growth of breast cancer cells were also observed in tumor xenograft mouse models. Maggiolini and colleagues indicated that G-1 (0.5 mg/kg/d) promotes the growth of SK-BR-3 breast cancer cell tumor xenografts (18, 46). On the contrary, other investigators showed that G-1 treatment suppressed the growth of SK-BR-3 xenograft tumors (23). A very common impression from previous reports is that low concentrations of G-1 stimulate cell proliferation, while high concentrations of G-1 suppress cell growth. In the current study, we found that although very low concentrations of G-1 (1–10 nmol/L) had no obvious effects on the growth of six breast cancer cell lines, relatively low concentrations of G-1 (100 nmol/L) significantly suppressed the growth of examined TNBC cancer cell lines. Micromolar levels of G-1 consistently resulted in the inhibition of breast cancer cell growth as evidenced by multiple parameters: (i) detachment of breast cancer cells from culture plates within hours, (ii) suppression of cancer cell growth in traditional 2-dimensional culture systems and in anchorage-independent culture systems, and (iii) activation of caspases and induction of cell death. Importantly, we found that injection of G-1 (5 mg/kg/d) inhibited growth of tumors derived from MDA-MB-231 cells in xenograft mouse models, suggesting that G-1 is able to exert its tumor-suppressive action under physiologic conditions. Although the circulating levels of G-1 were not determined in our study, previous reports (15) and our in vivo results indicates that a dosage of approximately 5 mg/kg/d is sufficient to achieve the drug concentrations required to suppress cancer cell growth in mice. Importantly, our in vivo study demonstrated that G-1 treatment has no effect on animal body weight during treatment. Our more recent data further demonstrated that G-1 treatment suppressed breast and ovarian tumor progression in xenograft mouse models, but had no effect on ovarian physiology (indicated by normal ovulation and corpus luteum formation in the ovary, Lv and colleagues, unpublished observations). These in vivo studies indicate that G-1 is well tolerated in the body. Our results indicate that G-1 is a promising candidate drug for breast cancers, especially for TNBC.

The molecular mechanism by which G-1 regulates cancer cell proliferation is unclear. Previous research showed that activation of GPER by G-1 in breast cancer cells triggers the activation of the EGFR signaling pathway (18–19, 23). Other groups showed that G-1 suppresses cancer cell proliferation via GPER-mediated activation of P53 and P21 signaling pathways (20). It is worth noting that previous studies speculated that GPER is the mediator of G-1 actions, on both growth-inhibiting and growth-promoting processes (5–8, 14, 15, 19–22, 24). However, no any direct evidence was provided in those studies to indicate the activation of GPER and the subsequent activation of G-proteins. Our previous studies in ovarian cancer cells showed not only that G-1 does not stimulate cell proliferation but that it suppresses cell proliferation in a GPER-independent manner (9, 10). We found that knockdown of GPER did not affect the inhibitory action of G-1 on the proliferation of ovarian cancer cells. Moreover, blocking the function of GPER using the GPER-selective antagonist G-15 (31) did not affect the ability of G-1 to suppress ovarian tumor cell proliferation (9, 10). Most importantly, we found that G-1 also suppressed proliferation of HEK293 cells (9, 10), which lack GPER (47). In support of our observations in ovarian cancer cells, we found that in the TNBC cell line MDA-MB-231, which has no (or very low if any) GPER expression, G-1 inhibited cell growth with a concentration as low as 100 nmol/L. Moreover, micromolar levels of G-1 consistently suppressed proliferation and induced apoptosis in MCF-7, SK-BR-3, and four TNBC cell lines, which have different expression status of ERs. In addition, the putative GPER antagonist G-15 did not block the growth-suppressive effects of G-1 in breast cancer cells. Our data clearly indicate that the action of G-1 in breast cancer cells has no direct association with the expression of classic and nonclassic ERs. G-1 is able to suppress breast cancer cell proliferation in a GPER-independent manner.

How G-1 induces breast cancer cell death is unclear. Employing the high-resolution laser scanning confocal microcopy and time-lapse imaging techniques, our present studies revealed the dynamic process of G-1–induced cell death. G-1 had no effect on quiescent cells (such as luteinized granulosa cells, data not shown). However, once cells start to divide, G-1 treatment leads to the formation of many spindle-like microtubule asters in the dividing cells within several hours, whereas the normal mitotic spindles fail to form. The elongated prophase during cell division triggers the apoptotic signaling pathway, leading to the formation of apoptotic bodies and eventually cell death. Biochemical and immunocytochemical studies showed that G-1 does not affect the entry of mitosis. Flow cytometry and apoptotic analysis indicated that the caspase signaling pathway is involved in G-1–induced breast cancer cell death.

By using high-resolution laser scanning confocal microscopy and time-lapse microscopy, we provide direct evidence to show that G-1 suppresses breast cancer cell proliferation and induces cancer cell apoptosis by disrupting microtubule assembly. These results are consistent with our observations in ovarian cancer cells (10). Our data from both ovarian and breast cancer cells indicate that G-1 is a novel microtubule-targeting agent (MTA). MTAs are classified into two groups: microtubule-stabilizing agents such as paclitaxel, which exert their function by promoting tubulin polymerization and increasing microtubule assembly in cells, and microtubule-destabilizing agents such as vinca alkaloids, which inhibit tubulin polymerization and depolymerize microtubules to interrupt microtubule dynamics. Our results from high-resolution laser scanning microscopy showed that treatment of the breast cancer cells with G-1 induced significant reduction of tubulin filaments, indicating that G-1 is a microtubule-destabilizing agent. Supporting this observation, G-1 abrogated formation of microtubules in both the tubulin polymerization assay and the microtubule sedimentation assay.

MTAs are known to interact with tubulin through at least 4 binding sites: the laulinmilide, taxane/epthilone, vinca alkaloid, and colchicine sites (36). The colchicine binding competing analysis showed that G-1, similar to 2-ME (36), competes with colchicine for tubulin binding, indicating that G-1 binds to the colchicine binding sites in the breast cancer cells to interrupt the tubulin polymerization, microtubule assembly and the subsequent spindle formation, leading to the arrest of breast cancer cells in the prophase of mitosis. Colchicine binding to β-tubulin results in curved tubulin dimer. This dimer prevents it from adopting a straight structure, due to a steric clash between colchicine and α-tubulin, which inhibits microtubule assembly (48). It is possible that G-1 may bind to the tubulin monomer or dimer to block the tubulin filament elongation and microtubule assembly. Further studies are necessary to provide deeper understanding on the interaction between G-1 and tubulins.

Our findings in the present study have significant clinical relevance. The biological responses in the in vitro studies and the tumor-suppressing mechanism of G-1 are similar to 2-ME. As a candidate drug that targets the colchicine-binding site, 2-ME failed to reach sufficient plasma levels to evaluate its therapeutic potential in phase I and II clinical trials (49). Results from our xenograft mouse models indicate that a dose of G-1 (5 mg/kg/d) was well-tolerated and able to achieve sufficient levels to suppress breast tumor growth. Moreover, 2-ME easily lost its antitumor activity in vivo due to oxidation at position 17 and conjugation between positions 17 and 3 (50). G-1 has a more stable chemical structure at similar positions compared with 2-ME (3). These unique features suggested that G-1 is an “improved version of 2-ME,” ensuring the potential application of this drug to further preclinical and clinical studies.

In conclusion, our results provide direct evidence that G-1, the putative GPER agonist, is a novel microtubule-targeting agent that binds to colchicine-binding site of tubulin protein to disrupt microtubule dynamics during mitosis, leading to the apoptosis of breast cancer cells and suppression of tumor growth. Importantly, previous physiologic and pharmacologic studies have shown that pharmacologic doses of G-1 played favorable roles in circulatory, immune, and nervous system in mice (4). Our in vivo studies showed that injection of mice with G-1 suppressed growth of breast and ovarian tumor cells, but had no effects on mouse body weight, social behavior, and function of reproductive physiology in a xenograft mouse model (Lv and colleagues; unpublished data). The low toxicity and the potent inhibitory effect on growth of breast cancer cells, especially on the growth of TNBC cells in vitro and in vivo, indicate that G-1 is a promising chemotherapeutic drug for better treatment of breast cancers.

No potential conflicts of interest were disclosed.

Conception and design: X. Lv, C. Wang

Development of methodology: X. Lv, C. Wang

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): X. Lv, C. He, C. Huang, G. Hua, J. Dong, C. Wang

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): X. Lv, C. He, Z. Wang, J.S. Davis, C. Wang

Writing, review, and/or revision of the manuscript: X. Lv, S.W. Remmenga, K.J. Rodabough, A.R. Karpf, J. Dong, J.S. Davis, C. Wang

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): X. Lv, C. Wang

Study supervision: C. Wang

We thank Janice A. Taylor and James R. Talaska of the Advanced Microscopy Core Facility at the University of Nebraska Medical Center for providing assistance with confocal microscopy.

This work was partially supported by the Eunice Kennedy Shriver National Institute of Child Health and Human Development (5R00HD059985; to C. Wang); The National Cancer Institute (R01CA197976; to C. Wang); The Olson Center for Women's Health (to C. Wang); The Fred & Pamela Buffett Cancer Center (LB595; to C. Wang); The Colleen's Dream Foundation (to C. Wang), The Marsha Rivkin Center for Cancer Research (The Barbara Learned Bridge Funding Award; to C. Wang), the COBRE Grant from the Nebraska Center for Cell Signaling/NIGMS (5P30GM106397-02; to K.R. Johnson and C. Wang), and Department of Veterans Affairs Research and Development Program (to J.S. Davis).

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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