Inhibition of Copper Transport Induces Apoptosis in Triple-Negative Breast Cancer Cells and Suppresses Tumor Angiogenesis

This article is featured in Highlights of This Issue, p. 871

Breast cancer is responsible for more than 250,000 new cases and 40,000 deaths among women, yearly, despite continuous efforts to improve treatment (1). Breast cancer is a biologically complex disease in its histology, molecular classification, response to therapy and mortality rates (2, 3). Five intrinsic molecular subtypes (luminal A, luminal B, HER2-enriched, claudin-low, and basal-like) have been identified using comprehensive gene expression analysis of human breast cancer tissue, cell lines and mouse models (2, 3). Triple-negative breast cancer (TNBC) lacks estrogen and progesterone receptors, the HER2, and is often associated with basal-like subtype representing a complex, clinically aggressive form of the disease (4). Although 30% to 40% of patients with early-stage TNBC benefit from treatment with anthracycline and taxane-based chemotherapy, TNBC is difficult to control if it becomes resistant to treatment and spreads to distant organ sites (5–7). No specific chemotherapy agents are able to cease metastatic spread, and most patients with TNBC die from advanced disease within 20 months post progression (8). A lack of molecular targets, the adaptive behavior of cancer cells and the microenvironment contributing to tumor progression are barriers to successful therapy. To improve patient outcomes, novel treatment approaches targeting intracellular pathways and pathways involved in cross-talk between cancer cells and the tumor microenvironment are critical.

Copper and copper-dependent proteins are emerging therapeutic targets due to their involvement in cell proliferation, survival, angiogenesis, and metastasis (9, 10). Elevated levels of copper in blood and tumor tissue of patients with cancer are correlated with disease progression (11). Copper-dependent superoxide dismutase (SOD1) is an important modulator of oxidative stress in cancer cells (12, 13), and the lysyl oxidase family of proteins require copper to stabilize the extracellular matrix, which contributes to the formation of a pre-metastatic niche (14, 15). Cells maintain a network of proteins (CTR1, ATP7A, ATP7B) that shuttle copper across membranes to regulate copper homeostasis to prevent damage from the redox-active metal, and deliver copper to secreted copper-dependent proteins (16). Intracellular transport of copper is mediated by chaperone proteins, ATOX1 and CCS, that supply copper to the copper-dependent ATP7A and ATP7B (Cu-ATPases), and to SOD1, respectively (17). CCS is also required for copper-mediated activation of HIF-1α to promote VEGF expression (18). ATOX1 has a possible role in metastasis: It promotes inflammatory neovascularization (19), wound closure (20), and breast cancer cell migration (21). Altogether, this evidence demonstrates that copper chaperone proteins, ATOX1 and CCS, are attractive targets for anti-cancer therapy.

Current approaches to targeting copper homeostasis include the inhibition of copper-dependent enzymes by global copper chelation, and the use of copper ionophores to elevate or redistribute copper and overwhelm the antioxidant capacity of cancer cells (22). Recently, the chelator tetrathiomolybdate (TM) was evaluated in a Phase II clinical trial for patients with breast cancer with high risk of recurrence. TM suppresses angiogenesis in preclinical models (23), with its activity ascribed to the inhibition of SOD1 (13, 24), LOX (25) and NF-kβ (23). TM treatment reduced serum copper levels and improved survival of patients with breast cancer, including patients with TNBC, demonstrating proof of principle for this approach (26). In a preclinical model, copper depletion with TM did not affect tumor growth, although it did reduce lung metastases. Despite the promising results of this trial, TM is insufficient to reduce the tumor burden in highly vascularized tumors, and may not inhibit tumor progression once an angiogenic switch has occurred (27).

An alternative approach to targeting cellular copper homeostasis is the inhibition of copper chaperones to impede copper transfer to copper-dependent enzymes and disrupt copper homeostasis. DCAC50 is a small molecule designed to inhibit intracellular copper transport by blocking the highly similar copper transfer interfaces of both ATOX1 and CCS, which prevents the protein–protein interactions necessary for copper transfer to the Cu-ATPases and SOD1, respectively (28). In leukemia and lung cancer cell lines DCAC50 reduced cell proliferation and tumor growth in xenograft mouse models without significant side effects (28).

Given the promising results of DCAC50 anticancer activity and the benefit of copper depletion in patients with breast cancer, we studied the impact of the inhibition of copper chaperones in a broad panel of cell lines of the most aggressive breast cancer phenotype, TNBC. We describe the effects of DCAC50 on the proliferation, viability, copper homeostasis, and redox status of TNBC cells. We also investigate the ability of DCAC50 to inhibit tumor growth and angiogenesis in breast cancer xenograft mouse models, and test its activity in combination with paclitaxel. This work illustrates the potential of copper transporters as targets for breast cancer treatment, shows the impact of the intracellular inhibition of copper homeostasis in TNBC, and investigates the mechanism of action.

DCAC50 was synthesized as previously reported (28) and was characterized by NMR and UV/Vis spectroscopy. Binding to ATOX1 was confirmed by fluorescence assay (28). DCAC50, ammonium tetrathiomolybdate (TM, 99.97%) and paclitaxel (Sigma) stocks were prepared in DMSO. L-buthionine-sulfoximine (BSO, 97%, Sigma) was prepared in water.

Breast cancer cell lines were obtained from the ATCC and were cultured in RPMI-1640 medium supplemented with 10% FBS and antibiotic–antimycotic solution (Gibco). SUM149 cells were a gift from Dr. Perou (UNC, Chapel Hill) and were cultured in HuMEC with 5% FBS (Gibco). HMEC and HUVEC, were purchased from Lonza and cultured in MEGM or EGM-2 medium (Lonza). Cell lines were grown at 37°C and 5% carbon dioxide for less than 20 passages after thawing to conduct described experiments, tested negative for Mycoplasma contamination and validated for species and unique DNA profile by the provider using short tandem repeat analysis. All assays were performed according to the manufacturer's instructions and after allowing cells to adhere overnight.

Cell lysates were collected in RIPA buffer (Sigma), sonicated (3 × 10 seconds) and centrifuged (10 minutes at 14,000 × rcf). Protein concentration was determined using Thermo Scientific PierceBCA Protein Assay. Equal amount of total protein for each lysate was analyzed by SDS-PAGE and transferred to polyvinylidene difluoride membrane (ImmobilonFL, Merk Millipore). The loading controls were from the same experimental samples. All proteins were analyzed on the same blot. The membrane was scanned using Odyssey IR Scanner. Images were analyzed using Image Studio Light (LI-COR).

About 5,000 to 10,000 cells per well were plated into a 96-well plate for each drug concentration (n = 3–6). DCAC50, TM, BSO, and paclitaxel solutions were prepared as serial dilutions with total DMSO matched to vehicle control. Constant ratios of DCAC50 and paclitaxel were used when treated in combination. Solutions (2X concentrates) were added to the cells in media for 72 hours at 37°C and CellTiter 96-AQueous One Solution Assay (Promega) was performed. Data were fitted into a variable slope (four-parameter) model using GraphPad Prism. For paclitaxel and DCAC50 combination, fraction of viable cells was used to calculate combination index (CI) and dose-reduction index (DRI) using CompuSyn software (29). Alternatively, cell proliferation was assessed with The IncuCyte S3 Live-Cell Analysis System (Essen Instruments) using quantitative metrics derived after phase-contrast image acquisition, and presented as the percentage of confluence.

Cells were plated and treated per the proliferation assay and assayed with Caspase-Glo-3/7 assay (Promega). For the AnnexinV and propidium iodide assay, the Dead Cell Apoptosis Kit (Invitrogen) was used. Cells were plated in 6-well plates, treated, collected by trypsinization, stained and analyzed with LSR-Fortessa 4–15 (Bechton and Dickenson) instrument and FlowJo software.

Cells were plated in 6-well plates, treated, trypsinized, washed with PBS, collected in acid-washed tubes, digested in trace metal grade concentrated nitric acid (overnight at 37°C, with shaking) and diluted to 2% nitric acid with ultrapure water. ⁶⁵Cu content was determined by inductively coupled plasma mass spectrometry (ICP-MS; Agilent 7700x) with a germanium internal standard.

A total of 10⁶ cells/well were seeded in 6-well plates containing a silicon nitride membrane (1.5 × 1.5 mm × 500 nm, Silson, UK), serum-starved for 24 hours and full-serum media were added one hour before treatment. Cells were washed with PBS, fixed with paraformaldehyde (3.7%, pH 7.4, 20 minutes, room temperature), washed with trace metal grade ammonium acetate (100 mmol/L), then ultrapure water, and air-dried.

XFM was conducted at beamline 2-ID-D at the Advanced Photon Source. Samples were irradiated with a 10.1 keV beam focused through a zone plate and order-sorting aperture. X-ray fluorescence maps were generated by “fly scanning” (1 second dwell, 0.5-μm steps). Fluorescent photons were collected by a Ge detector (UltraLEGe, Canberra) at 90° to the incident beam.

Analysis was performed using MAPS software (30) as described elsewhere (31). The nuclear region of the cell was defined as the region of high P and Zn content.

Cells were plated into 8-well glass chamber slides, coated with poly-L-lysine (Sigma), treated and fixed as for XFM microscopy, washed with PBS, permeabilized (0.1% Triton-X100; Sigma) and blocked with Protein Block Goat Serum (Biogenex Laboratories), stained with primary, then secondary antibodies. Cells were analyzed using Olympus IX-83 microscope controlled by Metamorph software, Olympus UPlanSAPO 40 × (oil, N.A. 1.25) objective, Xcite 120 LED (Lumen Dynamics) light source, Image EMX2 CCD (Hamamatsu) camera, and ImageJ software.

Cells were plated into 10-cm² dishes, treated, collected by trypsinization, stained with 10 μmol/L DCFDA reagent (30 minutes, 37°C) and washed with PBS. FACS analysis was performed using an LSR-Fortessa 4–15 (Becton and Dickenson) instrument and FlowJo software.

A total of 20,000 cells were seeded in 96-well plates. The luminescence-based GSH/GSSG-Glo Assay (Promega) was used to determine the GSH:GSSG ratio or to calculate total GSH levels.

Cells were seeded in 10-cm² dishes. For total SOD activity, treated cells were collected by trypsinization, washed with PBS. The cell pellet was lysed in lysis buffer (Trevigen; 30 minutes) and the supernatant was collected after centrifugation (10 minutes, 4°C, 10,000 × g). Total SOD activity was determined using the SOD activity assay (Sigma). For SOD1 activity, treated cells were washed, scraped into PBS and centrifuged. The pellet was suspended in three volumes of 50 mmol/L phosphate buffer (pH 7.6) and sonicated. Cell lysates were separated by native PAGE at 4°C. The gel was stained as in (32) with modifications (see Supplementary Methods).

HuVEC (20,000 cells/cm²) were plated into 48-well plates coated with phenol red-free Matrigel (Corning, 30 minutes, 37°C), in presence of DCAC50, TM or DMSO. Cell network-formation was imaged after 16 to 18 hours incubation at 37°C, using an Olympus IX81 inverted microscope with the Olympus Zero Drift Correction auto re-focusing system (Olympus) with a Hamamatsu Orca Flash 4.0 sCMOS camera (Hamamatsu Photonics) run by Slidebook 5.0 software (Intelligent Imaging Innovations), using a ×10 objective with adapter and Angiogenesis Analyzer for ImageJ (NIH; ref. 33).

Cells were seeded in 24-well plates and treated in phenol red-free media. Media was collected and centrifuged to remove debris. Equal volumes of sample and reagent from Amplite Fluorimetric Lysyl Oxidase Assay Kit (AAT Bioquest) were mixed. The reactions were incubated in the dark (37°C) and fluorescence (540 nm excitation/590 nm emission) was measured after 60 minutes.

All animals were humanely handled and monitored for health conditions according to the Institutional Animal Care and Use Committee (IACUC) approved protocols. Eight-week-old female Foxn1nu/nu (Harlan) mice were anesthetized via inhalation with 2% vaporized isoflurane and were unilaterally injected with 4 × 10⁶ MDA-MB-468 or 3 × 10⁶ MDA-MB-231 cells (100 μL, 50% Matrigel) into the fourth inguinal mammary gland at the base of the nipple. Seven days post cellular implantation, animals were randomly and blindly assigned to treatment groups (6–8 per group), and treated with intraperitoneal injections of 50 mg/kg/d DCAC50 or DMSO (vehicle control). Tumor growth and weight of the animals were monitored twice weekly. Tumor measurements were performed using calipers to calculate tumor volume using formula 1/2(Length × Width²). The assessment was blind to treatment groups and was performed by the same person throughout the study.

Frozen tumors were thawed in RPMI-1640 media containing 10 mg/mL BSA on ice, and prepared as described previously (34). Leica TCS SP8 confocal laser scanning microscope, white light laser, Leica HCX PL APO 10X/0.4 NA dry objective (2.2 mm working distance), and a SuperZ galvometric scanning stage were used for imaging.

Elevated copper levels in serum and tissues of patients with cancer (11) are a sign of altered copper homeostasis. Elevated mRNA levels of copper transport proteins have been observed in human cancers (28, 35). In breast cancer tissues ATOX1 mRNA levels were upregulated relative to normal breast tissue, but no significant change was observed for CCS (35). Because mRNA levels do not always correlate with protein abundance, we studied the protein expression of ATOX1 and CCS in a panel of breast cancer cell lines. We included luminal and HER2-positive subtypes, as well as basal-like and claudin-low TNBC subtypes, and compared protein levels with normal human mammary epithelial cells (HMEC). Protein expression levels of ATOX1 were elevated in all breast cancer cells lines whereas CCS expression levels were more variable (Fig. 1A and B; Supplementary Fig. S1). The highest levels of ATOX1 were observed in luminal MCF7 (5.4-fold), claudin-low, MDA-MB-157 (6.3-fold) and basal-like MDA-MB-468 (2.1-fold) cell lines. CCS protein levels were most elevated in luminal MCF7 (4.8-fold), claudin-low HCC1395 (4.1-fold) and basal-like HCC1806 (2.0-fold) cell lines. Levels of both proteins, ATOX1 and CCS, were elevated in luminal (MCF7, T47D, ZR75-1), HER-2-positive (ZR75-30), basal-like (HCC1937, HCC1806, MDA-MB-468) and claudin-low (MDA-MB-231, MDA-MB-436, HCC1395) cell lines. Because DCAC50 targets the activities of ATOX1 and CCS, we continued the study with representative cell lines of different subtypes showing variable elevated levels of these proteins.

The cytotoxicities of the intracellular copper transport inhibitor, DCAC50, and of the copper chelator, TM, previously shown to inhibit copper-dependent enzymes in breast cancer, were evaluated using a dose-response profile in breast cancer cells. To evaluate apoptosis, we measured the activity of caspase-3/7, and estimated percentages of apoptotic cells determined by simultaneous staining with propidium iodide and AnnexinV. Treatment with DCAC50 reduced cell proliferation in a dose-dependent manner in all studied cell lines. Calculated IC₅₀ doses varied from 5 to 10 μmol/L in breast cancer cells, and higher efficacy was observed in basal-like cell lines (Fig. 1C and D). Next, we evaluated the ability of DCAC50 to induce apoptosis in the most aggressive breast cancer subtype, TNBC. Inhibition with DCAC50 resulted in activated caspase-3/7, and significantly increased percentages of apoptotic cells in all cell lines (except MDA-MB-231), but especially in basal-like cell lines (Fig. 2A–C). Copper chelation with TM did not affect cell proliferation in studied cell lines (except for HCC1395, IC₅₀ ∼60 μmol/L, and HCC1806, IC₅₀ ∼90 μmol/L), and did not induce apoptosis (Fig. 2A–C; Supplementary Fig. S2A). In HMEC, no significant changes in caspase-3/7 activity were observed after treatment with DCAC50; however, DCAC50 did reduce cell proliferation (IC₅₀ = 3.5 ± 0.1 μmol/L, Supplementary Fig. S2B–S2C). Thus, treatment with DCAC50 attenuates cell proliferation and leads to apoptosis-induced cancer cell death.

Copper chaperone ATOX1, together with Cu-ATPases, are part of the copper export system that helps maintain copper homeostasis. Although targeting copper homeostasis using chelators, such as TM, is expected to reduce cellular copper levels, the inhibition of copper transfer from ATOX1 to Cu-ATPases with DCAC50 is expected to increase cellular copper content. ICP-MS analysis revealed that overall copper content in TM-treated cells is decreased, whereas copper content in DCAC50-treated claudin-low cells is significantly increased, compared with control cells (Fig. 3A; Supplementary Fig. S3A). Surprisingly, only a modest non-significant increase in copper levels was detected in basal-like cells, which are most sensitive to DCAC50. Baseline copper levels were lower in all basal-like cells (<9 pg Cu/μg protein) than in claudin-low cells (>12 pg Cu/μg protein), indicating that basal-like cells may be more sensitive to marginal copper overload.

The inhibition of copper chaperones and changes in cellular copper levels are likely to lead to changes in spatial copper distribution. To better understand the impact of DCAC50 on copper homeostasis in TNBC, XFM imaging was used to determine intracellular copper distribution (Fig. 3B). In DCAC50-treated MDA-MB-231 cells, where total intracellular copper level increased with treatment, there was no significant change in the nuclear to cytoplasmic copper concentration ratio compared with control (Fig. 3B). In HCC1395 cells there was a 20% increase in the nuclear to cytoplasmic copper ratio from 1.1 to 1.3 (Supplementary Fig. S3B). In DCAC50-treated MDA-MB-468 cells, despite limited copper accumulation observed by ICP-MS, the ratio of nuclear to cytoplasmic copper concentration compared with control increased 40% from 1.2 to 1.7 (Fig. 3B). Importantly, there is no corresponding change in the localization of iron or zinc across cell lines or treatments (Supplementary Fig. S3C and S3D), indicating that the copper redistribution is not an artifact of sample preparation nor is it indicative of differences in cell cycle across treatment groups. Thus, inhibition of copper transport with DCAC50 alters copper homeostasis resulting in increased copper levels and/or increased localization of copper to the nucleus. The greatest change in copper distribution with DCAC50 treatment was observed in basal-like MDA-MB-468 cells, which is the most sensitive of the three imaged cell lines to the drug.

Inhibition of ATOX1 and CCS by DCAC50 may in turn affect the level or distribution of copper transport proteins. However, the protein levels of ATP7B, ATOX1, CCS or CTR1 were unchanged after treatment with DCAC50 (Supplementary Fig. S4A). The copper transporter ATP7B is known to translocate between the perinuclear space and the plasma membrane to metallate secreted copper proteins and maintain appropriate cellular copper levels (16). DCAC50 is designed to disrupt copper transfer from ATOX1 to ATP7B (28), and therefore may prevent ATP7B from translocating to the plasma membrane where it facilitates copper export. To test this hypothesis, we performed immunofluorescence staining of ATP7B in MDA-MB-468 cells after treatment with DCAC50. Upon treatment, ATP7B was mostly found in the perinuclear region of MDA-MB-468 cells (Fig. 3C). In contrast, we observed diffuse staining of ATP7B in DMSO-treated cells. Thus, in basal-like MDA-MB-468 cells, DCAC50 disrupts copper homeostasis altering copper distribution and ATP7B localization.

To facilitate proliferation and promote survival cancer cells frequently have to adapt to high levels of oxidative stress. Copper is intimately linked with the cellular redox status via its inherent redox activity, SOD1 activity and interactions with GSH. Thus, increased copper levels and the inhibition of copper transfer to SOD1 with DCAC50 are expected to induce oxidative stress in TNBC cells ultimately leading to cellular damage. To determine the impact of DCAC50 on cellular redox status we used a cytosolic sensor of general oxidative stress, DCFDA. Treatment with DCAC50 resulted in higher DCF fluorescence intensity generated by the oxidation of DCFDA. The fluorescence signal was higher after 24 hours treatment with DCAC50 and by 72 hours had increased 2-fold compared with control cells (Fig. 4A and B). Copper chelation by TM had no impact on cellular oxidative stress as measured by DCF fluorescence, except for an increase observed in MDA-MB-231 cells.

Another indicator of the cellular redox environment is the oxidation state of glutathione. Lower ratios of reduced to oxidized glutathione (GSH:GSSG) were observed in all TNBC cells lines after treatment with DCAC50, indicating increased oxidative stress. Interestingly, total GSH levels at baseline were overall higher in claudin-low cells in comparison with basal-like cells, suggesting claudin-low cells possess higher antioxidant capacity and thus, may be less sensitive to the oxidative stress triggered by DCAC50 (Fig. 4C and D; Supplementary Fig. S4B). To test whether exhausting cellular antioxidant capacity by depleting GSH levels augments DCAC50′s effects, we evaluated cell proliferation in claudin-low MDA-MB-436 and HCC1395 cells after treatment with DCAC50 in presence of the GSH-synthesis inhibitor, BSO. GSH levels were reduced without affecting cell proliferation after treatment with BSO, and addition of BSO to DCAC50 resulted in significantly reduced cell proliferation compared with DCAC50 alone (Supplementary Fig. S4C and S4D). This finding further supports our hypothesis that lowering cellular antioxidant capacity increases susceptibility to DCAC50.

Copper-dependent SOD1 is a known regulator of cellular redox balance, and DCAC50 has been shown to inhibit SOD1 activity in lung cancer cells, presumably by blocking copper transfer from CCS to SOD1 (28). Control experiments with TM showed reduced total SOD and SOD1 activity with copper chelation (Fig. 4E and F). However, in most TNBC cells total SOD (SOD1 and SOD2) activity increased with DCAC50 treatment after 24 hours and increased further at 72 hours (Fig. 4E), perhaps in response to elevated oxidative stress. Specific SOD1 activity normalized to SOD1 expression decreased in DCAC50-treated MDA-MB-231 and HCC1806 cells (Fig. 4F; Supplementary Fig. S4E–S4G), as expected from the inhibition of copper transfer from CCS to SOD1. However, MDA-MB-468 and HCC1395 cell lines exhibited an increase in SOD1-specific activity, which may be due to the alternative pathways for copper delivery to the enzyme (36).

The increased DCF fluorescence, lower GSH:GSSG ratio and increased total SOD activity are consistent with increased oxidative stress resulting from disruption of copper transport by DCAC50. In contrast, global copper chelation with TM has no impact on DCF fluorescence and generally reduces total SOD and SOD1 activities. Taken together, these results suggest that the general copper dyshomeostasis caused by DCAC50 is a greater contributor to the observed oxidative stress than inhibition of copper transfer from CCS to SOD1.

Copper has been shown to directly stimulate angiogenesis whereas copper depletion by TM inhibits angiogenesis (23). Considering the recently reported pro-angiogenic activity of ATOX1 in endothelial cells (19), we tested if blocking ATOX1 activity with DCAC50 could suppress in vitro network formation in HuVEC cells. DCAC50 reduced the number of nodes, junctions, segments and branches and the length of these elements in a network-formation assay. A smaller effect was observed for TM at the tested dose. The treatment also inhibited extracellular LOX activity, but no dose-dependent effect on cell proliferation or apoptosis was observed with DCAC50 in the HuVEC cells (Fig. 5A–C; Supplementary Fig. S5A–S5B). Thus, in addition to the anticancer effects, DCAC50 may also modulate the microenvironment by blocking angiogenesis, which may prevent vascular remodeling that is required for tumor progression.

Taking into account the promising anticancer activity of DCAC50 in vitro, we evaluated the efficacy of copper transport inhibition in xenograft mouse models. In MDA-MB-468 xenografts, treatment with DCAC50 resulted in the inhibition of tumor growth and reduced tumor volumes, in comparison with the control (vehicle) group of mice (Fig. 5D; Supplementary Fig. S6A). Importantly, in addition to the inhibition of tumor growth in MDA-MB-468 xenografts, we observed significantly suppressed angiogenesis, as indicated by the reduced blood vessel area in DCAC50-treated mice, detected by staining with CD31⁺ and CD105⁺, vascular and neovascular endothelial cell markers, respectively (Fig. 5E–G). Reduced angiogenesis was also observed in MDA-MB-231 xenografts; although tumor volumes did not change overall at the end of treatment, slower tumor growth in 3 of 8 mice was observed (Supplementary Fig. S6B–S6D). The lower sensitivity of the MDA-MB-231 model to DCAC50 treatment is in concordance with our data observed in vitro. Thus, inhibition of intracellular copper transport by DCAC50 can suppress tumorigenesis by targeting both tumor cells and the tumor microenvironment, but its anti-tumor activity varies due to different sensitivities of TNBC cells to the disruption of copper homeostasis.

Treatment of patients with TNBC relies heavily on chemotherapy, including anthracyclines, taxanes, platinum agents and their combinations. The combination of chemotherapy with targeted therapy may result in fewer adverse effects and improve therapeutic efficacy, particularly in advanced cases. To explore this concept we evaluated the benefits of DCAC50 treatment in combination with paclitaxel. We conducted multi-drug combination dose–response analysis in TNBC cells according to Chou-Talalay (ref. 29; Fig. 6A). Co-treatment of paclitaxel and DCAC50 resulted in a CI less than one, suggesting additive to synergistic cytotoxicity (Fig. 6B). In addition, we observed a favorable dose reduction for both drugs, when cell lines were treated with the above combination, as indicated by DRI values ranging from 1.60 to 3.45 for DCAC50 and 2.04 to 3.72 for paclitaxel. Although the efficacy of the treatment combination leads to the additive cytotoxicity in MDA-MB-231, MDA-MB-436, MDA-MB-468 and HCC1395 cell lines (CI ranging from 0.9 to 1.1), we observed a synergistic response in HCC1187 and HCC1806 cell lines (CI < 0.9). Because the cytotoxicity of DCAC50 in tumor cells is a consequence of elevated oxidative stress, we hypothesized that the greater cytotoxicity of the co-treatment could be explained by increased oxidative stress, compared with paclitaxel alone. Co-treatment of paclitaxel and DCAC50 resulted in significantly higher oxidative stress in all examined cancer cell lines, in comparison with paclitaxel alone (Fig. 6C). Oxidative stress in paclitaxel-treated cells is reduced, compared with DMSO-treated cells. Thus, combination of DCAC50 with paclitaxel increases oxidative stress in tumor cells resulting in greater cytotoxicity.

In breast cancer, elevated levels of copper and copper-related proteins are associated with advanced disease (37–39), remodeling of the tumor microenvironment (14, 26) and resistance to chemotherapy (39, 40). Thus, copper homeostasis and the copper proteome are targets for breast cancer therapy. We have investigated an emerging approach to targeting copper-dependent cellular functions via the inhibition of intracellular copper transport in TNBC using the novel small-molecule DCAC50. We demonstrate that DCAC50 reduces cell proliferation and induces apoptosis through the inhibition of copper transport, disruption of cellular copper homeostasis, and the generation of oxidative stress in a panel of TNBC cell lines. Furthermore, DCAC50 enhances the cytotoxicity of paclitaxel in vitro, and inhibits tumor growth and angiogenesis in a xenograft mouse model.

The inhibition of ATOX1 and CCS activity with DCAC50 is expected to disrupt copper homeostasis and the copper proteome. In our study, significant increases in total intracellular copper levels were observed with DCAC50 treatment in claudin-low cells and modest increases in basal-like cells. Copper distribution was changed, with copper accumulating in the nuclei in MDA-MB-468 cells, to a lesser extent in HCC1395 nuclei or not at all in MDA-MB-231 nuclei. Attenuated cell proliferation and induced apoptosis were observed in all DCAC50-treated cell lines. Thus, the alteration of copper homeostasis, observed as an increase in total copper content and/or changes in copper distribution, leads to cytotoxicity. Nuclear localization of copper has previously been reported in Atox1^−/− mouse embryonic fibroblasts (41), in ATP7A-deficient skin fibroblasts (42) and in Atp7b^−/− mouse hepatocytes (43). However, the consequences of nuclear copper localization on cell proliferation, and the question of how copper enters the nuclei when the copper transport pathway is perturbed, remain to be elucidated. ATOX1 has been identified as a nuclear transcription factor and may undergo copper-dependent translocation to the nucleus (44). However, ATOX1 distributions were unchanged in DCAC50-treated cells (Supplementary Fig. S4A).

Various copper transporters regulate cellular copper content and its distribution. Cu-ATPases—downstream targets of ATOX1—translocate between the perinuclear space associated with TGN, where they supply copper to cuproenzymes, and the plasma membrane, to export copper from the cell (16). Copper binding is necessary for Cu-ATPase re-localization, and Cu-ATPase mutants unable to bind copper are restricted to the TGN (45). Thus, the inhibition of copper transfer from ATOX1 is expected to alter the distribution of Cu-ATPases. In MDA-MB-468 cells, DCAC50 changed the localization of ATP7B resulting in enhanced ATP7B clustering near the nuclei, which is indicative of impaired copper homeostasis. However, DCAC50 did not induce noticeable changes in protein levels of ATP7B, ATOX1, CTR1 or CCS, indicating no effect of the treatment on protein synthesis or degradation. In summary, DCAC50 causes changes in ATP7B and copper localization, but the detailed mechanism by which copper is translocated to the nucleus and how this impacts cell proliferation and survival remains unclear.

Copper homeostasis and copper-dependent proteins are intimately linked with the redox status of cells. Intracellular copper homeostasis is tightly regulated to protect cells from exposure to the redox-active metal. Free copper can induce superoxide generation and bind glutathione to create an oxidizing environment. On the other hand, copper-dependent SOD1 and total SOD (SOD1 and SOD2) activities are responsive to superoxide levels and protect cells from excessive oxidative stress. ATOX1 itself has a central role linking copper and redox homeostasis (46). Therefore, disruption of copper homeostasis and the copper proteome is expected to disrupt cellular redox status. We have observed increased oxidative stress in all DCAC50-treated cells, measured by general ROS levels and the change in the GSH:GSSG ratio. Overall, oxidative stress precedes the induction of apoptosis in cancer cells, suggesting that elevation of ROS is the reason for cytotoxicity. Inhibition of intracellular copper transport by DCAC50 is expected to inhibit the activity of copper-dependent SOD1. In our study, total SOD activity increased over time with increasing ROS levels, presumably in an effort to modulate oxidative stress. At the same time, SOD1 activity, normalized to SOD1 expression, decreased with DCAC50 treatment in some cell lines, as expected with the inhibition of copper transfer from CCS to SOD1, but increased in others. SOD1 has a half-life of more than 30 hours in mammalian cells (47) and alternative pathways exist for the delivery of copper to the enzyme (36), which may mitigate the impact of DCAC50 on SOD1 activity.

ATOX1 and CCS protein expression levels and IC₅₀ values for DCAC50 were established in eight breast cancer cell lines and in normal mammary epithelial cells (HMEC). Low micromolar IC₅₀ values were obtained in all cell lines, with no clear correlation between copper chaperone protein levels and sensitivity to DCAC50. Ultimately, six TNBC cell lines with variable elevated levels of ATOX1 and CCS were selected to investigate the effects of copper transport inhibition. We found that DCAC50 inhibited proliferation and induced apoptosis more effectively in basal-like than in claudin-low cell lines. To explain the broadly different responses of these subtypes of cells to DCAC50, we must consider cellular copper and redox statuses. High copper levels at baseline and large increases in intracellular copper levels with treatment were observed in claudin-low cells (MDA-MB-231, MDA-MB-436, HCC1395), yet DCAC50 was less effective in these cells lines. Basal-like MDA-MB-468 cells, which had lower baseline copper levels that increased only modestly with treatment, exhibited noticeable disruption to copper distribution and were particularly sensitive to DCAC50-induced apoptosis. Denoyer and colleagues (48) have argued that the sensitivity of cancer cells to ROS-generating copper ionophores may be attributed to limited antioxidant capacity rather than to high basal copper levels. Our data indicate that the ability of breast cancer cells to control ROS production is an important contributor to sensitivity to DCAC50. One of the major cellular antioxidant molecules, glutathione, binds copper and has been shown to protect cells from ROS toxicity (49, 50). In mouse embryonic fibroblasts, when glutathione levels are low, ATOX1 deficiency has been shown to disrupt copper homeostasis leading to cell death (51). This evidence suggests that when glutathione levels are low and ATOX1 is limited, cells are susceptible to copper dyshomeostasis and oxidative stress. In our study, basal-like cells (MDA-MB-468, HCC1187, HCC1806) had overall lower copper levels and lower total GSH levels at baseline, in comparison with claudin-low cells. Thus, basal-like cells may have a lower threshold for copper overload, and may be more sensitive to copper dyshomeostasis and oxidative stress when ATOX1 is inhibited by DCAC50. In contrast, claudin-low cells possess higher levels of glutathione and resist oxidative stress caused by DCAC50 treatment. Our results demonstrate that the GSH synthesis inhibitor, BSO, potentiates DCAC50 inhibition of cell proliferation in claudin-low HCC1395 and MDA-MB-436 cells by depleting glutathione levels and thus exhausting cellular antioxidant defense.

Of all the TNBC cell lines evaluated in this study, MDA-MB-231 cells are most resistant to DCAC50′s cytotoxic effects, which may be a consequence of distinct genetic alterations promoting cell growth and survival. MDA-MB-231 cell line is an invasive, highly metastatic claudin-low cell line that bears activating mutations in Kras, Braf, and Pdgfra, which have previously been shown to upregulate cell proliferation and survival through activation of MAPK and PI3K (52, 53). Moreover, MDA-MB-231 cells can upregulate the antiapoptotic gene Bcl2a1 upon tumor progression (54). Thus, activation of these alternative signaling pathways may contribute to the ability of MDA-MB-231 cells to resist DCAC50 treatment. In comparison with cancer cells, the disruption of copper homeostasis with DCAC50 triggered a limited response in normal cells (HMEC and HuVEC). DCAC50 attenuated cell proliferation in HMEC and inhibited network-formation in HuVEC (without reducing cell proliferation), but no induction of apoptosis was detected in either of the normal cell lines. DCAC50 has previously been shown to have a minimal effect on the proliferation of other normal human cell lines (28). The ability of cells to activate pathways to resist copper toxicity and oxidative stress, may explain heterogeneous sensitivity to DCAC50 and to the associated copper overload.

Reactive oxygen species are relevant to paclitaxel cytotoxicity, and resistance to paclitaxel relies, at least partially, on cellular total antioxidant capacity (55). Co-treatment with DCAC50 elevates oxidative stress, and is expected to enhance cytotoxicity in paclitaxel-treated cells. We observed increased production of oxidative stress and improved efficacy in cells treated by paclitaxel in combination with DCAC50, in comparison with paclitaxel alone. We hypothesize that the efficacy of the combination is due to the interference of multiple pathways involved in the defense against oxidative stress. The combination was effective to reduce treatment doses for both drugs, and may help reduce paclitaxel-associated side effects and reverse paclitaxel-associated resistance in tumor cells.

The findings presented here support our hypothesis that DCAC50 inhibits copper transport and alters copper homeostasis, disrupting the redox balance and causing cytotoxicity in cancer cells. The varying efficacy of DCAC50 reflects the heterogeneity of breast cancer cell lines and suggests that the success of drugs targeting copper homeostasis may depend on baseline cellular copper and redox statuses. Our findings also reveal the differences between targeting copper homeostasis, via the intracellular copper transport pathway, versus global copper chelation. Recent studies suggest that copper chelation is clinically important, especially for high-risk patients with breast cancer and in the prevention of angiogenesis and metastasis. In our study, TM did not have a significant effect on cancer cell growth or survival, however it did inhibit endothelial cell network-formation in vitro, in accordance with its known antiangiogenic activity (23). Importantly, DCAC50 targets tumor cell viability as well as the tumor microenvironment, thus the inhibition of intracellular copper transport has the potential to become an effective strategy to treat breast cancer.

We conclude that dual effects of triggering apoptosis in tumor cells and modulating the tumor vascular environment are extremely attractive and should prompt further evaluation of the copper transport pathway as a therapeutic target in breast cancer.

C. He has ownership interest (including stock, patents, etc.) in Accent Therapeutics and Epican Genetech, and is a consultant/advisory board member for Accenter Therapeutics. O.I. Olopade is a consultant/advisory board member for Tempus, CancerIQ, and HLF. No potential conflicts of interest were disclosed by the other authors.

Conception and design: O. Karginova, C.M. Weekley, O.I. Olopade

Development of methodology: O. Karginova, C.M. Weekley

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): O. Karginova, C.M. Weekley, A. Raoul, A. Alsayed, T. Wu

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): O. Karginova, C.M. Weekley, A. Raoul, A. Alsayed, S.S.-Y. Lee

Writing, review, and/or revision of the manuscript: O. Karginova, C.M. Weekley, C. He, O.I. Olopade

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): O. Karginova, C.M. Weekley, T. Wu

Study supervision: O. Karginova, C. He, O.I. Olopade

Use of the Advanced Photon Source at Argonne National Laboratory was supported by the U.S. Department of Energy, Office of Science, and Office of Basic Energy Sciences, under contract DE-AC02-06CH11357. We thank the U.S. Department of Energy, under contract number DE-FG02-07ER15865 (to C. He), for partial support of this work. We acknowledge funding from Breast Cancer Research Foundation FP049439 (to O.I. Olopade), the National Health and Medical Research Council (Australia; to C.M. Weekley; CJ Martin Overseas Biomedical Fellowship APP1090612), and NIBIB K99 EB022636 (to S.S.-Y. Lee). We thank Dr. Andrei Karginov for helpful comments and assistance with cell imaging, and Dr. Barry Lai for his assistance with XFM imaging.

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.