Antitumor Activity of Metal-Chelating Compound Dp44mT Is Mediated by Formation of a Redox-Active Copper Complex That Accumulates in Lysosomes

Neoplastic cells have high requirements for iron (Fe) due to their generally higher rates of proliferation than normal cells (1). In fact, neoplastic cells express enhanced transferrin receptor 1 (TfR1) levels relative to their normal counterparts (2) and take up Fe from transferrin (Tf) at a rapid rate (3), making them selectively sensitive to Fe chelation. Cancer cells also take up more copper (Cu) than their normal counterparts, as this metal is essential for angiogenesis and metastasis (4).

Considering the crucial roles of these metals, development of novel Fe and Cu chelators has become a promising anticancer strategy (1, 5). Indeed, the chelator, Triapine (3-aminopyridine-2-carboxaldehyde thiosemicarbazone; Fig. 1), which inhibits tumor growth, has entered clinical trials (1). However, the di-2-pyridylketone thiosemicarbazone (DpT) chelators possess far greater antitumor activity and selectivity than Triapine (6, 7).

One of the most effective DpT chelators is di-2-pyridylketone-4,4,-dimethyl-3-thiosemicarbazone (Dp44mT; Fig. 1; ref. 6). Dp44mT shows marked and selective activity against tumor xenografts in mice (6, 7). The proposed mechanism of action of Dp44mT involves Fe chelation and redox cycling of its Fe complex to generate reactive oxygen species (ROS; refs. 6, 8, 9). The potent and antitumor activity of Dp44mT has been verified by others (10–12). In particular, Rao and colleagues reported topoisomerase IIα inhibition (10) and broad and specific antitumor activity (12). Additional modes of anticancer activity reported for Dp44mT include inhibition of the metastasis suppressor protein, Ndrg-1 (1), and modulation of the cell-cycle control proteins of the cyclin family (A, B, D1, D2, and D3) as well as cyclin-dependent kinase 2 (1, 5).

Dp44mT not only binds Fe, but also Cu, and both complexes are redox active, contributing to their marked cytotoxicity (13). Considering: (i) the potential of lysosomes as a therapeutic target; (ii) their key role in metal metabolism (14, 15); and (iii) their pronounced susceptibility to ROS (15, 16), our goals were to examine the effects of the DpT compounds on Cu and Fe and their interaction with lysosomes. Our work has elucidated a novel strategy for the design of new therapeutics that activate the lysosomal apoptotic pathway by binding redox-active Cu.

2-Hydroxy-1-naphthaldehyde isonicotinoyl hydrazone (311), di-2-pyridylketone-2-methyl-3-thiosemicarbazone (Dp2mT), di-2-pyridylketone-4-ethyl-3-thiosemicarbazone (Dp4eT), di-2-pyridylketone-4-allyl-3-thiosemicarbazone (Dp4aT), di-2-pyridylketone-4-methyl-3-thiosemicarbazone (Dp4mT), Dp44mT, di-2-pyridylketone-4-phenyl-3-thiosemicarbazone (Dp4pT), and pyridoxal isonicotinoyl hydrazone (PIH) were synthesized as described (6, 8). Desferrioxamine (DFO) was from Novartis. Buthionine sulfoximine (BSO), d-penicillamine (d-pen), tetrathiomolybdate (TM), and Trientine (Trien) were purchased from Sigma-Aldrich. Triapine was from Vion Pharmaceuticals.

Human SK-N-MC neuroepithelioma cells, MCF-7 breast cancer cells, DMS-53 lung carcinoma cells, and human umbilical vein endothelial cells (HUVEC) were obtained from the American Type Culture Collection and grown as described (17, 18). Cells were used within 2 months of purchase after resuscitation of frozen aliquots. Cell lines were authenticated on the basis of viability, recovery, growth, morphology, and also cytogenetic analysis, antigen expression, DNA profile, and isoenzymology by the provider.

Procedures involved in preparing ⁵⁹Fe-transferrin and conducting cytotoxicity assays have been described previously (17, 18).

Efflux studies examining the ability of chelators to mobilize ⁵⁹Fe or ⁶⁴Cu (10 μCi/mL; ⁶⁴CuCl₂; ANSTO) from cells were done by standard methods (17). Briefly, cells were prelabeled for 3 hours at 37°C with ⁶⁴Cu (10 μCi) or ⁵⁹Fe-transferrin (⁵⁹Fe-Tf; 0.75 μmol/L), washed 4 times on ice and reincubated with medium (control) or medium and chelators (25 μmol/L) for up to 6 hours at 37°C and then harvested (17). Radioactivity was measured in the pellet and supernatant using a γ-scintillation counter (Wallac Wizard 3; Perkin Elmer).

Complexes were prepared by adding equimolar ⁶⁴Cu and chelator. SK-N-MC cells were incubated with the complexes for 3 hours at 37°C, washed 4 times on ice, reincubated for 3 hours at 37°C in control media, and the percentage of ⁶⁴Cu remaining cell associated assessed (17).

SK-N-MC cells (3 × T175 flasks, 80% confluent) were prelabeled for 3 hours at 37°C with ⁶⁴Cu (10 μCi), washed and reincubated at 37°C with medium (control) or Dp44mT (5 μmol/L) for 3 hours at 37°C. Cells were added to extraction buffer (0.25 mol/L sucrose, 10 mmol/L Tris-HCl) and disrupted using a Dounce “B” homogenizer on ice. To prevent damage to the released lysosomes, disruption of cells was discontinued when approximately 50% of the intact cells were disrupted. The suspension was centrifuged 1,000 × g for 15 minutes at 4°C, the pellet discarded, and supernatant centrifuged at 16,000 × g for 15 minutes at 4°C to yield a crude lysosomal/mitochondrial fraction. As a further control, enzyme marker analysis for the lysosomal specific enzyme acid phosphatase (Sigma) showed that this fraction was enriched with lysosomes (as determined by spectrophotometric assessment at 405 nm).

Potentiometric titrations were conducted as described (8).

Distribution of acridine orange (Sigma) was used to determine lysosomal membrane permeability (LMP) as previously reported (18) and was quantified by flow cytometry (19). Briefly, cells were incubated for 15 minutes at 37°C with acridine orange (20 μmol/L), washed 3 times with PBS, and then incubated for 30 minutes at 37°C with 25 μmol/L chelator or reagent. Cathepsin D release was examined by fluorescence microscopy as previously reported (20) using a pepstatin A–BODIPY FL conjugate (Invitrogen) that selectively binds to cathepsin D. LysoTracker red (Invitrogen) was used to confirm colocalization of cathepsin D with lysosomes. For details, see Supplementary Methods.

Assessment of Bid cleavage was assessed by Western blotting by standard methods (6) using antibodies to Bid (Cell Signaling Technology) and β-actin (clone AC-1; Sigma).

Tetramethylrhodamine ethyl ester (TMRE; Invitrogen) partitions to the mitochondrial matrix (21). SK-N-MC cells were incubated with 20 nmol/L of TMRE for 0.25 hours at 37°C and then incubated with 5 μmol/L of Cu[Dp44mT] for 0.5 to 2 hours at 37°C. Mitochondrial damage was examined by flow cytometry (21).

This assay was conducted using a kit (Calbiochem). SK-N-MC cells were treated at 25 μmol/L for up to 24 hours at 37°C with Cu(II), Fe(III), Dp44mT, Cu[Dp44mT], or Fe[Dp44mT]₂. As a control, BSO (100 μmol/L) was added as a glutathione synthesis inhibitor (22).

Studies without cells were conducted as described with H₂DCF (5 μmol/L; ref. 23). As a positive control, Fe(III) at 5 μmol/L was reduced to Fe(II) using cysteine (100 μmol/L) in 150 mmol/L acetate buffer (pH = 5.0). Hydrogen peroxide (100 μmol/L) was then added to initiate hydroxyl radical generation. To confirm hydroxyl radical production, dimethyl sulfoxide (DMSO; 10% v/v) was used, as it has been reported to be an effective hydroxyl radical scavenger (24, 25). Cu[Dp44mT] and other reagents (at 5 μmol/L, except DMSO at 10% v/v) were added to examine hydroxyl radical production. Intracellular oxidation of H₂DCF studies were conducted as described in previous studies (6, 23).

Data were compared using Student's t test. Results were expressed as mean ± SD (number of experiments) and considered to be statistically significant when P < 0.05.

To assess the antineoplastic mechanisms of the DpT analogues (Fig. 1), their ability to remove ⁶⁴Cu from prelabeled cells was compared with their interaction with ⁵⁹Fe. The activity of DpT chelators was compared with the Fe chelators, DFO, PIH, and 311 (17), and Cu chelators, d-pen, Trien, and TM (Fig. 1; ref. 1). The DpT chelators, Dp4mT, Dp44mT, Dp4eT, Dp4aT, and Dp4pT, were found to be ineffective at 25 μmol/L at inducing ⁶⁴Cu release from cells (leading to ⁶⁴Cu retention) relative to when cells were incubated with control medium (Fig. 2A). In these studies, SK-N-MC cells were used as their response to chelators is well characterized (6, 17).

The results above were in contrast to their ability to mobilize ⁵⁹Fe, where the DpT chelators significantly (P < 0.001) induced a more than 700% increase in ⁵⁹Fe release relative to control cells (Fig. 2B). Hence, there was a difference in the ability of DpT chelators to mobilize these metals. The only exception was the negative control, Dp2mT, which failed to induce significant ⁶⁴Cu or ⁵⁹Fe efflux (Fig. 2A and B). By design, Dp2mT cannot bind metals (6), showing the importance of metal binding for DpT chelator activity. The chelators DFO, PIH, and 311 were included as positive controls to increase ⁵⁹Fe efflux, and we also examined their effects on ⁶⁴Cu release. Only 311 showed a marked interaction with ⁶⁴Cu, significantly (P < 0.001) reducing its efflux to 43% ± 10% of the control (Fig. 2A). The Cu chelators, d-pen and Trien, were ineffective at inducing ⁶⁴Cu efflux at 25 μmol/L (Fig. 2A). However, the Cu chelator, TM, significantly (P < 0.05) reduced ⁶⁴Cu efflux to 76% ± 2% of the control. The low activity of the Cu chelators, d-pen, Trien, and TM, at mobilizing ⁶⁴Cu was unexpected on the basis of their in vivo efficacy (1). However, their mechanism of action in cell culture is unclear, and the efficacy in vivo could relate to extracellular rather than intracellular chelation (26). These efflux experiments were repeated using MCF-7 (Fig. 2C and D), and to provide a comparison with tumor cells, normal HUVECs were used (Fig. 2E and F). Generally, the active chelators accumulated ⁶⁴Cu similarly across the 3 cell types (Fig. 2A–F). In terms of the effect of Dp44mT on inhibiting ⁶⁴Cu release, its effect was greater in HUVECs and MCF-7 cells than SK-N-MC.

The kinetics of ⁶⁴Cu and ⁵⁹Fe release were assessed over a range of reincubation times (0.5–6 hours) using SK-N-MC cells and HUVECs that were prelabeled with ⁶⁴Cu or ⁵⁹Fe-Tf for 3 hours at 37°C (Supplementary Fig. S1A–D). These results showed that Dp44mT and 311 decreased ⁶⁴Cu release relative to the control while both ligands markedly increased ⁵⁹Fe efflux as a function of time. In addition, dose–response curves showed that 311 and Dp44mT decreased ⁶⁴Cu release compared with the control, leading to ⁶⁴Cu accumulation in cells (Supplementary Fig. S1E). With Dp44mT, this effect was reversible as its concentration increased to more than 5 μmol/L, and at these higher concentrations, cellular damage results in the release of the Cu–Dp44mT complex from the cells. In contrast, at lower concentrations, the cells are still intact preventing ⁶⁴Cu release. In terms of ⁵⁹Fe efflux, 311 and Dp44mT markedly increased cellular ⁵⁹Fe release as a function of concentration (Supplementary Fig. S1F). Collectively, the active DpT group of chelators markedly prevented ⁶⁴Cu efflux from all cell types, leading to ⁶⁴Cu accumulation.

The results indicating intracellular accumulation of DpT–⁶⁴Cu complexes were confirmed by studies where SK-N-MC cells were labeled for 3 hours at 37°C with preformed ⁶⁴Cu–chelator complexes. The cells were then washed and reincubated for 3 hours at 37°C in control media and the percentage of ⁶⁴Cu remaining cell associated then quantified. In control cells incubated with ⁶⁴Cu only, only 4% ± 1% of ⁶⁴Cu remained intracellular (Fig. 3A). Hence, most ⁶⁴Cu had been released during the reincubation. Cells labeled with the ⁶⁴Cu complexes of the active DpT analogues (Dp4mT, Dp44mT, Dp4eT, Dp4aT, and Dp4pT) and then reincubated with control medium, showed that 39% to 68% of ⁶⁴Cu remained cell associated (Fig. 3A). In contrast, ⁶⁴Cu complexes of Dp2mT, d-pen, and Trien were no more effective than the control at retaining intracellular ⁶⁴Cu (Fig. 3A). The ⁶⁴Cu–TM complex also led to intracellular ⁶⁴Cu accumulation relative to the control. These results explain the low efflux of ⁶⁴Cu observed with the active DpT analogues (Fig. 2A), as their ⁶⁴Cu complexes become “trapped” within cells.

To examine the subcellular localization of the accumulated ⁶⁴Cu, SK-N-MC cells were labeled for 3 hours at 37°C with ⁶⁴Cu, washed, and reincubated for 3 hours at 37°C with Dp44mT (5 μmol/L) or control media. The distribution of ⁶⁴Cu in the cytosol and crude lysosomal/mitochondrial fraction was examined by differential centrifugation. In cells reincubated with Dp44mT, a significantly (P < 0.001) greater proportion of ⁶⁴Cu accumulated in the lysosomal/mitochondrial fraction (Fig. 3B).

Considering this observation, we hypothesized that because of the polyprotic nature of Dp44mT (8), it was trapped in acidic lysosomes as it became positively charged. To assess this, speciation studies of the chelator as a function of pH were conducted. At a pH of 7.4, 100% of the ligand is in its neutral state (Dp44mT), allowing facile transport through membranes, whereas at a lysosomal pH of 5, 16% is charged leading to lysosomal accumulation (Dp44mT⁺; Fig. 3C). It should be noted that while 16% of the ligand is protonated at a pH of 5, this leads to accumulation of the ligand over time. This occurs due to the process of: (i) the neutral ligand entering the lysosome; (ii) the ligand becoming protonated and charged at lysosomal pH (pH = 5), preventing its passage out of the organelle; (iii) the charged ligand-binding copper in the lysosome; and (iv), as the so formed copper complex is probably positively charged, it also cannot escape the lysosome accounting for the ⁶⁴Cu accumulation in this compartment (see Fig. 3B).

To further explore the potential effect of Dp44mT and its Cu complex (Cu[Dp44mT]) on the lysosome, we implemented the lysosomotropic fluorophore, acridine orange (Fig. 3D) that accumulates within lysosomes (18). High lysosomal concentrations of acridine orange give a red fluorescence, whereas lower cytosolic and nuclear concentrations give a green fluorescence (18).

Examining control cells by fluorescence microscopy, a granular red fluorescence consistent with acridine orange concentration in lysosomes was found (Fig. 3D; ref. 18). However, incubation of cells with 25 μmol/L of Cu[Dp44mT] for 0.5 hours resulted in a marked loss of red fluorescence and the disappearance of red vesicles consistent with increased LMP, as well as the appearance of apoptotic bodies (Fig. 3D). In contrast, the Fe[Dp44mT]₂ complex, Dp44mT, CuCl₂, or FeCl₃ (at 25 μmol/L), had no significant effect relative to the control (Fig. 3D). The lack of activity of Dp44mT or Fe[Dp44mT]₂ is due to the short incubation period (0.5 hours) used, which was optimal for detecting the effect of Cu[Dp44mT]. Indeed, Dp44mT and Fe[Dp44mT]₂ only induce significant cytotoxicity after 24 hours (Fig. 6A). These data using fluorescence microscopy were confirmed upon quantification by flow cytometry, where only the Cu[Dp44mT] complex (25 μmol/L) significantly (P < 0.001) reduced red fluorescence (Supplementary Fig. S2).

To further examine the effect of Cu[Dp44mT] on the lysosome, we examined the intracellular distribution of LysoTracker red and a lysosomal enzyme, cathepsin D (18), utilizing a fluorescent probe (pepstatin A–BODIPY FL) that binds to cathepsin D (20). Control cells stained with LysoTracker red and pepstatin A–BODIPY FL showed a granular/vesicular pattern consistent with lysosomes (Fig. 4A). The overlay of LysoTracker red and pepstatin A–BODIPY FL–stained cells showed colocalization. After a 0.5-hour incubation with 25 μmol/L of Cu[Dp44mT], the granular, lysosomal type pattern disappeared, with the fluorescence becoming evenly distributed within the cytosol (Fig. 4A). This observation was consistent with Cu[Dp44mT]-induced LMP, confirming results with acridine orange (Fig. 3D). Neither Cu(II) as CuCl₂ (25 μmol/L) or Dp44mT (25 μmol/L) alone had any significant effect on LysoTracker red or cathepsin D staining (Fig. 4A).

To show a relationship between altered LMP and apoptosis induction, Western blotting assessed cleavage of the proapoptotic Bcl-2 family member, BH3-interacting domain death agonist (Bid), by lysosomal proteases (ref. 27; Fig. 4B). As a positive control, cells were incubated with cisplatin (20 mmol/L) that induces Bid cleavage (27). SK-N-MC cells incubated with 5 μmol/L of Cu[Dp44mT] or 20 mmol/L of cisplatin for 2 hours showed significant (P < 0.001) cleavage of Bid (Fig. 4B). This confirmed redistribution of lysosomal proteases to the cytosol (Fig. 4A) leading to Bid cleavage and provides a direct link between lysosomal damage and apoptosis induction by Cu[Dp44mT].

We next examined whether mitochondria were also damaged. Mitochondrial stability was determined by measuring the reduction in mitochondrial membrane potential using TMRE (21), whereas LMP was assayed using acridine orange via flow cytometry. As in Fig. 4C, the damage induced by 5 μmol/L of Cu[Dp44mT] to lysosomes was significantly (P < 0.05–0.001) more apparent than that to mitochondria after 1 or 2 hours, indicating that lysosomes were more sensitive to Cu[Dp44mT].

To understand why lysosomes were so sensitive to Cu[Dp44mT], we assessed whether the Fe or Cu complexes of Dp44mT generate oxidative stress that could damage lysosomes. These studies in a cell-free system were conducted under lysosomal like conditions at a pH of 5.0 and in the presence of cysteine (ref. 28; Fig. 5A and B). Oxidative stress was determined by the oxidation of nonfluorescent H₂DCF to fluorescent DCF, a well-characterized probe for assessing redox stress (6, 23). The high redox activity of 5 μmol/L of Cu[Dp44mT] was significantly (P < 0.01) greater than 5 μmol/L of CuCl₂, FeCl₃, and Fe[Dp44mT]₂ after only 2 minutes (Fig. 5A). Addition of the 5 μmol/L of Cu chelator TM to 5 μmol/L of Cu[Dp44mT] totally prevented its activity, consistent with the ability of TM to bind Cu from Cu[Dp44mT], as shown in Fig. 5A and B.

Considering these results, the potential of Cu[Dp44mT] to induce intracellular ROS in SK-N-MC cells was also assessed using H₂DCF. At 5 μmol/L, Cu[Dp44mT] caused a significant (P < 0.001) increase in intracellular H₂DCF oxidation to 273% ± 20% of that found for control cells after a 0.5-hour incubation (Fig. 5C). No increase in H₂DCF oxidation was observed with Cu(II) as CuCl₂, Fe(III) as FeCl₃, Dp44mT alone, or Fe[Dp44mT]₂, even at a 5-fold higher concentration (25 μmol/L; Fig. 5C). Collectively, these experiments show the marked redox activity of Cu[Dp44mT].

To examine the redox activity of Cu[Dp44mT] on physiologic substrates, levels of the well-described indicator of oxidative stress, glutathione (GSH), and oxidized GSH (GSSG; ref. 22) were determined. Over 0.5 to 24 hours incubations with SK-N-MC cells, 25 μmol/L of Cu[Dp44mT] significantly (P < 0.001) reduced the GSH/GSSG ratio to 0% to 7% of the control (Fig. 5D), confirming that Cu[Dp44mT] possesses pronounced redox activity. In comparison, Dp44mT (25 μmol/L), the GSH synthesis inhibitor, BSO (100 μmol/L), or Fe[Dp44mT]₂ (25 μmol/L) also reduced the GSH/GSSG ratio to 4% to 20% of the control but only after 24 hours (Fig. 5D).

The studies above indicate that Dp44mT enters cells and forms a redox-active Cu complex that damages lysosomes. Considering this, we examined the cytotoxicity of Dp44mT and its Cu and Fe complexes. As shown in Fig. 6A, after a 3-hour incubation, 5 μmol/L of Cu[Dp44mT] significantly (P < 0.001) decreased viability to 28% ± 4% of the control. In contrast, Dp44mT or Fe[Dp44mT]₂ (both at 5 μmol/L) did not significantly decrease viability within 3 hours but showed activity at 24 hours (Fig. 6A). These results show the pronounced cytotoxicity of Cu[Dp44mT] relative to Fe[Dp44mT]₂ or Dp44mT.

To determine the importance of Cu chelation to the cytotoxicity, we incubated cells for 72 hours at 37°C with increasing Dp44mT concentrations, Dp44mT in the presence of the Cu chelator, TM, or the preformed TM–Cu complex, which blocks the ability of TM to bind cellular Cu (Fig. 6B). The addition of TM to Dp44mT markedly prevented the ability of the latter to reduce cellular viability up to a Dp44mT concentration of 2.5 μmol/L, whereas the preformed Cu-TM complex (which cannot bind cellular Cu) had no significant rescue effect on Dp44mT cytotoxicity (Fig. 6B).

To confirm that the ability of TM to rescue Dp44mT cytotoxicity was due to Cu chelation, we used the structurally unrelated Cu chelator, bathocuproine disulfonate (BCS), in identical studies and observed a similar, but less pronounced, rescue effect on Dp44mT cytotoxicity (Fig. 6B). Like TM, BCS decreased Dp44mT cytotoxicity, whereas its Cu complex (BCS-Cu) had no significant influence (Fig. 6B), as its binding site was saturated with Cu. Notably, BCS or TM alone or their Cu complexes had no significant effect on viability (Supplementary Fig. S3).

As Dp44mT concentration increased, the ability of the Cu chelators BCS or TM to prevent cytotoxicity was markedly reduced after 72 hours (Fig. 6B). This may reflect chelation of Fe by Dp44mT which becomes significant at these higher concentrations and leads to Fe deprivation and cytotoxicity (6). Indeed, while the binding of Cu by Dp44mT is crucial for its antitumor efficacy, its Fe chelation efficacy is also important (6, 8).

Because GSH plays an important role in buffering redox stress (22) and considering the marked redox activity of Cu[Dp44mT], as shown in Fig. 5A–C, further studies examined its effect on GSH (Fig. 6C–F). To investigate this, the effect of GSH supplementation or depletion on viability was assessed by incubating cells in the presence of N-acetylcysteine (NAC) that enhances GSH levels (22), or the GSH synthesis inhibitor, BSO, that decreases GSH (22). Indeed, a 24-hour incubation of SK-N-MC cells with NAC (5 mmol/L) or BSO (100 μmol/L) significantly (P < 0.001) increased and decreased the GSH/GSSG ratio to 155% ± 5% and 4% ± 2% (n = 3) of the control, respectively.

The addition of NAC (5 mmol/L) with Cu[Dp44mT] significantly (P < 0.001) prevented the decrease in viability due to Cu[Dp44mT] after 3 and 24 hours (Fig. 6C), and similar results were observed as a function of concentration over 72 hours (Fig. 6D). Supplementation with NAC also significantly (P < 0.01) protected against the decreased viability observed with Fe[Dp44mT]₂ or Dp44mT alone after 72 hours (Fig. 6C and D), and thus decreased their cytotoxicity. Further, morphologic studies using acridine orange also showed that NAC preserved lysosomal integrity in the presence of Cu[Dp44mT], as shown in Supplementary Fig. S4.

In contrast to NAC, the addition of BSO with Dp44mT and Fe[Dp44mT]₂ significantly (P < 0.001) enhanced their cytotoxicity particularly at 24 hours (Fig. 6E). However, the addition of BSO with 5 μmol/L of Cu[Dp44mT] did not increase its cytotoxicity (Fig. 6E). This may be due to the marked redox activity of Cu[Dp44mT] relative to Dp44mT and Fe[Dp44mT]₂ at high concentrations (Fig. 5A), which rapidly depresses GSH levels (Fig. 5D).

As a function of chelator or complex concentration (0.02–0.25 μmol/L), BSO potentiated cytotoxicity of Dp44mT, Fe[Dp44mT]₂, and Cu[Dp44mT] after 72 hours (Fig. 6F). Hence, at low Cu[Dp44mT] concentrations (i.e., 0.02–0.25 μmol/L; Fig. 6F), BSO clearly potentiated the cytotoxicity of this complex after 72 hours, in contrast to higher Cu[Dp44mT] concentrations (i.e., 5 μmol/L; Fig. 6E).

In this investigation, we showed that incubation of cells with Dp44mT leads to retention of its ⁶⁴Cu complex. The probable reason for this is the ionization characteristics of Dp44mT (8). At physiologic pH, Dp44mT is neutral and permeates cell membranes (6, 8). However, in the lysosomal compartment (pH ∼5; ref. 29), an increased proportion of Dp44mT becomes positively charged leading to accumulation, ROS formation, and LMP.

Dp44mT induces apoptosis using several cell types and a tumor model in vivo (6). Moreover, we showed that apoptosis occurred via the mitochondrial pathway, where decreased Bcl-2 and increased Bax expression occurred along with holo-cytochrome c (h-cytc) release and caspase activation (6). Our study suggests that these apoptotic events could be caused by Cu[Dp44mT]-induced redox stress that results in LMP, causing redistribution of lysosomal cathepsins to the cytosol (Fig. 4A) and concomitant cleavage of Bid into its proapoptotic form (Fig. 4B). Indeed, cathepsins can cleave Bid, which migrates to mitochondria and induces outer membrane permeabilization that is dependent on proapoptotic Bax (30). Bax plays a role in inducing release of mitochondrial h-cytc, thereby activating the caspase cascade (31), which we showed occurs after Dp44mT treatment (6). Hence, LMP by redox-active Cu[Dp44mT] could result in downstream effects on mitochondria that lead to h-cytc release.

The Dp44mT-Cu complex showed far greater cytotoxicity as a function of time than either the Fe complex or the ligand alone. This was also reflected in the faster kinetics of lysosomal rupture (Fig. 3D), cathepsin D release (Fig. 4A), and decrease in GSH/GSSG ratio (Fig. 5D) induced by Cu[Dp44mT]. As these events all occur over the same time scale, they suggest a coherent and coordinated series of events that ultimately induces significant cytotoxicity. Intriguingly, the importance of Cu in mediating the cytotoxicity of the free ligand was shown by the rescue effect of the nontoxic Cu chelators, TM and BCS. This indicated that formation of a redox-active Cu[Dp44mT] complex is important for Dp44mT activity. Moreover, it was shown that Cu[Dp44mT] results in a marked decrease in GSH and the cytotoxicity of this complex can be reduced by NAC, which enhances GSH levels (22). Conversely, cytotoxicity can be potentiated by the GSH inhibitor, BSO. These results indicate that lysosomal damage caused by Cu[Dp44mT] was directly due to its redox activity. On the basis of these data, we propose a model of the mechanism of action of Dp44mT (Fig. 7).

Another aspect of the mechanism of action of Dp44mT is that it shows selectivity against tumor cells in vitro and in vivo (7). The basis for this relates, in part, to the greater uptake and metabolism of Cu and Fe for essential processes in cancer cells relative to normal cells (5). Under such conditions, the lysosome could be more active in terms of its metal metabolism, and hence, more susceptible to Dp44mT. It is also known that lysosomal autophagic pathways in cancer cells are abnormal due to the monoallelic deletion of the essential autophagy regulator, beclin1 (32). As such, metal recycling due to autophagy could be disturbed and lead to differences between neoplastic and normal cells in their response to agents such as Dp44mT.

In summary, we have dissected the mechanism of action of Dp44mT and showed that it accumulates in lysosomes due to its unique ionization characteristics. Our investigation reveals the marked redox activity of Cu[Dp44mT] leads to LMP that induces cell death. Moreover, this study shows that targeting lysosomes can lead to potent and selective anticancer therapeutics. Knowledge of this mechanism can be used to design more potent cytotoxic agents that affect the lysosomal apoptosis pathway.

No potential conflicts of interest were disclosed.

D.R. Richardson thanks the National Health and Medical Research Council for a Senior Principal Research Fellowship and Project Grant support. D.B. Lovejoy and P.J. Jansson thank the Cancer Institute NSW for Fellowship support.

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