Copper(II) complex enhanced chemodynamic therapy through GSH depletion and autophagy flow blockade

Abstract

Three copper(II) complexes C1–C3 were synthesized and fully characterized as chemodynamic therapy (CDT) anticancer agents. C1–C3 showed greater cytotoxicity than their ligands toward SK-OV-3 and T24 cells. Particularly, C2 showed high cytotoxicity toward T24 cells and low cytotoxicity toward normal human HL-7702 and WI-38 cells. Mechanistic studies demonstrated that C2 oxidized GSH to GSSG and produced ˙OH, which induced mitochondrial dysfunction and ER stress, finally leading to apoptosis of T24 cells. In addition, C2 inhibited autophagy by blocking autophagy flow, thereby closing the self-protection pathway of oxidative stress to enhance CDT. Importantly, C2 significantly inhibited T24 tumor growth with 57.1% inhibition in a mouse xenograft model. C2 is a promising lead as a potential CDT anticancer agent.

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Introduction

Although platinum anticancer drugs are commonly used for the treatment of various solid tumors, they lack specificity and selectivity to cancer cells which result in serious side effects.^1–5 Therefore, how to reduce the toxicity and side effects of metal complex drugs, improve their efficacy and overcome drug resistance have been the focus of medicinal chemists.

Compared with normal cells, the concentration of reactive oxygen species (ROS) in cancer cells is higher, and cancer cells are more sensitive to the increase in ROS levels.^6,7 Modulation of ROS levels in tumor cells could enhance efficacy and reduce side effects. Bu et al. took advantage of the tumor microenvironment (TME) of weak acidity, hypoxia, and higher levels of H₂O₂ and glutathione (GSH) and proposed a new treatment strategy called chemodynamic therapy (CDT) by utilizing the Fenton/Fenton-like reaction. CDT uses the Fenton/Fenton-like reaction catalyzed by metal ions (e.g. Fe²⁺, Mn²⁺, Co²⁺, Ti³⁺ and Cu⁺) to convert excess H₂O₂ in tumor tissues into highly cytotoxic ˙OH to induce oxidative stress, which subsequently induces cancer cell apoptosis.^8–12 Due to the TME, CDT exhibits higher specificity of ROS generation in tumor tissues and lower side effects on normal tissues.^13,14 However, overexpression of intracellular glutathione (GSH) in tumor cells, which has a strong scavenging effect on ROS, is an obstacle for CDT.^15–17 Thus, reducing the level of intracellular GSH is desirable to avoid tumor resistance and improve the efficacy of CDT.^18–20 Tai et al. reported a novel bimetallic Cu(II) complex enhancing CDT by oxidizing GSH into glutathione (GSSG).²¹ Copper(II) complexes are expected to reduce GSH concentration and have been used to enhance CDT for cancer treatment. Cancer cells could initiate protective autophagy to remove impaired organelles and toxic proteins involved in ROS generation for detoxification and maintaining homeostasis. Therefore, inhibiting the autophagy of cancer cells to enhance the effect of CDT and induce apoptosis is another effective anticancer strategy.^22–24

The coordination of transition metal ions with bioactive ligands to form complexes is an effective strategy to improve the activity of metal-based anticancer drugs.^25,26 Quinoline is the key core structure of many antitumor agents, and its derivatives are often chosen as ligands to coordinate with metal ions to form anticancer metal complexes.^27–31 Our group has reported many metal complexes with quinoline as the ligand including Cu(II) quinoline complexes that can be used as CDT agents, with remarkable antitumor activity.^32–34 As a continuation of our previous work, we synthesized dichloro-substituted quinoline derivative Cu(II) complexes (C1–C3) as CDT agents. Cu(II) could deplete GSH to produce Cu(I), which further catalyzes the conversion of H₂O₂ into highly cytotoxic ˙OH. At the same time, C2 enhances CDT by inhibiting autophagy flow.

Experimental

Materials

All chemicals were obtained commercially and used without further purification unless otherwise specified. All compounds were initially dissolved in DMF with an initial concentration of 2 mM.

Syntheses of L¹–L³

2-(2-Fluorophenyl)-3-aminoquinoline was obtained according to the steps reported in the references.^34–37 Then 2-(2-fluorophenyl)-3-aminoquinoline and 2-hydroxybenzaladehyde derivatives were dissolved in CH₃OH by continuously stirring at 80 °C for 5–12 h. The product quinolone ligand was obtained by cooling, filtering, and further recrystallizing with CH₃OH.

L¹ (yield: 90%): HRMS: m/z 409.0321 [M − H]⁻. ¹H NMR (400 MHz, DMSO-d₆): δ 12.41 (s, 1H), 9.10 (s, 1H), 8.45 (s, 1H), 8.11 (d, J = 2.0 Hz, 2H), 8.09 (t, J = 1.5 Hz, 2H), 7.90 (s, 1H), 7.83 (ddd, J = 8.4, 6.9, 1.5 Hz, 1H), 7.71 (ddd, J = 8.2, 6.9, 1.2 Hz, 1H), 7.66–7.58 (m, 2H), 7.57 (ddd, J = 9.9, 4.9, 2.1 Hz, 2H), 7.41–7.29 (m, 2H), 7.24 (d, J = 2.8 Hz, 1H). ¹³C NMR (100 MHz, DMSO-d₆): δ 165.52, 159.80 (d, J_C–F = 245.3 Hz), 158.00, 151.90, 146.81, 140.85, 136.91, 132.44, 132.08 (d, J_C–F = 3.3 Hz), 131.81 (d, J_C–F = 8.4 Hz), 130.49, 129.36, 128.62, 128.34, 128.24, 126.92 (d, J_C–F = 15.8 Hz), 125.15 (d, J_C–F = 3.0 Hz), 124.78, 121.05, 119.63, 119.09, 116.07 (d, J_C–F = 21.7 Hz). ¹⁹F NMR (376 MHz, DMSO-d₆): δ −115.04.

L² (yield: 92%): HRMS: m/z 409.0324 [M − H]⁻. ¹H NMR (400 MHz, DMSO-d₆): δ 13.15 (s, 1H), 9.17 (s, 1H), 8.51 (s, 1H), 8.11 (t, J = 7.8 Hz, 2H), 7.84 (ddd, J = 8.4, 6.8, 1.5 Hz, 1H), 7.79–7.68 (m, 3H), 7.69–7.55 (m, 2H), 7.44–7.33 (m, 2H). ¹³C NMR (100 MHz, DMSO-d₆): δ 165.09, 159.77 (d, J_C–F = 245.3 Hz), 155.31, 151.88, 146.86, 141.02, 133.15, 132.07 (d, J_C–F = 3.3 Hz), 131.80 (d, J_C–F = 8.3 Hz), 131.32, 130.55, 129.35, 128.63, 128.27, 128.26, 126.88 (d, J_C–F = 15.7 Hz), 125.14 (d, J_C–F = 3.2 Hz), 124.71, 123.10, 121.89, 121.21, 116.04 (d, J_C–F = 21.7 Hz). ¹⁹F NMR (376 MHz, DMSO-d₆): δ −115.25.

L³ (yield: 87%): HRMS: m/z 409.0323 [M − H]⁻. ¹H NMR (400 MHz, DMSO-d₆): δ 12.41 (s, 1H), 9.10 (s, 1H), 8.46–8.42 (m, 1H), 8.10 (ddd, J = 8.0, 4.5, 1.3 Hz, 2H), 7.89 (s, 1H), 7.82 (ddd, J = 8.4, 6.9, 1.4 Hz, 1H), 7.71 (ddd, J = 8.2, 6.9, 1.2 Hz, 1H), 7.66–7.52 (m, 2H), 7.42–7.29 (m, 2H), 7.22 (s, 1H). ¹³C NMR (100 MHz, DMSO-d₆): δ 163.25, 159.81 (d, J_C–F = 245.3 Hz), 159.48, 152.06, 146.72, 141.85, 136.10, 133.05, 132.07 (d, J_C–F = 3.5 Hz), 131.64 (d, J_C–F = 8.2 Hz), 130.31, 129.33, 128.60, 128.40, 128.14, 127.10 (d, J_C–F = 15.8 Hz), 125.00 (d, J_C–F = 3.3 Hz), 124.11, 121.64, 120.47, 119.13, 115.93 (d, J_C–F = 21.8 Hz). ¹⁹F NMR (376 MHz, DMSO-d₆): δ −115.01.

Syntheses of C1–C3

Cu(OH)₂, ligands, methanol and trichloromethane were mixed in a sample bottle and placed at 65 °C for 72 h. The products C1–C3 were harvested.

C1 (yield: 80%): Anal. Calc. for C₄₄H₂₄CuF₂Cl₄N₄O₂: C 59.78; H 2.74; N 6.34; O 3.62%, found: C 59.81; H 2.77; N 6.30; O 3.59%. HRMS: m/z 919.9566 [M + K]⁺.

C2 (yield: 81%): Anal. Calc. for C₄₄H₂₄CuF₂Cl₄N₄O₂: C 59.78; H 2.74; N 6.34; O 3.62%, found: C 59.76; H 2.71; N 6.32; O 3.66%. HRMS: m/z 881.9975 [M + H]⁺, 903.9767 [M + Na]⁺.

C3 (yield: 82%): Anal. Calc. for C₄₄H₂₄CuF₂Cl₄N₄O₂: C 59.78; H 2.74; N 6.34; O 3.62%, found: C 59.80; H 2.71; N 6.31; O 3.65%. HRMS: m/z 919.9573[M + K]⁺.

¹H NMR analysis of C1–C3 and GSH reaction

Equimolar amounts of copper(II) complexes and GSH were dissolved in deuterated DMSO and D₂O at the volume ratios of 1 : 5 and 60 : 1, and sonicated for 2 h at r.t. to detect the ligand and GSSG on an NMR spectrometer, respectively.

Intracellular ROS detection

T24 cells were treated with L² (12 μM) and C2 (12 μM), respectively. After 8 hours, the cells were stained with DCFH-DA. Finally, the cells were imaged under a fluorescence microscope.

Intracellular ˙OH determination

T24 cells were treated with C2 (12 μM). After 8 hours, the cells were stained with ˙OH detection dye (MitoROS OH580 working solution). The cells were washed with PBS and immediately imaged.

Intracellular ATP determination

T24 cells were treated with C2 (12, 15, and 18 μM) for 48 hours, respectively. The cells were collected and washed and ATP detection was performed following manufacturer's instructions. An ATP kit was purchased from Solarbio (China).

ΔΨ_m detection

T24 cells were treated with C2 (12, 15, and 18 μM), respectively. After 24 hours, the cells were stained with ΔΨ_m detection dye (JC-1 probe). Subsequently, the treated cells were further washed with PBS and the changes of ΔΨ_m were immediately detected.

Apoptosis and cell cycle assay

T24 cells were treated with C2 (12, 15, and 18 μM), respectively. After 24 hours, the cells were stained with Annexin V for 15 min, followed by staining with PI. After incubation, apoptosis was evaluated by flow cytometry.

T24 cells were treated with C2 (12, 15, and 18 μM), respectively. After 24 hours, the treated cells were collected and incubated with ice ethanol at minus 20 °C at least overnight. The cells were then treated with a cell cycle detection dye (PI/RNase staining solution) for half an hour. Finally, the cell cycle distribution was detected.

Protein expression analysis

T24 cells were incubated with C2 (12, 15, and 18 μM) for 48 hours, respectively. The cells were collected by centrifugation and lysed on ice for half an hour with the lysis buffer. The protein was collected by centrifugation and denatured at high temperature. Proteins with different molecular weights were separated, transformed, blocked, and incubated with primary and secondary antibodies, and finally developed.

Cellular autophagosome detection

T24 cells were incubated in a six-well plate overnight, transfected with adenovirus carrying AdPlus-mCherry-GFP-LC3B. Then, the cells were treated with C2 (18 μM) for 48 h. Yellow fluorescent spots were observed under a fluorescence inverted microscope.

Antitumor activity in vivo

Eighteen BALB/c nude mice were assigned to the vehicle group, C2 group and cisplatin group. The administration method was intraperitoneal injection. The C2 group was intraperitoneally (IP) treated with C2 at 20 mg kg⁻¹ (vehicle was 5% DMF and 1% Tween 80 in saline, v/v) daily, the vehicle group was treated with vehicle daily and the cisplatin group was treated with cisplatin at 2 mg kg⁻¹ (saline) every 2 days. Body weight of mice and length (l) and width (w) of T24 tumors were recorded every 2 days. The tumor volume was calculated using the formula: v = lw²/2.

Results and discussion

Synthesis and structural characterization

Three dichloro-substituted quinoline derivatives L¹–L³ were synthesized (Scheme 1). Their structures were determined by NMR spectroscopy and HRMS. As shown in Scheme 2, L¹–L³'s corresponding copper(II) complexes C1–C3 were synthesized by reacting L¹–L³ with Cu(OH)₂ in the presence of methanol and trichloromethane. The structures of C1–C3 were characterized by HRMS, FT-IR spectroscopy and elemental analysis. The crystal structures of C1–C3 were determined by single-crystal X-ray diffraction analysis. Their structures are mononuclear. The Cu(II) center was tetra-coordinated by two nitrogen atoms and two oxygen atoms from the two ligands, respectively (Fig. 1). Further details including the bond lengths and bond angles of C1–C3 are summarized in Tables S1 and S2.†

Scheme 1 Synthesis of L¹–L³. Reagents and conditions: CH₃OH, 80 °C, 5–12 h.

Scheme 2 Synthesis of C1–C3. Reagents and conditions: Cu(OH)₂, CH₃OH and CHCl₃, 65 °C, 72 h.

Fig. 1 The crystal structures of C1–C3. Hydrogen atoms and solvent molecules have been omitted for clarity.

Stability of copper(II) complexes under physiological conditions

The stability of C1–C3 in PBS solution (pH = 7.4) containing 1% DMF was determined by UV-Vis spectroscopy. As shown in Fig. S22,† there were no obvious blue-shift and red-shift in the absorption characteristic peaks of C1–C3 with an increase in time (0, 24, and 48 h), indicating that the copper(II) complexes were stable under physiological conditions for 48 hours at room temperature.

Reaction of C1–C3, GSH and H₂O₂

First, the reaction of C1–C3 with GSH was monitored by ¹H NMR spectroscopy. Because the water solubility of C1–C3 was not high, 60 : 1 and 1 : 5 ratios of DMSO-d₆ and D₂O were used to detect the ligands and GSSG, respectively. When the ratio of DMSO-d₆ to D₂O was 60 : 1, the proton signal of the corresponding ligand was observed in the ¹HNMR spectrum. When the ratio of DMSO-d₆ to D₂O was 1 : 5, compared with the ¹HNMR spectrum of GSH, an obvious signal appeared around 3.15 ppm, belonging to the β-CH₂ group of Cys of GSSG, indicating that GSH was oxidized to GSSG by C1–C3 (Fig. S24–S29†).

The EPR signals of ˙OH in the solution mixture of C1–C3, GSH and H₂O₂ are displayed in Fig. 2. A stronger 1 : 2 : 2 : 1 signal of typical ˙OH could be detected after addition of the ˙OH trapping agent 5,5-dimethyl-1-pyrroline N-oxide (DMPO). However, there is no 1 : 2 : 2 : 1 signal in the solution mixture without copper(II) complexes. Therefore, these three copper(II) complexes could be excellent CDT reagents after GSH depletion.

Fig. 2 EPR spectra for detection of the ˙OH signal. [Cu(II) complex] = 50 μM; [GSH] = 50 μM; [H₂O₂] = 8 mM; and [DMPO] = 100 mM.

In vitro cytotoxicity

The antiproliferative activities of L¹, L², L³, C1, C2 and C3 against human cancer cells Hep-G2, MGC80-3, T24, and SK-OV-3 and human normal cells WI-38 and HL-7702 were investigated by MTT assay. The antiproliferative activities of copper(II) complexes C1–C3 towards T24 and Hep-G2 cells were stronger than those of their ligands, which indicated that ligand coordination with copper(II) enhanced the cytotoxicity in T24 and Hep-G2 cells. The overall in vitro antiproliferative activities of C1–C3 towards T24 cells were higher than those of Hep-G2 cells. C2 showed the strongest cytotoxicity towards T24 cells among the three copper(II) complexes, and the IC₅₀ of C2 was 11.6 ± 1.5 μM. In addition, the cytotoxicity of C2 towards normal cell lines HL-7702 and WI-38 was significantly lower than that for T24 cells. Thus, C2 was selected for further mechanism study (Table 1).

Table 1 IC₅₀ (μM) values of L¹ to L³ and C1 to C3 against various cell lines

	Hep-G2	MGC80-3	T24	SK-OV-3	WI-38	HL-7702
L^1	>20	>20	>20	>20	>20	>20
L^2	>20	18.5 ± 1.2	>20	>20	>20	>20
L^3	>20	17.3 ± 2.8	>20	>20	>20	>20
C1	15.6 ± 2.5	19.0 ± 3.9	15.2 ± 1.3	13.7 ± 0.7	>20	>20
C2	16.9 ± 2.5	16.0 ± 1.9	11.6 ± 1.5	15.3 ± 1.6	>20	19.6 ± 1.7
C3	17.6 ± 3.3	>20	12.2 ± 1.6	13.2 ± 0.9	>20	>20

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Cellular GSH/GSSG ratio and ROS production

To confirm whether C2 oxidized GSH to GSSG in T24 cells, the cellular GSH/GSSG ratio was detected. The GSH/GSSG ratio in C2 (18 μM)-treated T24 cells decreased from 13.7 ± 1.2 to 2.4 ± 0.2 (Fig. 3A). Furthermore, the intracellular ROS production triggered by L² and C2 was investigated with the classic ROS probe DCFH-DA. As shown in Fig. 3B, the green fluorescence intensity of T24 cells treated with C2 was much higher than that of L², indicating that ligands coordination with copper ions increased the production of ROS. Furthermore, the intracellular ˙OH generation triggered by C2 was investigated. The red fluorescence intensity in T24 cells was enhanced after the addition of C2 (Fig. 3C), which indicated that C2 could increase ˙OH generation in T24 cells.

Fig. 3 (A) Intracellular GSH/GSSG ratio after treatment with C2 (18 μM) for 48 h in T24 cells. (B) Intracellular ROS levels after treatment with L² (18 μM) and C2 (18 μM) for 8 h in T24 cells, respectively. (C) Intracellular ˙OH levels after treatment with C2 (18 μM) for 8 h in T24 cells.

Mitochondrial membrane potential detection

The increase in the ROS level can inhibit the electron transfer chain function, resulting in the change of mitochondrial membrane potential (ΔΨ_m) and apoptosis.^38,39 Therefore, the effect of C2 on ΔΨ_m in T24 cells was measured using a JC-1 kit. As shown in Fig. 4A, with the increase in the concentration of C2, ΔΨ_m decreased in a concentration dependent manner. The ΔΨ_m decrease increased from 8.07% (control) to 18.4% (12 μM), 22.5% (15 μM), and 37.5% (18 μM), indicating that C2 induced mitochondrial dysfunction.

Fig. 4 (A) ΔΨ_m decreased in T24 cells by C2 (12, 15 and 18 μM) treatment. (B) Cellular ATP level decreased in T24 cells by C2 (12, 15 and 18 μM) treatment.

Mitochondria are key organelles of ATP generation in cells.⁴⁰ Cancer cells tend to produce more ATP than normal cells, so mitochondria tend to be targeted by anticancer drugs.⁴¹ A dose-dependent decline in cellular ATP levels was observed in T24 cells treated with C2. The ATP level decreased to 71.2 ± 3.7% (12 μM), 44.4 ± 3.5% (15 μM) and 39.1 ± 4.9% (18 μM) of that of control cells (Fig. 4B). These results indicated that C2 induced mitochondrial dysfunction.

ER stress induced by C2

ROS and Ca²⁺ signaling pathways overlap and interact with each other.⁴² ROS affect Ca²⁺ influx and intracellular Ca²⁺ storage, and subsequently Ca²⁺ can increase the production of ROS.⁴³ It has been reported that many drugs induced apoptosis and attenuated the development of tumors through intracellular Ca²⁺ overload.⁴⁴ The effects of C2 on the Ca²⁺ concentration in T24 cells were detected by flow cytometry. Fig. 5A shows that the level of cellular Ca²⁺ increased with the increase of C2 concentration. To further investigate whether the Ca²⁺ concentration was the initiator of ER stress, the expression levels of Phospho-PERK, CHOP and Phospho-eIF2α were detected. After incubation with C2 for 24 h, the levels of Phospho-PERK, CHOP and Phospho-eIF2α in T24 cells were all increased in a dose dependent manner (Fig. 5B). Up-regulation of Phospho-PERK, CHOP and Phospho-eIF2α occurred, which is a typical sign of ER stress. Together, these results indicated that C2 increased Ca²⁺ concentration and induced ER stress.^45–47

Fig. 5 (A) Flow cytometry assay of intracellular Ca²⁺ with Fluo-3AM in T24 cells. (B) The expression of ER stress related proteins (Phospho-PERK, CHOP and Phospho-eIF2α) in T24 cells after treatment with C2.

T24 cell apoptosis induced by C2

When mitochondrial dysfunction and ER stress occur, apoptosis might be initiated. With the increase of C2 concentration, the percentage of apoptotic cells (Q2 + Q3) increased gradually, which was enhanced from 13.16% (12 μM), 18.82% (15 μM) to 54.4% (18 μM) (Fig. 6A).

Fig. 6 (A) Determination of apoptosis in T24 cells after treatment with C2 (12, 15 and 18 μM) for 24 h by flow cytometry. (B) Flow cytometry analysis with PI staining showing the sub-G1 ratio in T24 cells after treatment with C2 (12, 15 and 18 μM) for 24 h.

Subsequently, we investigated the proportion of sub-G1 induced by C2 through PI staining to confirm whether C2 could induce T24 cell apoptosis. Compared to that of vehicle-treated cells (5.62%), the proportion of sub-G1 induced by C2 was 15.13% (12 μM), 17.54% (15 μM), and 39.97% (18 μM), respectively (Fig. 6B). The sub-G1 ratio proportion confirmed that C2 could induce T24 cell apoptosis.

C2 induced mitochondria-mediated apoptosis

A mitochondria-mediated apoptosis pathway is critical for the regulation of apoptosis under various stimuli. To study the apoptotic pathway activated by C2, the expression of proteins involved in the mitochondria-mediated apoptosis pathway was detected. As shown in Fig. 7, C2 could up-regulate Bax protein and down-regulate Bcl-2 protein expression. In addition, with the increase in the concentration of C2, the activation percentage of caspase-9 increased from 1.20% (control) to 6.46% (12 μM), 9.06% (15 μM), and 19.05% (18 μM), and the activation percentage of caspase-3 increased from 3.10% (control) to 8.50% (12 μM), 15.01% (15 μM), and 41.31% (18 μM). Such observations indicated that C2 induced apoptosis in T24 cells via the mitochondria-mediated intrinsic apoptotic pathway.

Fig. 7 C2 induced cell apoptosis via the mitochondria-mediated apoptotic pathway. (A) The expression of Bax and Bcl-2 in T24 cells after treatment with C2 (12, 15, 18 μM) for 48 h. C2 (12, 15, 18 μM) triggered caspase-9 (B) and caspase-3 (C) activities in T24 cells.

C2 induced autophagy inhibition enhanced CDT

The increase of ROS triggers autophagy to protect cancer cells. Under the action of oxidative stress, cancer cells can degrade macromolecules and organelles in damaged cells by activating their own protective autophagy pathway, leading to effective removal of waste in vivo and reduction of the toxicity of reactive oxygen species such as ˙OH towards cancer cells.^23,48 Therefore, inhibiting the autophagy of cancer cells is an effective means to enhance CDT, so that cancer cells cannot “self-detoxify” under oxidative stress and can eventually undergo apoptosis.²² A virus vector expressing the mCherry-GFP-LC3B autophagy double label was used to investigate autophagy flow. As shown in Fig. 8A, the number of yellow fluorescent spots in cells increased after incubation with C2, indicating the increase of autophagosomes in cells. The increase of autophagosomes was further confirmed by western blot analysis. Fig. 8B shows that the expression of p62 and LC3B-II increased after C2 treatment, indicating the increase of autophagosomes in cells. The result indicated that autophagy flow was inhibited by C2. Rapamycin (Rapa) was used to further detect whether autophagy inhibited the production of intracellular ˙OH. As shown in Fig. 8C, Rapa (200 nm) decreased the ROS level induced by C2. Therefore, inhibiting autophagy of cancer cells by C2 enhanced CDT.

Fig. 8 (A) The autophagosomes (orange arrows) induced by C2 were investigated using the mCherry-GFP-LC3B autophagy double label. (B) The expression levels of p62 and LC3B-II in T24 cells after treatment with C2 (12, 15, and 18 μM) for 48 h. (C) Rapa decreased the ROS level induced by C2 (12, 15, and 18 μM).

C2 showed potential anticancer activity in T24 xenograft mice

BALB/c nude mice bearing T24 xenograft tumors were used to investigate the in vivo anticancer activity of C2. Eighteen mice were randomly assigned to the C2 group, cisplatin group, and vehicle control group, with six mice in each group, and they were intraperitoneally injected. Body weight and tumor volume were recorded. As shown in Fig. 9A, the tumor volume in the control group increased to 1430.8 mm³ on the 15th day. In comparison, the tumor volume increased to 598.5 mm³ in the C2 group and 564.7 mm³ in the cisplatin group. It can be seen that C2 and cisplatin inhibited tumor growth, and the tumor growth rates (T/C%) were 41.8% and 37.2%, respectively. After the mice were sacrificed on the 15th day, the tumors were excised (Fig. 9B), weighed and recorded to assess the tumor inhibition rate. Fig. 9C shows that the tumor inhibition rate of C2 (57.1%) was similar to that of cisplatin (62.8%). In addition, C2 showed no obvious effect on the body weight and organs including the spleen, lungs, heart, liver, and kidneys of mice (Fig. 9D and E).

Fig. 9 (A) Effects of vehicle control (5% DMF in saline, v/v, once daily), C2 (20 mg per kg per day) and cisplatin (2 mg per kg per 2 days) on the growth of tumor volume. (B) Tumor weight and tumor inhibition rate. (C) Photographs of tumors excised from mice. (D) Body weight change was monitored every two days. (E) Pathological sections of major organs.

Conclusion

Three quinoline copper(II) complexes have been synthesized as CDT agents. C2 exhibited cytotoxicity against human bladder cancer T24 cells but much lower cytotoxicity against human normal liver HL-7702 cells and lung fibroblast WI-38 cells. C2 decreased the GSH/GSSG ratio and produced ˙OH in T24 cells to induce mitochondrial dysfunction and ER stress, which led to apoptosis and sub-G1 phase arrest. Moreover, C2 blocked autophagy flow to enhance the CDT effect. C2 exhibited a high anticancer effect with a tumor inhibition of 57.1% in the T24 xenograft mouse model. C2 is a promising lead to act as a potential CDT anticancer agent.

Conflicts of interest

The authors declare no competing financial interest.

Acknowledgements

This work was supported by the National Natural Science Foundation of China (Grant 22077022) and Natural Science Foundation of Guangxi Province of China (Grants: AD17129007 and GUIKEZY22096015).

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Footnotes

† Electronic supplementary information (ESI) available. CCDC 2115672–2115674 for complexes C1–C3. For ESI and crystallographic data in CIF or other electronic format see DOI: https://doi.org/10.1039/d2dt04108a

‡ These authors contributed equally to this work.

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