Mitochondria‐Directed Redox Phospholipid Polymers for Cancer Therapy

ABSTRACT

Intracellular redox regulation is increasingly recognized as compartment‐specific, with each organelle maintaining a distinct redox environment. Mitochondria are a major source of reactive oxygen species (ROS), and many cancer cells exhibit elevated basal ROS levels, making mitochondrial redox modulation an attractive strategy for redox‐based cancer therapy. However, efficient mitochondrial delivery of redox‐active agents remains challenging. Small‐molecule redox agents often require hydrophobicity that compromises their water solubility and increases their non‐specific toxicity. Although synthetic polymers improve solubility and biocompatibility, they are predominantly taken up via endocytosis, which often requires additional elements for endosomal escape to achieve efficient cytosolic access and mitochondrial delivery. In this study, we developed mitochondria‐directed phospholipid polymers based on 2‐methacryloyloxyethyl phosphorylcholine (MPC) and redox‐active ferrocene using a simple copolymerization strategy. We leveraged the cell‐penetrating characteristics of amphiphilic MPC copolymers bearing hydrophobic ferrocene units to access the cytosol and introduced a mitochondria‐directed motif by copolymerizing a triphenylphosphonium (TPP)‐containing monomer. The TPP‐bearing polymer exhibited higher in vitro anticancer activity than the non‐directed polymer in CT26 mouse colon cancer cells. Confocal co‐localization analysis revealed that the non‐directed polymer was widely distributed throughout the cells, whereas the TPP‐bearing polymer showed significantly higher mitochondrial co‐localization. The TPP‐bearing polymer also induced higher intracellular ROS levels than the non‐directed polymer. Moreover, intratumoral administration of the TPP‐bearing polymer significantly suppressed CT26 tumor growth in mice. Overall, this study provides a simple design strategy for organelle‐directed redox‐active polymers toward redox‐based cancer therapy.

A mitochondria‐directed redox polymer containing an antioxidant‐oxidizing ferrocene unit and a triphenylphosphonium unit is prepared by a simple copolymerization strategy for cancer treatment. Incorporation of an appropriate amount of the directed unit enhances anticancer activity, elevates intracellular ROS levels, induces apoptosis, and suppresses in vivo tumor growth after intratumoral administration.

Abbreviations

¹H NMR

proton nuclear magnetic resonance

AIBN

2,2′‐azobisisobutyronitrile

AMPA

2,2′‐azobis(2‐methylpropionamidine) dihydrochloride

CCK‐8

Cell Counting Kit‐8

CV

cyclic voltammetry

DLS

dynamic light scattering

DMEM

Dulbecco's modified Eagle's medium

DMSO

dimethyl sulfoxide

D‐PBS

Dulbecco's phosphate‐buffered saline

Fc

ferrocene

Fc⁺

ferrocenium

FITC

fluorescein 5‐isothiocyanate

GPC

gel permeation chromatography

GSH

glutathione

GSSG

glutathione disulfide

HBSS

Hank's balanced salt solution

HPLC

high‐performance liquid chromatography

IC50

half‐maximal inhibitory concentration

Mn

number‐average molecular weight

MPC

2‐methacryloyloxyethyl phosphorylcholine

Mw

weight‐average molecular weight

MWCO

molecular weight cutoff

NAD(P)⁺

oxidized nicotinamide adenine dinucleotide (phosphate)

NAD(P)H

reduced nicotinamide adenine dinucleotide (phosphate)

PI

propidium iodide

PMF

poly(MPC‐co‐VFc)

PMFT

poly(MPC‐co‐VFc‐co‐VTPP)

ROS

reactive oxygen species

SD

standard deviation

SHE

standard hydrogen electrode

THF

tetrahydrofuran

TPP

triphenylphosphonium

UV–Vis

ultraviolet–visible

VFc

vinyl ferrocene

VTPP

4‐vinylbenzyl(triphenyl)phosphonium chloride

1. Introduction

The intracellular redox state governs redox signaling and metabolic pathways, playing a central role in cellular health [1, 2]. In many cancer cells, genetic alterations and metabolic rewiring elevate reactive oxygen species (ROS) levels relative to those in normal cells [3, 4]. Cancer cells tend to upregulate antioxidant systems to maintain redox balance [5]. This dependence on antioxidant defenses creates a vulnerability to redox perturbation that can be therapeutically exploited. Accordingly, approaches increasing intracellular ROS production or decreasing antioxidant capacity show potential to effectively induce cancer cell death [3, 4, 6, 7].

Recently, the intracellular redox state is increasingly considered in a compartment‐specific manner rather than as a uniform whole‐cell property [8, 9, 10, 11, 12]. Redox couples, such as glutathione (GSH)/GSH disulfide (GSSG) and NAD(P)H/NAD(P)⁺, and ROS levels vary across organelles [8, 13, 14, 15]. Therefore, studies have investigated strategies for organelle‐directed modulation of redox balance [16, 17, 18]. Mitochondria are particularly attractive targets, as they are a major endogenous source of ROS and couple redox balance to energy metabolism and apoptotic pathways [19, 20]. Therefore, new strategies for the controlled modulation of mitochondrial redox balance are needed.

Despite growing interest in mitochondria‐directed redox therapy, its effective implementation remains challenging. Many mitochondria‐directed small molecules and redox agents require hydrophobic structures for intracellular delivery, which often result in low aqueous solubility and increase non‐specific interactions with biomolecules, causing unwanted cytotoxicity [21, 22, 23]. Synthetic polymers overcome this issue by providing high aqueous solubility and reduced non‐specific interactions [24, 25, 26]. However, most polymers are internalized mainly via endocytosis, and efficient escape from endosomes, followed by delivery to mitochondria, generally requires additional and often complex molecular designs [27, 28, 29]. Therefore, a platform with a simple molecular design is needed to provide high aqueous solubility and bioinertness and enable cytosolic access to reach mitochondria and modulate mitochondrial redox balance.

We previously developed redox‐active phospholipid polymers for cancer therapy by copolymerizing 2‐methacryloyloxyethyl phosphorylcholine (MPC) with a redox‐active monomer [30, 31, 32, 33]. MPC polymers exhibit high biocompatibility and minimize non‐specific interactions with biomolecules [34, 35]. Furthermore, amphiphilic MPC polymers translocate into the cytosol via simple diffusion, providing intracellular access without relying on endocytosis [36, 37, 38, 39]. Based on this platform, we hypothesized that introducing a mitochondria‐directed unit containing triphenylphosphonium (TPP) [23, 40] could increase mitochondrial localization and influence anticancer activity.

In this study, we designed a mitochondria‐directed redox‐active polymer containing MPC, vinyl ferrocene (VFc), and 4‐vinylbenzyl(triphenyl)phosphonium chloride (VTPP) (Figure 1). We investigated whether the effect of TPP incorporation on the physicochemical properties and anticancer activity of the polymers depended on its content. Based on these results, the polymer showing the highest anticancer activity was selected for subsequent evaluation of mitochondrial co‐localization, intracellular ROS levels, and apoptosis in CT26 mouse colon cancer cells. Its in vivo antitumor activity was also evaluated in a CT26 tumor model.

FIGURE 1.

(a) Chemical structures of poly(MPC‐co‐VFc) (PMF) and poly(MPC‐co‐VFc‐co‐TPP) (PMFT). (b) Schematic illustration of cancer treatment using the mitochondria‐directed redox phospholipid polymer PMFT. Created with https://www.biorender.com.

2. Materials and Methods

2.1. Materials

MPC and VTPP were purchased from Sigma‐Aldrich (St. Louis, MO, USA). VFc, 2,2′‐azobis(2‐methylpropionamidine) dihydrochloride (AMPA), and fluorescein 5‐isothiocyanate (FITC) were purchased from Tokyo Chemical Industry (Tokyo, Japan). Additionally, 2,2′‐azobisisobutyronitrile (AIBN) was purchased from Kanto Chemical (Tokyo, Japan). All other chemicals and solvents were commercially available and used as received without further purification.

2.2. Synthesis of Redox Phospholipid Polymers

Redox phospholipid polymers were synthesized via free‐radical polymerization using AIBN as an initiator. PMF was synthesized as previously reported, and the same batch used in our previous study was employed in this work [33]. To synthesize poly(MPC‐co‐VFc) (PMF), 6.0 mmol of MPC, 4.0 mmol of VFc, and 0.5 mmol of AIBN were dissolved in an ethanol/tetrahydrofuran (THF) mixture (4/1, v/v; 10 mL). To synthesize poly(MPC‐co‐VFc‐co‐TPP)‐1 (PMFT‐1), 5.9 mmol of MPC, 4.0 mmol of VFc, 0.10 mmol of VTPP, and 0.50 mmol of AIBN were dissolved in an ethanol/THF mixture (5/1, v/v; 12 mL) to yield a total monomer amount of 10 mmol. The reaction solutions for PMFT‐5 and PMFT‐10 were prepared using the same procedure with monomer feed ratios listed in Table 1. The reaction solutions were purged with argon gas for 15 min and polymerized at 65°C for 48 h. After polymerization, the polymer solutions were reprecipitated into a diethyl ether/chloroform mixture (9/1, v/v). The precipitates were collected via filtration, dissolved in deionized water, and transferred to a dialysis membrane (Spectra/Por 7; MWCO, 1 kDa; Repligen, Waltham, MA, USA) for dialysis for 4 days. The dialyzed polymer solutions were lyophilized to obtain the polymer powders.

TABLE 1.

Properties of the redox phospholipid polymers synthesized in this study.

Polymers	Monomer unit composition (mol%)		Mw (×103) c	M w /M n c	Hydrodynamic size (nm) d	Redox potential (V vs. SHE) e
	In feed MPC/VFc/VTPP a	In copolymer MPC/VFc/VTPP b
PMF	60/40/0	53/47/0	3.7	1.9	7.1	+0.47
PMFT‐1	59/40/1	47/52/1	4.5	1.5	7.7	+0.47
PMFT‐5	55/40/5	42/52/6	3.9	2.0	8.0	+0.48
PMFT‐10	50/40/10	39/50/11	2.7	2.1	7.1	+0.49

^a MPC, 2‐methacryloyloxyethyl phosphorylcholine; VFc, vinyl ferrocene; VTPP, 4‐vinylbenzyl(triphenyl)phosphonium chloride.

^b Calculated via proton nuclear magnetic resonance (¹H‐NMR) and ultraviolet (UV)–visible (Vis) spectroscopy.

^c Weight‐average (M _w) and number‐average (M _n) molecular weights were measured via gel permeation chromatography (GPC) using polyethylene glycol standards.

^d Determined via dynamic light scattering (DLS).

^e Determined via cyclic voltammetry (CV).

The polymer molar composition was determined via proton nuclear magnetic resonance (¹H NMR) spectroscopy (AVANCE III HD 500 MHz; Bruker, Billerica, MA, USA; Figure S1) and ultraviolet–visible spectroscopy (UV‐1800; Shimadzu, Kyoto, Japan). The molecular weight was evaluated via gel permeation chromatography (HPLC LC‐4000; JASCO, Tokyo, Japan) using a mixture of methanol/water (7/3, v/v) containing 10 mM LiBr for PMF and a mixture of 0.1 M NaNO₃ aqueous solution/acetonitrile (4/1, v/v) for PMFT as the eluent (Figure S2). The particle size was measured via dynamic light scattering (DLS; Zetasizer Nano ZS; Malvern Panalytical, Worcestershire, UK).

2.3. Synthesis of FITC‐Labeled Redox Phospholipid Polymers

FITC‐labeled redox phospholipid polymers (FITC–PMF and FITC–PMFT‐1) were prepared via free‐radical polymerization using AMPA as an initiator. To synthesize AMPA–PMF, 6.0 mmol of MPC, 4.0 mmol of VFc, and 0.50 mmol of AMPA were dissolved in a methanol/THF mixture (4/1, v/v; 10 mL). To synthesize AMPA–PMFT‐1, 5.9 mmol of MPC, 4.0 mmol of VFc, 0.10 mmol of VTPP, and 0.50 mmol of AMPA were dissolved in a methanol/THF mixture (4/1, v/v; 10 mL). The reaction solutions were purged with argon gas for 15 min and polymerized at 65°C for 48 h. After polymerization, the polymer solutions were transferred to a dialysis membrane (Spectra/Por 7; MWCO, 1 kDa) and dialyzed for 4 days. Finally, the dialyzed polymer solutions were lyophilized to obtain the polymer powders.

AMPA–PMF and AMPA–PMFT‐1 were labeled with FITC. To synthesize FITC–PMF, AMPA–PMF (0.10 g) and FITC (0.17 g) were dissolved in an ethanol/dimethyl sulfoxide (DMSO) mixture (1/4, v/v; 10 mL). To synthesize FITC–PMFT‐1, AMPA–PMFT‐1 (0.20 g) and FITC (0.33 g) were dissolved in an ethanol/DMSO mixture (2/3, v/v; 10 mL). The reaction mixtures were stirred at 60°C for 24 h. After the reaction, the polymer solutions were transferred to a dialysis membrane (Spectra/Por 7; MWCO, 1 kDa) and dialyzed for 14 days. The dialyzed polymer solutions were subsequently lyophilized to obtain FITC‐labeled polymer powders.

2.4. Electrochemical Measurements and Oxidation of Redox Phospholipid Polymers

Cyclic voltammetry (CV) was performed using a three‐electrode electrochemical cell. A carbon electrode, Ag/AgCl (saturated KCl), and a platinum wire were used as the working, reference, and counter electrodes, respectively. The polymers were dissolved in Dulbecco's phosphate‐buffered saline (D‐PBS; FUJIFILM Wako Pure Chemical, Osaka, Japan) at 1.0 mg/mL. CV measurements were conducted using a potentiostat (VMP3; BioLogic Science Instruments, Seyssinet‐Pariset, France) at a scan rate of 50 mV/s for five consecutive cycles. The redox potential was determined from the CV curves of the fifth cycle using the following equation:

$$E=\left(E_{\mathrm{ap}}+E_{\mathrm{cp}}\right)/2$$ (1)

where E _ap is the anodic peak potential and E _cp is the cathodic peak potential.

For electrochemical oxidation of the polymers, PMF and PMFT were dissolved in D‐PBS at 10 mg/mL, and an electrode potential of +0.60 V vs. Ag/AgCl (sat. KCl) was applied until the oxidation current reached a constant value. All polymer concentrations are expressed on a total polymer mass basis.

2.5. Cell Culture

CT26 mouse colon carcinoma cells were purchased from the American Type Culture Collection. CT26 cells were seeded in 100‐mm culture dishes (Wuxi NEST Biotechnology, Wuxi, Jiangsu, China) at a density of 1.0 × 10⁶ cells/dish and cultured in 10 mL of RPMI‐1640 medium (FUJIFILM Wako Pure Chemical) supplemented with 10% fetal bovine serum (Corning, Corning, NY, USA) and 1% penicillin–streptomycin solution (FUJIFILM Wako Pure Chemical) at 37°C in a humidified incubator with 5% CO₂.

MB49 mouse bladder carcinoma cells were purchased from EMD Millipore (Temecula, CA, USA). MB49 cells were seeded in 100‐mm culture dishes at a density of 1.0 × 10⁶ cells/dish and cultured in 10 mL of Dulbecco's modified Eagle's medium (DMEM; FUJIFILM Wako Pure Chemical) supplemented with 10% fetal bovine serum and 1% penicillin–streptomycin solution at 37°C in a humidified incubator with 5% CO₂.

NIH3T3 mouse embryonic fibroblasts were purchased from the American Type Culture Collection. NIH3T3 cells were seeded in 100‐mm culture dishes at a density of 3.5 × 10⁵ cells/dish and cultured in 10 mL of DMEM (FUJIFILM Wako Pure Chemical) supplemented with 10% fetal bovine serum and 1% penicillin–streptomycin solution at 37°C in a humidified incubator with 5% CO₂.

Human mesenchymal stem cells (hMSCs) were purchased from Lonza (Basel, Switzerland). hMSCs were seeded in 100‐mm culture dishes at a density of 3.5 × 10⁵ cells/dish and cultured in 10 mL of mesenchymal stem cell growth medium (MSCGM; Lonza) at 37°C in a humidified incubator with 5% CO₂.

2.6. Cell Viability Assay

Cell viability was evaluated using the Cell Counting Kit‐8 (CCK‐8; Dojindo, Kumamoto, Japan). CT26, MB49, NIH3T3, or hMSCs were seeded in a 96‐well plate (Thermo Fisher Scientific, Waltham, MA, USA) at a density of 2.5 × 10³ cells/well in 100 μL of RPMI‐1640 medium (for CT26 cells), DMEM (for MB49 and NIH3T3 cells), or MSCGM (for hMSCs) and incubated for 24 h at 37°C in a humidified incubator with 5% CO₂. Polymer solutions in D‐PBS (10 μL) of each solution were added to each well to achieve the desired final concentration. After 24 h of incubation, the medium was removed, and the cells were washed twice with 100 μL of Hank's balanced salt solution (HBSS; FUJIFILM Wako Pure Chemical). The wells were replenished with 100 μL of fresh culture medium, followed by the addition of 10 μL of CCK‐8 reagent. After incubation at 37°C for 2 h, the absorbance at 450 nm was measured using a microplate reader (iMark Microplate Absorbance Reader; Bio‐Rad Laboratories, Hercules, CA, USA). Medium without cells was used as the blank, and cells treated with 10 μL of D‐PBS were used as the control cells.

2.7. Evaluation of Intracellular Localization

CT26 cells were seeded in the Slide & Chamber 2‐well chambers (WATSON, Tokyo, Japan) at a density of 5.0 × 10⁴ cells/well in 2 mL of RPMI‐1640 medium and incubated for 24 h. The medium was replaced with 1 mL of RPMI‐1640 medium, and FITC–PMF and FITC–PMFT‐1 solutions in D‐PBS (1.0 mg/mL, 1 mL) were added to the culture medium and incubated for 1 h. After removing the medium, the cells were washed twice with HBSS and incubated in 2 mL of HBSS containing MitoTracker Deep Red FM (500 nM; Thermo Fisher Scientific) and Hoechst 33342 (5 μg/mL; Thermo Fisher Scientific) for 15 min. Then, HBSS was removed, and the cells were washed twice with HBSS and fixed with 1 mL of 4% paraformaldehyde solution (Narabyouri Research, Nara, Japan) at room temperature (20°C–25°C) for 15 min. After removing paraformaldehyde, the cells were washed thrice with D‐PBS. Fluoromount (2–3 drops; Cosmo BIO, Tokyo, Japan) was applied to the cells, which were sealed with a coverslip, and the samples were kept at room temperature for 30–45 min. The cells were observed using a confocal laser scanning microscope (LSM 980 with Airyscan 2; Carl Zeiss, Oberkochen, Germany) with 405, 488, and 639‐nm excitation lasers. Co‐localization between the FITC signal and MitoTracker signal was quantified using ImageJ software.

2.8. Evaluation of Intracellular and Mitochondrial ROS Levels

Intracellular and mitochondrial ROS levels were evaluated using the CellROX Green Flow Cytometry Assay Kit (Thermo Fisher Scientific) and MitoBright ROS Deep Red (Dojindo), respectively. CT26 cells were seeded in a 6‐well plate at a density of 1.5 × 10⁵ cells/well in 3.0 mL of RPMI‐1640 medium and incubated for 24 h. Electrochemically oxidized PMF or PMFT‐1 in D‐PBS (10 mg/mL; 0.15 mL) was added to the culture medium and incubated for 1 h. The control group received an equal volume of D‐PBS. After incubation, the medium was removed, and the cells were washed twice with 3.0 mL of HBSS and detached using trypsin. The cell suspension was centrifuged at 100 × g for 5 min, and the cells were stained with CellROX Green and MitoBright ROS Deep Red, respectively, according to the manufacturers' instructions. Fluorescence was measured via flow cytometry (Accuri C6 Plus; BD Biosciences, Franklin Lakes, NJ, USA).

2.9. Apoptosis Assay

Apoptosis was assessed using the FITC Annexin V Apoptosis Detection Kit with PI (BioLegend, San Diego, CA, USA). CT26 cells were seeded in a six‐well plate at a density of 7.5 × 10⁴ cells/well in 3.0 mL of RPMI‐1640 medium and incubated for 24 h. Electrochemically oxidized PMF or PMFT‐1 in D‐PBS (10 mg/mL, 0.15 mL) was added to the culture medium and incubated for 24 h. The control group received an equal volume of D‐PBS. After 24 h, the supernatant was collected and retained. The cells were washed twice with 1.0 mL HBSS, and the washes were collected. Adherent cells were detached using trypsin, and all fractions were pooled and centrifuged at 500 × g for 5 min. The cell pellet was washed twice with HBSS and resuspended in 100 μL of Annexin V Binding Buffer. Annexin V (5 μL) and PI (10 μL) were added, and the cells were incubated at room temperature for 15 min in the dark. Subsequently, 400 μL of Annexin V Binding Buffer was added, and the samples were analyzed via flow cytometry (Accuri C6 Plus).

2.10. Evaluation of Antitumor Activity

All animal experiments were approved by the Animal Experiment Ethics Committee of the School of Engineering, Nagoya University (G250006). Female BALB/c mice (4 weeks old; SLC, Shizuoka, Japan) were subcutaneously injected into the left flank with 3.0 × 10⁵ CT26 cells. When tumor volume reached 30–50 mm³, D‐PBS (control) or electrochemically oxidized PMFT‐1 (10 mg/mL; 100 μL) dissolved in D‐PBS was intratumorally administered once daily on days 0–9. Each group consisted of five mice. Tumor size was measured every 3 days using a caliper, and tumor volume was calculated as follows:

$$\text{Tumor volume}\left({\mathrm{mm}}^3\right)={\left(\text{short diameter}\left[\mathrm{mm}\right]\right)}^2\times \left(\text{long diameter}\left[\mathrm{mm}\right]\right)/2$$ (2)

The mice were euthanized when tumor volume exceeded 2000 mm³.

2.11. Statistical Analyses

The mean values between two groups and among multiple groups were compared using the Welch's t and Tukey's tests, respectively, with EZR (Saitama Medical Center, Jichi Medical University, Japan) [41]. Statistical significance was set at p < 0.05.

3. Results

3.1. Synthesis and Characterization of Mitochondria‐Directed Redox Phospholipid Polymers

To investigate the effect of VTPP content on the physicochemical properties and anticancer activity, a PMFT series was synthesized with different monomer feed ratios (PMFT‐1, PMFT‐5, and PMFT‐10; Table 1). The compositions of the resulting polymers did not markedly deviate from their corresponding feed ratios. Moreover, no clear differences in hydrodynamic size, weight‐average molecular weight (M _w), and dispersity (M _w/M _n) were observed between PMFT and PMF.

The redox activities of the polymers were evaluated via CV (Figure 2). The cyclic voltammograms showed apparent redox potentials of +0.47 V, +0.47 V, +0.48 V, and +0.49 V versus standard hydrogen electrode (SHE) for PMF, PMFT‐1, PMFT‐5, and PMFT‐10, respectively, indicating a gradual positive shift with increasing VTPP content.

FIGURE 2.

Cyclic voltammograms of 1.0 mg/mL PMF and PMFT in Dulbecco's phosphate‐buffered saline (D‐PBS) at a scan rate of 50 mV/s.

3.2. In Vitro Anticancer Activity of Mitochondria‐Directed Redox Phospholipid Polymers

Next, the anticancer activity of the polymers against CT26 cells was evaluated. The electrochemically oxidized polymers decreased cell viability in a concentration‐dependent manner (Figure 3a). Notably, the reduced forms of the polymers were evaluated under the same conditions and did not show a comparable decrease in viability (Figure S3). The lack of anticancer activity of the reduced forms of the polymers supports the redox state‐dependence of their anticancer effects and suggests that VTPP primarily modulates intracellular delivery rather than acting as a cytotoxic moiety.

FIGURE 3.

In vitro anticancer activity. (a) Viability of CT26 mouse colon cancer cells incubated in culture medium supplemented with electrochemically oxidized PMF or PMFT (0.15–1.0 mg/mL) for 24 h. Cell viability was normalized to that of the control (cultured in the absence of polymers). Data are represented as the mean ± standard deviation (SD; n = 3). (b) Half‐maximal inhibitory concentration (IC₅₀) values of PMF and PMFT against CT26 cells calculated from three independent experiments (n = 3). Error bars represent the SD. Asterisks indicate significant differences (*p < 0.05).

Half‐maximal inhibitory concentration (IC₅₀) values were calculated from the viability data of the oxidized polymers (Figure 3b). In the oxidized polymer series, PMFT‐1 exhibited a lower IC₅₀ than PMF, indicating enhanced anticancer activity upon VTPP incorporation. However, a further increase in VTPP content attenuated anticancer activity. PMFT‐5 and PMFT‐10 showed higher IC₅₀ values, and the IC₅₀ of PMFT‐10 approached that of PMF. Based on these results, PMFT‐1 was used for subsequent evaluations of the mitochondria‐directed redox polymers. The oxidized forms of PMF and PMFT‐1 were also evaluated in MB49 bladder carcinoma cells, NIH3T3 fibroblasts, and hMSCs (Figure S4). Both polymers exhibited lower cytotoxicity toward the non‐cancerous NIH3T3 fibroblasts and hMSCs than toward the CT26 and MB49 cancer cells.

3.3. Intracellular Localization of Mitochondria‐Directed Redox Phospholipid Polymers

To evaluate mitochondrial association of the polymers, FITC‐labeled polymers (FITC–PMF and FITC–PMFT‐1) were synthesized, and their intracellular localization was examined via confocal laser scanning microscopy. CT26 cells were incubated with FITC‐labeled polymers (0.50 mg/mL) for 1 h (Figure 4a). FITC–PMF fluorescence was broadly distributed throughout the cells, consistent with the reported intracellular distribution of non‐directed amphiphilic MPC polymers [36]. In contrast, FITC–PMFT‐1 fluorescence was predominantly cytoplasmic and showed greater overlap with the mitochondrial signals. Line‐scan analysis showed that the fluorescence intensity profile of FITC–PMFT‐1 more closely matched the mitochondrial signal than that of FITC–PMF (Figure S5). Additionally, co‐localization was quantified using Pearson's correlation coefficient (Figure 4b). FITC–PMFT‐1 showed a significantly higher coefficient than FITC–PMF, indicating enhanced mitochondrial co‐localization of the VTPP‐containing redox phospholipid polymer.

FIGURE 4.

In vitro subcellular localization of PMF and PMFT‐1. (a) Representative mitochondrial co‐localization images of CT26 cells incubated in the presence of 0.5 mg/mL of fluorescein 5‐isothiocyanate (FITC)–PMF and FITC–PMFT‐1 for 1 h. Scale bar, 5 μm. Mitochondria and nuclei were stained with MitoTracker Deep Red FM and Hoechst 33342, respectively. (b) Pearson's correlation coefficients. Co‐localization of FITC–PMF and FITC–PMFT‐1 with mitochondria was analyzed using the ImageJ software. Data are represented as the mean ± SD (n = 20 cells from two independent experiments). Asterisks indicate significant differences (***p < 0.001).

3.4. Effects of Mitochondria‐Directed Redox Phospholipid Polymers on Intracellular ROS Levels and Apoptosis

We hypothesized that the enhanced anticancer activity of PMFT‐1 is associated with elevated intracellular ROS levels promoted by the incorporation of the mitochondria‐directed VTPP unit. To verify this, intracellular and mitochondrial ROS levels in CT26 cells were evaluated after 1 h of incubation with PMF or PMFT‐1 (Figure 5 and Figure S6). PMFT‐1‐treated cells showed higher intracellular and mitochondrial ROS levels than PMF‐treated cells, consistent with the increased mitochondrial co‐localization of PMFT‐1.

FIGURE 5.

Intracellular reactive oxygen species (ROS) levels in CT26 cells treated with electrochemically oxidized PMF or PMFT‐1. (a) Representative intracellular ROS levels in CT26 cells incubated in medium supplemented with D‐PBS (control; black), 0.5 mg/mL of PMF (gray), or 0.5 mg/mL of PMFT‐1 (green) for 1 h. CT26 cells were stained with CellROX Green and analyzed via flow cytometry. (b) Fold change in mean fluorescence intensity reflecting intracellular ROS levels. Data are represented as the mean ± SD of three independent experiments (n = 3). Asterisks indicate significant differences (*p < 0.05).

To examine whether the increased intracellular ROS levels were associated with apoptotic cell death, an apoptosis assay based on Annexin V and PI co‐staining was performed. Flow cytometry analysis showed that the polymer‐treated groups exhibited higher proportions of early and late apoptotic cells than the control group (Figure 6). Notably, PMFT‐1‐treated cells showed a higher proportion of apoptotic cells (early and late apoptotic cells, 91.6%) than PMF‐treated cells (83.5%). Overall, PMFT‐1 induced apoptosis in CT26 cells more effectively than PMF, consistent with increased mitochondrial co‐localization and higher intracellular ROS levels.

FIGURE 6.

Apoptosis assay of CT26 cells treated with electrochemically oxidized PMF or PMFT‐1. Representative flow cytometry plots of CT26 cells after 24 h of incubation in culture medium supplemented with D‐PBS (control), 0.5 mg/mL of PMF, or 0.5 mg/mL of PMFT‐1. The cells were stained with Annexin V and propidium iodide (PI) and analyzed via flow cytometry.

3.5. In Vivo Antitumor Activity of Mitochondria‐Directed Redox Phospholipid Polymers

As a preliminary in vivo evaluation, CT26 tumor‐bearing BALB/c mice were intratumorally injected with PMFT‐1. The tumors grew progressively in the control group (D‐PBS), and the mean tumor volume was approximately 2000 mm³ on day 21 (Figure S7). In the PMFT‐1‐treated group, the mean tumor volume was approximately 1200 mm³ on day 21, indicating significant suppression of tumor growth compared to that in the control group (Figure S7). Moreover, no marked body‐weight loss was observed in PMFT‐1‐treated mice (Figure S8).

4. Discussion

In this study, we developed mitochondria‐directed redox phospholipid polymers for enhanced anticancer activity. Redox‐active molecules induce oxidative stress via different mechanisms depending on their redox properties. We previously showed that ferrocene‐containing redox polymers oxidize intracellular antioxidants, such as GSH, thereby inducing oxidative stress‐mediated cancer cell death [30, 31]. Although TPP is widely used as a mitochondria‐directed moiety, its contribution is not necessarily determined simply by its presence or absence. The extent of TPP incorporation can influence the physicochemical properties and anticancer activity of the resulting polymers, and the optimal design may depend on the carrier platform [42, 43, 44]. Consistent with this concept, PMFT‐1, which contained 1 mol% VTPP, showed the highest in vitro anticancer activity, whereas higher VTPP content attenuated anticancer activity relative to PMFT‐1. Given that PMF and PMFT‐1 had comparable redox potentials and that PMFT‐5 and PMFT‐10 had slightly more positive potentials, this activity profile is unlikely to be explained by differences in redox potential. These results suggest that the effect of VTPP content on anticancer activity is more likely associated with changes in intracellular delivery or subcellular localization than with changes in the redox potential of the VFc units. Therefore, VTPP content should be optimized, not simply maximized, in this polymer platform.

Subcellular localization is a critical determinant of redox‐active polymer function because ROS generation and antioxidant buffering are spatially organized within cells. In our system, FITC–PMFT‐1 showed higher mitochondrial co‐localization than FITC–PMF, and PMFT‐1 induced higher intracellular and mitochondrial ROS levels and a higher proportion of apoptotic cells than PMF in CT26 cells. Mitochondria are a major intracellular source of ROS, with superoxide (O^2·−) generated primarily by mitochondrial electron transport processes [45]. Mitochondrial GSH is present in the millimolar range and is considered to be broadly similar to cytosolic levels [46]. Therefore, increased mitochondrial co‐localization of PMFT‐1 may perturb local antioxidant buffering near ROS production sites, thereby contributing to elevated intracellular ROS levels and apoptosis.

As a preliminary in vivo evaluation, PMFT‐1 also showed in vivo antitumor activity after intratumoral administration in the CT26 tumor model. The PMFT‐1‐treated group showed significant suppression of tumor growth compared with the D‐PBS control group, and no marked body‐weight loss was observed. These results demonstrate the in vivo antitumor activity of PMFT‐1. However, further comparative in vivo studies are needed to evaluate the contribution of VTPP incorporation to antitumor efficacy. Further studies are also needed to evaluate the mitochondrial localization of PMFT‐1 in vivo and its effect on antitumor activity. Moreover, comprehensive toxicity assessments will be required to evaluate the in vivo safety of the redox phospholipid polymers. For broader therapeutic application, future molecular designs should combine systemic delivery with organelle‐level redox modulation by balancing stability during blood circulation, tumor tissue access, cellular entry, and mitochondrial localization.

Collectively, this study demonstrates that incorporation of an optimized amount of VTPP into ferrocene‐containing MPC‐based redox phospholipid polymers enhances in vitro anticancer activity. PMFT‐1, containing 1 mol% VTPP, showed elevated mitochondrial co‐localization and increased intracellular and mitochondrial ROS levels and apoptosis in CT26 cells, while suppressing tumor growth after intratumoral administration. These findings provide a basis for the development of mitochondria‐directed redox polymer platforms for cancer therapy.

Author Contributions

Yuma Kato: writing – original draft, writing – review and editing, visualization, methodology, investigation, formal analysis, data curation. Yuki Ogawa: writing – review and editing, writing – original draft, methodology, investigation, formal analysis, data curation, funding acquisition. Akira Ito: writing – review and editing, supervision, investigation, methodology, conceptualization. Masahiro Kaneko: conceptualization, methodology, investigation, supervision, funding acquisition, writing – review and editing, writing – original draft, visualization.

Funding

This work was supported by the Japan Society for the Promotion of Science (25K17894), the Japan Science and Technology Agency (JPMJSP2125), the Public Foundation of Chubu Science and Technology Center, and the Toyota Physical and Chemical Research Institute.

Ethics Statement

All animal experiments were approved by the Animal Experiment Ethics Committee of the School of Engineering, Nagoya University (G250006).

Consent

The authors have nothing to report.

Conflicts of Interest

The authors declare no conflicts of interest.

Supporting information

Figure S1: Chemical structure and proton nuclear magnetic resonance (¹H‐NMR) spectrum of poly(MPC‐co‐VFc‐co‐TPP)‐1 (PMFT‐1).

Figure S2: GPC chromatograms of PMFT.

Figure S3: Viability of CT26 mouse colon cancer cells after 24 h of incubation with as‐synthesized (non‐oxidized) PMF and PMFT (0.15–1.0 mg/mL). Cell viability was normalized to that of the untreated control (cultured in the absence of polymers). Data are represented as the mean ± standard deviation (SD; n = 3).

Figure S4: Viability of CT26 mouse colon cancer cells, MB49 mouse bladder cancer cells, NIH3T3 mouse embryonic fibroblasts, and human mesenchymal stem cells (hMSCs) after 24 h of incubation with electrochemically oxidized (a) PMF and (b) PMFT‐1 (0.15–1.0 mg/mL). Cell viability was normalized to that of the untreated control (cultured in the absence of polymers). Data are represented as the mean ± SD (n = 3).

Figure S5: Line‐scan profiles of FITC‐labeled polymers and mitochondria in CT26 cells. Fluorescence intensity profiles of (a) FITC–PMF and MitoTracker and (b) FITC–PMFT‐1 and MitoTracker were obtained along the indicated lines in the confocal fluorescence images. Scale bar, 5 μm.

Figure S6: Mitochondrial reactive oxygen species (MitoROS) levels in CT26 cells treated with electrochemically oxidized PMF or PMFT‐1. CT26 cells were incubated in medium supplemented with D‐PBS (control; black), 0.5 mg/mL of PMF (gray), or 0.5 mg/mL of PMFT‐1 (green) for 1 h, stained with MitoBright ROS Deep Red, and analyzed via flow cytometry.

Figure S7: Time course of tumor volume in subcutaneous CT26 tumor‐bearing mice. D‐PBS (control) or electrochemically oxidized PMFT‐1 (10 mg/mL; 100 μL) was intratumorally injected when the tumor volume reached 30–50 mm³. Daily administration was performed on days 0–9. Tumor size was monitored every 3 days. (a) Individual tumor growth curves for the control and PMFT‐1 groups. Each line represents one mouse (n = 5). (b) Tumor volumes (mean ± SD) in the control (black) and PMFT‐1 (green) groups (n = 5). Asterisks indicate significant differences (*p < 0.05).

Figure S8: Body weight changes in subcutaneous CT26 tumor‐bearing mice. Dulbecco's phosphate‐buffered saline (D‐PBS; control) or electrochemically oxidized PMFT‐1 was intratumorally injected when tumor volume reached 30–50 mm³. Daily administration was performed on days 0–9. (a) Individual body weight curves for the control (left) and PMFT‐1 (right) groups. Each line represents one mouse (n = 5). (b) Body weights (mean ± SD) in the control (black) and PMFT‐1 (green) groups (n = 5).

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.