Poly(l-Ornithine)-Based Polymeric Micelles as pH-Responsive Macromolecular Anticancer Agents

Anticancer peptides and polymers represent an emerging field of tumor treatment and can physically interact with tumor cells to address the problem of multidrug resistance. In the present study, poly(l-ornithine)-b-poly(l-phenylalanine) (PLO-b-PLF) block copolypeptides were prepared and evaluated as macromolecular anticancer agents. Amphiphilic PLO-b-PLF self-assembles into nanosized polymeric micelles in aqueous solution. Cationic PLO-b-PLF micelles interact steadily with the negatively charged surfaces of cancer cells via electrostatic interactions and kill the cancer cells via membrane lysis. To alleviate the cytotoxicity of PLO-b-PLF, 1,2-dicarboxylic-cyclohexene anhydride (DCA) was anchored to the side chains of PLO via an acid-labile β-amide bond to fabricate PLO(DCA)-b-PLF. Anionic PLO(DCA)-b-PLF showed negligible hemolysis and cytotoxicity under neutral physiological conditions but recovered cytotoxicity (anticancer activity) upon charge reversal in the weakly acidic microenvironment of the tumor. PLO-based polypeptides might have potential applications in the emerging field of drug-free tumor treatment.


Introduction
Even though early screening, advances in diagnostics, and improved therapeutic regimens have led to a decline in cancer mortality, malignancies remain the major cause of death [1]. Chemotherapy is one of the most important and indispensable methods for treating malignant tumors. However, chemotherapy is usually unsatisfactory due to insufficient drug accumulation in tumor tissues, poor aqueous solubility, drug resistance, severe off-target toxicity, and a high probability of metastasis [2][3][4][5]. To address these problems, a wide variety of polymeric micelles have been extensively explored to deliver anticancer drugs to the tumor site by embedding a drug in the hydrophobic core of the polymeric micelles or by conjugating the drug at the distal end, thus increasing the circulation time, improving the accumulation in tumor tissue, and reducing the toxicity of the drug itself [6][7][8][9][10]. Although significant advances in drug delivery systems have been achieved, there remain many challenges, such as burst release and potential off-target toxicity of small-molecule drugs, as well as their susceptibility to developing drug resistance [11,12]. To this end, new anticancer agents that afford high selectivity toward cancer cells and overcome multidrug resistance are in critical demand.
Host defense peptides are short cationic peptides which are widely found in nature and play an important role in immediate nonspecific defenses against various microbes, including bacteria, fungi, protozoa, and viruses [13]. Over the past few decades, inspired by host defense peptides, synthetic antimicrobial peptides (AMPs) have been extensively studied to combat bacteria via a membrane-lytic mechanism [14][15][16][17][18][19]. Cationic and amphiphilic NCA) and L-phenylalanine N-carboxyanhydride (LF-NCA) to produce poly(δ-benzyloxycarbonyl-L-ornithine)-b-poly(L-phenylalanine) (PZLO-b-PLF), followed by the deprotection of benzyloxycarbonyl groups of PZLO according to our previous report [36] to produce the final product PLO-b-PLF. PLO-b-PLF was modified with DCA to prepare PLO(DCA)-b-PLF. The detailed methods of synthesis and characterization are described in the Supplementary Materials section. Scheme 1. Synthetic route of PLO-based polypeptides.

Preparation and Characterization of Polymeric Micelles
PLO-b-PLF and PLO(DCA)-b-PLF were dissolved in PBS (0.01 M, pH 7.4) at a concentration of 1 mg/mL, followed by ultrasonication for 5 min. The particle size, polydispersity, and zeta potential of the polymeric micelles were determined by dynamic light scattering (DTS Zetasizer Nano, Malvern Instruments, Worcestershire, UK). The measurements were carried out for 3 runs per sample, and the results are presented as the mean ± standard deviation. The critical micelle concentration (CMC) of PLO-b-PLF in PBS (pH 7.4) or PLO(DCA)-b-PLF in bicarbonate buffer (pH 9.2) was determined by fluorescence spectroscopy using pyrene as a probe [34]. Briefly, polypeptide solutions with varying concentrations in the range of 0.015-2000 μg/mL were incubated with pyrene (6.16 × 10 −7 M) overnight at room temperature in the dark. The excitation spectra of these solutions were scanned from 300 to 360 nm at an emission wavelength of 395 nm using a fluorescence spectrometer (Horiba FluoroMax, Kyoto, Japan). The intensity ratios of from I339.0 to I334.0 were drawn as a function of the logarithm of polymer concentrations. Several drops of the polymeric micellar solution were placed on a carbon-coated 200 mesh copper grid and kept overnight at room temperature. The morphology of the polymeric micelles was then examined on a transmission electron microscope (TEM, JEM-F200, Tokyo, Japan) with an acceleration voltage of 200 kV. Scheme 1. Synthetic route of PLO-based polypeptides.

Preparation and Characterization of Polymeric Micelles
PLO-b-PLF and PLO(DCA)-b-PLF were dissolved in PBS (0.01 M, pH 7.4) at a concentration of 1 mg/mL, followed by ultrasonication for 5 min. The particle size, polydispersity, and zeta potential of the polymeric micelles were determined by dynamic light scattering (DTS Zetasizer Nano, Malvern Instruments, Worcestershire, UK). The measurements were carried out for 3 runs per sample, and the results are presented as the mean ± standard deviation. The critical micelle concentration (CMC) of PLO-b-PLF in PBS (pH 7.4) or PLO(DCA)b-PLF in bicarbonate buffer (pH 9.2) was determined by fluorescence spectroscopy using pyrene as a probe [34]. Briefly, polypeptide solutions with varying concentrations in the range of 0.015-2000 µg/mL were incubated with pyrene (6.16 × 10 −7 M) overnight at room temperature in the dark. The excitation spectra of these solutions were scanned from 300 to 360 nm at an emission wavelength of 395 nm using a fluorescence spectrometer (Horiba FluoroMax, Kyoto, Japan). The intensity ratios of from I 339.0 to I 334.0 were drawn as a function of the logarithm of polymer concentrations. Several drops of the polymeric micellar solution were placed on a carbon-coated 200 mesh copper grid and kept overnight at room temperature. The morphology of the polymeric micelles was then examined on a transmission electron microscope (TEM, JEM-F200, Tokyo, Japan) with an acceleration voltage of 200 kV.

Cell Viability Assay
The cytotoxicity of PLO-b-PLF and PLO(DCA)-b-PLF was evaluated by an alamarBlue assay [42]. All cells were seeded onto 96-well plates at a density of 6 × 10 3 cells in 100 µL of DMEM with 10% fetal bovine serum per well. After culturing for 24 h, the medium was replaced with fresh complete medium containing different concentrations of polypeptide ranging from 500 to 1.0 µg/mL. Wells without polypeptide treatment and without cells were set as the positive control and negative control, respectively. After incubation for 24 h, the medium was replaced with fresh complete medium containing 10% alamarBlue solution. Upon additional incubation for 2.5 h, the fluorescence intensity of each well was measured on an Infinite M200 microplate reader (Tecan, Zurich, Switzerland) at an excitation wavelength of 555 nm and emission wavelength of 590 nm. Cell viability was calculated by the following formula: Cell viability (%) = [(Fluorescence polypeptide − Fluorescence negative control )/(Fluorescence positive control − Fluorescence negative control )] × 100%

Hemolysis Assay
Fresh rat red blood cells were washed three times by suspending cells in PBS (pH 7.4) and then centrifuged at 3500 rpm for 10 min at 4 • C. The supernatant was removed, and the red blood cells were suspended in PBS (5%, v/v). Then, 50 µL of red blood cell suspension was added to 50 µL of PBS solution containing PLO-b-PLF or PLO(DCA)-b-PLF at various concentrations ranging from 62 to 8000 µg/mL in a 96-well microplate. The mixture was incubated at 37 • C for 1 h. The plate was then centrifuged at 3500 rpm for 10 min. Aliquots (30 µL) of the supernatant were transferred into the well of a 96-well microplate containing 70 µL PBS, and the absorbance was measured at 540 nm using an Infinite M200 microplate reader (Tecan, Zurich, Switzerland). The untreated blood cell suspension in PBS was used as the negative control, and a solution containing red blood cells lysed with 2% Triton X-100 was employed as the positive control. Each test was performed in three replicates. The percentage of hemolysis was calculated by the following formula:

Dead/Live Cell Staining
HepG2 cells were seeded onto 6-well plates at a density of 1.2 × 10 5 cells per well and cultured in 2 mL of complete DMEM at 37 • C for 24 h. Then, fresh complete DMEM containing predetermined concentrations of PLO(DCA)-b-PLF (0 × IC 50 , 0.5 × IC 50 , IC 50 , and 2 × IC 50 ) at pH 6.5 and 100 µg/mL PLO(DCA)-b-PLF at pH 7.4 was added and incubated for 24 h at 37 • C. The cells were washed with PBS (pH 7.4) three times and subsequently costained with calcein acetoxymethyl ester (calcein AM, 6 µM) and propidium iodide (PI, 2 µM) for 15 min at 37 • C. Finally, the cells were washed with PBS twice and imaged by fluorescence microscopy (Observer A1, Zeiss Merlin, Baden-Wuerttemberg, Germany). The excitation and emission wavelengths for calcein AM were 488 nm and 520 nm, respectively, while the excitation and emission wavelengths for PI were 530 nm and 620 nm, respectively.

Zeta Potential Measurement
HepG2 and HK-2 cells were plated in 6-well plates (5 × 10 5 cells/well) and cultured in complete DMEM for 24 h. The cells were harvested and resuspended in complete DMEM at pH 7.4 containing predetermined concentrations of PLO-b-PLF or PLO(DCA)-b-PLF. After incubation for 30 min at 37 • C, the cells were centrifuged and resuspended in 1 mL of H 2 O. The zeta potential of the cells was measured using dynamic light scattering (DTS Zetasizer Nano, Malvern Instruments, Worcestershire, UK). Measurements were carried out at 3 runs per sample, and the results are presented as the mean ± standard deviation.

Lactate Dehydrogenase (LDH) Leakage Assay
HepG2 cells were seeded in a 96-well plate at a density of 6 × 10 3 cells per well in 100 µL of complete DMEM and cultured for 24 h. The cells were then incubated with different concentrations of PLO-b-PLF in fresh complete medium for 45 min. Untreated cells were employed as a negative control for background LDH release, while cells treated with lysis buffer were set as the positive control for maximal LDH release. Subsequently, the 96-well plate was centrifuged at 3000 rpm for 5 min, and LDH release was determined according to the protocol provided by the supplier. Briefly, 50 µL of supernatant was incubated with 50 µL of working solution for 30 min, followed by the addition of 50 µL of stop solution. Absorbance at 490 nm was recorded on an Infinite M200 microplate reader (Tecan, Zurich, Switzerland). The percentage of LDH release was calculated by the following equation: LDH release (%) = [(absorbance sample − absorbance negative control )/(absorbance positive control − absorbance negative control )] × 100%

Flow Cytometry Study
HepG2 cells were seeded onto 6-well plates at a density of 1.5 × 10 5 cells per well and cultured in 2 mL of complete medium at 37 • C for 24 h. Then, the medium was replaced with fresh complete medium containing predetermined concentrations (0 × IC 50 , 0.25 × IC 50 , 0.5 × IC 50 , IC 50 , and 2 × IC 50 ) of PLO-b-PLF. After incubation for 60 min, the cells were washed with PBS, detached with trypsin, and centrifuged to discard the supernatant. Next, the harvested cells were subsequently suspended in the binding buffer and stained with an Annexin V-FITC Apoptosis Detection kit according to the protocol provided by the supplier. Finally, the Annexin V-FITC/PI-labeled cells were subjected to measurement on a BD Accuri™ C6 Flow Cytometer (Becton, Dickinson, and Company, Franklin Lakes, NJ, USA).

Morphological Visualization of Cancer Cells by Scanning Electron Microscopy (SEM)
HepG2 cells were seeded at a density of 1.2 × 10 5 /well on 10 mm × 10 mm sterilized coverslips in a 6-well plate and cultured in complete medium for 24 h, followed by incubation with predetermined concentrations (0 × IC 50 , 0.25 × IC 50 , 0.5 × IC 50 , IC 50 , 2 × IC 50 , and 4 × IC 50 ) of PLO-b-PLF in fresh complete medium for 60 min at 37 • C. The cells were then washed with PBS twice and fixed with 2.5% glutaraldehyde solution at 4 • C overnight. Subsequently, the medium was removed, and the cells were gradually dehydrated by serial incubation in 30%, 50%, 70%, 85%, 95%, and 100% ethanol solutions. Finally, the cells on coverslips were visualized by SEM (Gemini 300, Zeiss Merlin, Baden-Wuerttemberg, Germany).

Confocal Laser Scanning Microscopy (CLSM) Study
HepG2 cells were seeded on sterilized coverslips in a 6-well plate at a density of 1.2 × 10 5 cells per well in 2.0 mL complete medium and cultured for 24 h, followed by incubation with predetermined concentrations (0 × IC 50 , 0.5 × IC 50 , IC 50 , and 2 × IC 50 ) of PLO-b-PLF in fresh complete medium for 30 min at 37 • C. The cells were then rinsed with PBS three times and fixed with 4% paraformaldehyde for 20 min. Subsequently, the cells were washed with PBS three times and then sequentially incubated with DiO (2 µM) for 15 min and Hoechst (10 µg/mL, 1 mL) for 5 min to stain the cell membranes and cell nuclei, respectively. After washing with PBS three times, the coverslips were observed with a CLSM microscope (Carl Zeiss, LSM 800, Baden-Wuerttemberg, Germany).

In Vitro Cancer Cell Migration Assay
HepG2 cells were seeded in 6-well plates at a density of 1.5 × 10 5 cells per well. After incubation for 24 h at 37 • C, the plate surface was scratched with a 200 µL pipette tip to draw a gap of uniform width. The medium was replaced with fresh medium without fetal bovine serum and containing different concentrations (0.25 × IC 50 , 0.5 × IC 50 , IC 50 , and 2 × IC 50 ) of PLO(DCA)-b-PLF at pH 6.5 or 100 µg/mL PLO(DCA)-b-PLF at pH 7.4. The cells were then incubated at 37 • C. Images of the gaps were taken using a bright field microscope (Observer A1, Zeiss Merlin, Baden-Wuerttemberg, Germany) at 0 h and 24 h after scratching.

Synthesis and Characterization of PLO-b-PLF and PLO(DCA)-b-PLF
The block polymer PLO-b-PLF consists of poly(L-ornithine) as the hydrophilic block and poly(L-phenylalanine) as the hydrophobic segment. The synthetic route of PLO-b-PLF is depicted in Scheme 1, and the detailed procedures are described in the Supplementary Materials section. In the present study, the degree of polymerization (DP) of PLO was fixed at 30, while the DPs of PLF were set at 0, 4, 8, and 12. Hexylamine was employed to initiate the sequential ring-opening polymerization of ZLO-NCA and LF-NCA to produce PZLO-b-PLF (i.e., PZLO 30 (PZ1), PZLO 30 -b-PLF 4 (PZ2), PZLO 30 -b-PLF 8 (PZ3), and PZLO 30b-PLF 12 (PZ4)), followed by the deprotection of the benzyloxycarbonyl groups of PZLO according to our previous report [36] to produce the final products PLO 30 Figure S1, and the calculated results are listed in Table S1. Based on the 1 H NMR spectra, the DPs of PZLO and PLF in PZ1, PZ2, PZ3, and PZ4 were estimated to be 30.4 and 0.0, 30.4 and 4.09, 29.2 and 7.91, and 29.1 and 11.3, respectively ( Table S1). All of the calculated DP values are very close to the respective feeding ratios. The unimodal GPC peaks and their narrow distributions in Figure S1 indicate that the synthesis of PZLO-b-PLF proceeded in a controlled manner. Due to the self-assembly of PLO-b-PLF in aqueous solution, the characterization of PLO-b-PLF with 1 H NMR and GPC is difficult. For example, the GPC chromatograms of PLO-b-PLF show predominant peaks, exhibiting very large molecular weights, which can be ascribed to the self-assembled polymeric micelles ( Figure S2B). Consistently, the phenyl groups demonstrated very weak signals in the 1 H NMR spectrum of PLO-b-PLF due to the aggregation of hydrophobic PLF blocks ( Figure S2A). However, it should be noted that the deprotection procedures also proceeded in a controlled manner according to our previous report [43,44].
To reduce cytotoxicity, the cationic PLO-b-PLF was modified with DCA to prepare anionic PLO(DCA)-b-PLF, i.e., PLO 30 Figure S2A. The shift of the proton peak of g from 2.8 ppm to 3.1 ppm and the appearance of proton peak h at approximately 2.0 ppm indicated the successful modification of the DCA.

Characterization of PLO-b-PLF and PLO(DCA)-b-PLF Micelles
The amphiphilic block copolypeptides PLO-b-PLF and PLO(DCA)-b-PLF are expected to self-assemble into polymeric micelles in aqueous solution. The aggregation behavior, i.e., CMC, was investigated by fluorescence spectroscopy using pyrene as the probe according to a previously reported method [35]. The CMCs of cationic P2, P3, and P4 were determined to be 179.5 µg/mL, 51.3 µg/mL, and 28.8 µg/mL, while the CMCs of anionic PD2, PD3, and PD4 were calculated to be 162.2 µg/mL, 40.7 µg/mL, and 22.4 µg/mL, respectively ( Figure S3). An elongation of the hydrophobic PLF block led to a prominent decrease in the CMC values, and the small CMC values indicated that the polymeric micelles, self-assembled from PLO-b-PLF and PLO(DCA)-b-PLF (especially P4 and PD4), had high thermodynamic stability even when diluted in the blood stream.
Dynamic light scattering was employed to investigate the sizes and zeta potentials of polymeric micelles. As shown in Figure 1a, the hydrodynamic diameters of the polymeric micelles ranged from 52 nm to 113 nm. Elongation of the PLF block increases the particle size, whereas DCA modification slightly decreases the micellar diameter. The zeta potentials of the P2, P3, and P4 polymeric micelles are positive, ranging between 14.6 mV and 20.4 mV (Figure 1b), which can be ascribed to the protonated state of amino groups along the L-ornithine side chains under physiological conditions. Modification with DCA converted  (Figure 1b), which is indicative of the success of the DCA modification. The morphology of the polymeric micelles was examined by TEM ( Figure S4). The polymers P4 and PD4 adopted a spherical morphology with a relatively uniform particle size [35]. The smaller sizes of polymeric micelles obtained from the TEM measurements might be ascribed to the collapse and contraction of micellar structures during the process of sample preparation. polymeric micelles. As shown in Figure 1a, the hydrodynamic diameters of the polymer micelles ranged from 52 nm to 113 nm. Elongation of the PLF block increases the partic size, whereas DCA modification slightly decreases the micellar diameter. The zeta pote tials of the P2, P3, and P4 polymeric micelles are positive, ranging between 14.6 mV an 20.4 mV (Figure 1b), which can be ascribed to the protonated state of amino groups alon the L-ornithine side chains under physiological conditions. Modification with DCA co verted the charging state from positive to negative (Figure 1b), which is indicative of t success of the DCA modification. The morphology of the polymeric micelles was exam ined by TEM ( Figure S4). The polymers P4 and PD4 adopted a spherical morphology wi a relatively uniform particle size [35]. The smaller sizes of polymeric micelles obtaine from the TEM measurements might be ascribed to the collapse and contraction of micell structures during the process of sample preparation.

Hydrolysis of Acid-Labile PLO(DCA)-b-PLF
In PLO(DCA)-b-PLF, the DCA groups were anchored to the side chains of PLO wi acid-labile β-carboxylic amide linkages. It is expected that the β-carboxylic amide wou remain relatively stable under physiological conditions (pH 7.4), enabling a negative-t positive charge reversal to occur upon the hydrolysis of the β-carboxylic amide under t slightly acidic conditions of the tumor microenvironment (pH 6.5-6.8). The hydrolysis k netics of PLO(DCA)-b-PLF were investigated by 1 H NMR spectra and zeta potential mea urements. The evolution of the 1 H NMR spectra of PLO(DCA)-b-PLF as a function of i cubation time at pH 6.5 is shown in Figure S5B, in which the methylene peak of the orn thine side chains shifts from 3.1 ppm to 2.8 ppm as the β-carboxylic amide hydrolyze Two peaks (Ha and Hb) were integrated to obtain the hydrolysis kinetics of PLO(DCA)-

Hydrolysis of Acid-Labile PLO(DCA)-b-PLF
In PLO(DCA)-b-PLF, the DCA groups were anchored to the side chains of PLO with acid-labile β-carboxylic amide linkages. It is expected that the β-carboxylic amide would remain relatively stable under physiological conditions (pH 7.4), enabling a negative-topositive charge reversal to occur upon the hydrolysis of the β-carboxylic amide under the slightly acidic conditions of the tumor microenvironment (pH 6.5-6.8). The hydrolysis kinetics of PLO(DCA)-b-PLF were investigated by 1 H NMR spectra and zeta potential measurements. The evolution of the 1 H NMR spectra of PLO(DCA)-b-PLF as a function of incubation time at pH 6.5 is shown in Figure S5B, in which the methylene peak of the ornithine side chains shifts from 3.1 ppm to 2.8 ppm as the β-carboxylic amide hydrolyzes. Two peaks (H a and H b ) were integrated to obtain the hydrolysis kinetics of PLO(DCA)b-PLF under different pH values ( Figure S5C). The results showed that the β-carboxylic amide in PLO(DCA)-b-PLF is easily hydrolyzed at pH 6.5 but remains relatively stable at pH 7.4, thus realizing the negative-to-positive charge conversion required for antitumor activity. Meanwhile, the zeta potentials of PD4 as a function of incubation time at pH 6.5 and 7.4 were measured (Figure 2). The surface charge of PD4 was converted from negative to positive upon incubation at pH 6.5 after approximately 6.5 h, whereas PD4 remained negatively charged even after 48 h of incubation at pH 7.4. These results indicate that negative-to-positive charge reversal of PLO(DCA)-b-PLF could be achieved by the pHinduced hydrolysis of β-carboxylic amide under the slightly acidic conditions of the tumor microenvironment. and 7.4 were measured (Figure 2). The surface charge of PD4 was converted from negative to positive upon incubation at pH 6.5 after approximately 6.5 h, whereas PD4 remained negatively charged even after 48 h of incubation at pH 7.4. These results indicate that negative-to-positive charge reversal of PLO(DCA)-b-PLF could be achieved by the pH-induced hydrolysis of β-carboxylic amide under the slightly acidic conditions of the tumor microenvironment.

Cytotoxicity (Antitumor Activity) Assays
The cytotoxicity of PLO-b-PLF block copolymers was evaluated against six cancer cell lines by alamarBlue assay, and the corresponding IC50 values are listed in Table 1. PLO-b-PLF exhibited a broad spectrum of anticancer activity against all cancer cells, including MCF-7/ADR cells, with IC50 values ranging from 5.03 to 20.3 μg/mL. Notably, the IC50 value of P1 (in the absence of hydrophobic block and micelle formation) was close to those of P2, P3, and P4, bearing hydrophobic PLF segments, indicating that the presence of a hydrophobic segment and the increase in charge density as a result of micelle formation did not significantly improve the anticancer activity of PLO-b-PLF. Meanwhile, we measured the zeta potential of cancer cells, which resided at a very narrow range at about −35 mV (HepG2, −35.0 mV; MCF-7, −33.5 mV; Hela, −35.1 mV; A547, −35.8 mV). No obvious correlation between the IC50 values and zeta potentials of the cells was observed. The IC50 value might relate to the zeta potential of the cell as well as other factors, such as the cell membrane structure. Since P4 had a lower CMC value and, thus, higher hydrodynamic stability that might exhibit better resistance to the dilution conditions in vivo, it was selected as the representative polymer for the following studies. To alleviate the cytotoxicity of PD4 toward normal cells, anionic and charge-reversal PLO(DCA)-b-PLF was synthesized. The cytotoxicity of PD4 on representative HK-2 and

Cytotoxicity (Antitumor Activity) Assays
The cytotoxicity of PLO-b-PLF block copolymers was evaluated against six cancer cell lines by alamarBlue assay, and the corresponding IC 50 values are listed in Table 1. PLOb-PLF exhibited a broad spectrum of anticancer activity against all cancer cells, including MCF-7/ADR cells, with IC 50 values ranging from 5.03 to 20.3 µg/mL. Notably, the IC 50 value of P1 (in the absence of hydrophobic block and micelle formation) was close to those of P2, P3, and P4, bearing hydrophobic PLF segments, indicating that the presence of a hydrophobic segment and the increase in charge density as a result of micelle formation did not significantly improve the anticancer activity of PLO-b-PLF. Meanwhile, we measured the zeta potential of cancer cells, which resided at a very narrow range at about −35 mV (HepG2, −35.0 mV; MCF-7, −33.5 mV; Hela, −35.1 mV; A547, −35.8 mV). No obvious correlation between the IC 50 values and zeta potentials of the cells was observed. The IC 50 value might relate to the zeta potential of the cell as well as other factors, such as the cell membrane structure. Since P4 had a lower CMC value and, thus, higher hydrodynamic stability that might exhibit better resistance to the dilution conditions in vivo, it was selected as the representative polymer for the following studies. To alleviate the cytotoxicity of PD4 toward normal cells, anionic and charge-reversal PLO(DCA)-b-PLF was synthesized. The cytotoxicity of PD4 on representative HK-2 and LO2 cells at pH 7.4 is shown in Figure S6A,B. The survival rates of HK-2 and LO2 cells were above 50%, even when treated at a dose of 500 µg/mL PD4 for 24 h, which is indicative of a negligible toxicity of PD4 under physiological conditions. Under the slightly acidic conditions of tumors, the hydrolysis of DCA could recover the cationic state and thus afford anticancer activity. The cytotoxicity of DCA-modified PD4 on various cancer cells after incubation for 24 h at pH 6.5 was then determined ( Figure S7). The corresponding IC 50 values of PD4 on HepG2, A549, MCF-7, MCF-7/ADR, BT474, and HeLa cells were 20.1 µg/mL, 28.7 µg/mL, 29.0 µg/mL, 12.9 µg/mL, 37.2 µg/mL, and 48.4 µg/mL, respectively. These results indicate that PD4, modified with acid-sensitive DCA, could effectively kill cancer cells in weakly acidic cancer tissue without causing harm to normal cells. The anticancer effect of PD4 was directly demonstrated through a live/dead cell costaining measurement. HepG2 cells were incubated in the presence of PD4 under different pH values (6.5 or 7.4) and then stained with PI (red, dead cells) and calcein AM (green, live cells). As shown in Figure 3, few PI-positive cells were observed when treated with 100 µg/mL PD4 at pH 7.4. However, when the concentration of PD4 increased from 0.5 × IC 50 to 2 × IC 50 at pH 6.5, stronger PI staining was detected, indicating that more HepG2 cells were killed with increasing concentrations of PD4. The live/dead staining results are consistent with the alamarBlue assay and further demonstrate that the noncytotoxic PD4 could experience charge reversal at pH 6.5 to exert anticancer activity. acidic conditions of tumors, the hydrolysis of DCA could recover the cationic state and thus afford anticancer activity. The cytotoxicity of DCA-modified PD4 on various cancer cells after incubation for 24 h at pH 6.5 was then determined ( Figure S7). The corresponding IC50 values of PD4 on HepG2, A549, MCF-7, MCF-7/ADR, BT474, and HeLa cells were 20.1 μg/mL, 28.7 μg/mL, 29.0 μg/mL, 12.9 μg/mL, 37.2 μg/mL, and 48.4 μg/mL, respectively. These results indicate that PD4, modified with acid-sensitive DCA, could effectively kill cancer cells in weakly acidic cancer tissue without causing harm to normal cells.
The anticancer effect of PD4 was directly demonstrated through a live/dead cell costaining measurement. HepG2 cells were incubated in the presence of PD4 under different pH values (6.5 or 7.4) and then stained with PI (red, dead cells) and calcein AM (green, live cells). As shown in Figure 3, few PI-positive cells were observed when treated with 100 μg/mL PD4 at pH 7.4. However, when the concentration of PD4 increased from 0.5 × IC50 to 2 × IC50 at pH 6.5, stronger PI staining was detected, indicating that more HepG2 cells were killed with increasing concentrations of PD4. The live/dead staining results are consistent with the alamarBlue assay and further demonstrate that the noncytotoxic PD4 could experience charge reversal at pH 6.5 to exert anticancer activity.

Hemolysis Assay
The hemolytic activities of P4 and PD4 were determined against rat red blood cells, and the corresponding hemolytic percentage as a function of polypeptide concentrations is shown in Figure S8. For P4, hemolysis increased with increasing concentrations of P4. However, the hemolysis caused by PD4 remained at approximately zero even when the concentration of PD4 was as high as 4000 μg/mL. These results show that DCA modification can greatly reduce hemolytic toxicity, largely owing to the negatively charged state of PD4.

Hemolysis Assay
The hemolytic activities of P4 and PD4 were determined against rat red blood cells, and the corresponding hemolytic percentage as a function of polypeptide concentrations is shown in Figure S8. For P4, hemolysis increased with increasing concentrations of P4. However, the hemolysis caused by PD4 remained at approximately zero even when the concentration of PD4 was as high as 4000 µg/mL. These results show that DCA modification can greatly reduce hemolytic toxicity, largely owing to the negatively charged state of PD4.

Electrostatic Binding of PLO onto the Surface of Cancer Cells
The binding of cationic PLO-based polypeptides onto the surface of anionic cancer cells was evaluated by measuring the change in zeta potential of HepG2 cells as a function of different concentrations of P1 and P4 [45]. HepG2 cells were incubated with P1 or P4 for 30 min, centrifuged, and then resuspended prior to zeta potential measurements. The surface charge of HepG2 cells evolved from negative to positive with increasing concentrations of P1 and P4 ( Figure 4A), indicating that cationic PLO-based polypeptides bind to the surface of HepG2 cells via electrostatic interactions. It is interesting to note that, compared to P4, having micelle-forming characteristics, P1 showed faster and greater HepG2 cell-binding capacity, plausibly owing to its flexible molecular conformation and higher L-ornithine content. In contrast, the surface charge of normal HK-2 cells treated with anionic PD4 remained essentially constant with an increasing PD4 concentration, sug-gesting a low affinity of PD4 toward normal cells. These results revealed that electrostatic interactions between PLO-based polypeptides and cancer cells were responsible for their binding and represented the first step of anticancer action.
surface charge of HepG2 cells evolved from negative to positive with increasing concentrations of P1 and P4 ( Figure 4A), indicating that cationic PLO-based polypeptides bind to the surface of HepG2 cells via electrostatic interactions. It is interesting to note that, compared to P4, having micelle-forming characteristics, P1 showed faster and greater HepG2 cell-binding capacity, plausibly owing to its flexible molecular conformation and higher L-ornithine content. In contrast, the surface charge of normal HK-2 cells treated with anionic PD4 remained essentially constant with an increasing PD4 concentration, suggesting a low affinity of PD4 toward normal cells. These results revealed that electrostatic interactions between PLO-based polypeptides and cancer cells were responsible for their binding and represented the first step of anticancer action.

Effect of PLO-Based Polypeptides on the Membrane Permeability of Cancer Cells
Once bound to cancer cells, AMPs are expected to destroy the integrity of the cell membrane, thus killing cancer cells via physical action [26]. At this stage, the leakage of the cytoplasmic enzyme LDH from HepG2 cells upon incubation with P1 or P4 was detected. As shown in Figure 4B, a higher extracellular content of LDH was detected with the increase in the concentration of P1 or P4, indicating greater cell membrane damage. In addition, cells treated with P1 (in the absence of the hydrophobic segment) exhibited greater LDH leakage, suggesting that the presence of a hydrophobic segment did not significantly improve the membrane-disrupting ability of PLO-based polypeptides. These results are consistent with the IC 50 values and further show that PLO-based polypeptides exert their anticancer action via a mechanism involving physical membrane disruption ( Figure 4C).

Flow Cytometry Study
The Annexin V-FITC/PI apoptosis detection assay was used to investigate the anticancer mechanisms of the representative copolypeptide P4. Annexin V not only binds to the valgus phosphatidylserine which is exposed on the membrane outer leaflet of apoptotic cells, but also enters necrotic cells which are lacking membrane integrity and binds to phosphatidylserine on the inner leaflet of the bilayer [32]. In contrast, PI can only label the DNA of necrotic cells. As shown in Figure 5, compared to the control group, the proportion of necrotic cells was obviously improved with the increase in the concentration of P4. However, the percentage of apoptotic cells remained less than 5% in all P4 treatment groups. The apoptosis results implied that P4 damaged the cell membrane without obvious cell apoptosis.
greater LDH leakage, suggesting that the presence of a hydrophobic segment did not significantly improve the membrane-disrupting ability of PLO-based polypeptides. These results are consistent with the IC50 values and further show that PLO-based polypeptides exert their anticancer action via a mechanism involving physical membrane disruption ( Figure 4C).

Flow Cytometry Study
The Annexin V-FITC/PI apoptosis detection assay was used to investigate the anticancer mechanisms of the representative copolypeptide P4. Annexin V not only binds to the valgus phosphatidylserine which is exposed on the membrane outer leaflet of apoptotic cells, but also enters necrotic cells which are lacking membrane integrity and binds to phosphatidylserine on the inner leaflet of the bilayer [32]. In contrast, PI can only label the DNA of necrotic cells. As shown in Figure 5, compared to the control group, the proportion of necrotic cells was obviously improved with the increase in the concentration of P4. However, the percentage of apoptotic cells remained less than 5% in all P4 treatment groups. The apoptosis results implied that P4 damaged the cell membrane without obvious cell apoptosis.

Cell Membrane Disruption Viewed by SEM and CLSM
The morphology and structural changes of cancer cells upon incubation with different concentrations of P4 were further examined by SEM. HepG2 cells without PLO-b-PLF treatment showed a smooth and intact membrane surface ( Figure 6). Upon treatment with P4 at lower concentrations (0.25 × IC 50 and 0.5 × IC 50 ), the cell membrane surface became rough but remained intact. However, when the concentration of P4 was further increased (IC 50 , 2 × IC 50 , and 4 × IC 50 ), the cell membrane surface became rougher. Eventually, cell morphology was gradually lost.

Cell Membrane Disruption Viewed by SEM and CLSM
The morphology and structural changes of cancer cells upon incubation with different concentrations of P4 were further examined by SEM. HepG2 cells without PLO-b-PLF treatment showed a smooth and intact membrane surface ( Figure 6). Upon treatment with P4 at lower concentrations (0.25 × IC50 and 0.5 × IC50), the cell membrane surface became rough but remained intact. However, when the concentration of P4 was further increased (IC50, 2 × IC50, and 4 × IC50), the cell membrane surface became rougher. Eventually, cell morphology was gradually lost. The polypeptide-induced cell membrane disruption was further investigated by CLSM. DiO and Hoechst were employed as membrane-staining and nucleus-staining dyes, respectively. As shown in Figure 7, P4 showed dose-dependent cell membrane disruption. HepG2 cells treated with higher concentrations of P4 exhibited a stronger DiO signal, indicating that more dye resided in the cytoplasm due to cell membrane damage. In other words, higher concentrations of P4 led to more obvious cell membrane penetration. The CLSM results were consistent with the SEM results, further suggesting that P4 kills cancer cells through membrane lysis. The polypeptide-induced cell membrane disruption was further investigated by CLSM. DiO and Hoechst were employed as membrane-staining and nucleus-staining dyes, respectively. As shown in Figure 7, P4 showed dose-dependent cell membrane disruption. HepG2 cells treated with higher concentrations of P4 exhibited a stronger DiO signal, indicating that more dye resided in the cytoplasm due to cell membrane damage. In other words, higher concentrations of P4 led to more obvious cell membrane penetration. The CLSM results were consistent with the SEM results, further suggesting that P4 kills cancer cells through membrane lysis.

Migration Inhibition
Inhibiting cancer cell migration represents an anticancer activity of chemotherapeutics. The ability of PD4 to inhibit cancer cell migration at different pH values was evaluated using a scratch wound healing assay ( Figure S9). After 24 h of incubation, cells treated with PD4 at a dose of 0.25 × IC50 and pH 6.5 for 24 h showed reduced cell migration ( Figure   Figure 7. CLSM images of HepG2 cells before and after incubation with different concentrations of P4 for 30 min. Nuclei were stained with Hoechst (blue) while the cell membrane was stained with DiO (green).

Migration Inhibition
Inhibiting cancer cell migration represents an anticancer activity of chemotherapeutics. The ability of PD4 to inhibit cancer cell migration at different pH values was evaluated using a scratch wound healing assay ( Figure S9). After 24 h of incubation, cells treated with PD4 at a dose of 0.25 × IC 50 and pH 6.5 for 24 h showed reduced cell migration ( Figure S9D) compared to the untreated cells ( Figure S9B). Moreover, cells that were treated with 100 µg/mL PD4 at pH 7.4 experienced partial migration ( Figure S9C). These results demonstrate the importance of acidic conditions on the anticancer effect of PD4 and imply that PD4 can not only kill cancer cells but also prevent their metastasis.

Conclusions
Cationic PLO-b-PLF and anionic PLO(DCA)-b-PLF were synthesized, characterized, and evaluated as macromolecular anticancer agents. These amphiphilic block copolypeptides self-assemble into nanosized polymeric micelles in aqueous solution. PLO-b-PLF micelles bind to the surface of cancer cells via electrostatic interactions, disrupt the cancer cell membranes, and kill cancer cells via membrane lysis. This physical mechanism might afford PLO-b-PLF with broad-spectrum anticancer activity and alleviate the problem of drug resistance. The DCA modification of PLO-b-PLF, i.e., PLO(DCA)-b-PLF, prevents cytotoxicity and hemolytic activity under normal physiological conditions. However, the negative-to-positive charge reversal of PLO(DCA)-b-PLF as a result of the hydrolysis of the β-amide bond, under weakly acidic conditions restores cytotoxicity (anticancer activity), improves the anticancer selectivity against tumor cells. It should be noted that low extracellular pH is seen not only in tumors but also in inflammation; therefore, immune cell viability may be impaired. Although an enhanced permeation and retention effect is expected for nanosized PLO(DCA)-b-PLF micelles, the applicability of the pH-responsive antitumor activity of PLO(DCA)-b-PLF requires further careful evaluation. The in vivo antitumor evaluation of PLO(DCA)-b-PLF micelles is currently underway and will be reported in the future.  Table S1: Degree of polymerization (DP) and molecular weights of PZLO-b-PLF. Reference [46] is cited in the supplementary materials.