pH-Induced Charge-Reversal Amphiphile with Cancer Cell-Selective

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Biological and Medical Applications of Materials and Interfaces

pH-Induced Charge-Reversal Amphiphile with Cancer Cell-Selective Membrane-Disrupting Activity Yincheng Chang, Zehuan Huang, Yang Jiao, Jiangfei Xu, and Xi Zhang ACS Appl. Mater. Interfaces, Just Accepted Manuscript • Publication Date (Web): 04 Jun 2018 Downloaded from http://pubs.acs.org on June 4, 2018

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ACS Applied Materials & Interfaces

pH-Induced Charge-Reversal Amphiphile with Cancer Cell-Selective Membrane-Disrupting Activity Yincheng Chang, Zehuan Huang, Yang Jiao, Jiang-Fei Xu, and Xi Zhang*

Key Lab of Organic Optoelectronics & Molecular Engineering, Department of Chemistry, Tsinghua University, Beijing 100084 (China)

KEYWORDS: amphiphiles; membranes; charge reversal; anticancer; chemotherapy.

ABSTRACT: A charge-reversal amphiphile exhibiting charge conversion from negative to positive induced by pH is reported. It selectively kills cancer cells through cell membrane disruption. This amphiphile comprising an alkyl chain and anionic headgroup of acid-labile βcarboxylic amide (C16N-DCA) was prepared. In the microenvironment of normal cells with pH 7.4, the negatively charged C16N-DCA exhibited considerably reduced cytotoxicity. However, in the acidic microenvironment of cancer cells with pH 6.5-6.8, the headgroup charge of C16NDCA changed from negative to positive under hydrolysis of the acid-labile amide group. As a result, the generated cationic amphiphile displayed significant killing of cancer cells by disrupting their cell membranes. Such pH-selective cell killing bioactivity represents a new route of chemotherapy for anticancer strategies.

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INTRODUCTION

Chemotherapy is among the most important therapeutic modalities in clinical anticancer treatments.1-3 Most conventional chemotherapeutic agents such as cis-platinum,4 paclitaxel,5 camptothecin,6 kill cancer cells using mechanisms of DNA damage, antitubulin, enzyme inhibition and others. However, these methods cannot distinguish cancer cells from normal cells very well, resulting in severe side effects to cancer patients.7,8 To deal with this problem, several chemical approaches have been developed, such as targeted drug delivery,9-11 smart nanocarriers responding

to

tumor

microenvironments,12

supramolecular

chemotherapy

based

on

overexpressed tumor biomarkers and others.13-15 Although significant advances have been achieved, new chemotherapeutic agents that distinguish cancer cells from normal cells are required.16 Cationic amphiphiles commonly contain a hydrophilic cationic headgroup and hydrophobic alkyl chain.17,18 Through the disruption of anionic cell membranes,19 cationic amphiphiles show high toxicity toward both microbes and mammalian cells.20-24 We hypothesized that it could be possible to develop new cationic amphiphiles to selectively target cancer cell membranes. However, one challenge is the high similarity between normal cell and cancer cell membranes, which can result in non-selective cell killing.25,26 Therefore, a key point is to develop a safe and effective anticancer cationic amphiphile capable of distinguishing cancer cells from normal cells via this mechanism.


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To this end, we present a facile and feasible strategy to develop anticancer charge-reversal amphiphiles. Through introduction of acid-labile moieties, some research groups have endowed drug delivery systems with charge-reversal property to realize pH-induced release of DNA or clinical anticancer drugs.27-29 We proposed that the acid-labile moieties could be introduced into cationic amphiphiles. Such acid-labile charge-reversal amphiphiles could exhibit high anticancer activity in the acidic cancer microenvironments via cell membrane disruption, while displaying low cytotoxicity in neutral normal cell microenvironments. In this way, a new chemotherapeutic strategy to distinguish cancer cells from normal cells could be established with a nonconventional killing mechanism. As a proof of concept, an amphiphile bearing an alkyl chain and an anionic headgroup of acidlabile β-carboxylic amide (C16N-DCA) was designed. It was easily prepared through the reaction between primary amines of C16N and 1,2-dicarboxylic-cyclohexene anhydride (DCA) (Scheme 1). Thus, C16N-DCA was endowed with a negative charge ascribed to the carboxylic headgroup. In the microenvironment of normal cells with pH 7.4,30 the negatively-charged C16N-DCA might remain stable, thus exhibiting low cytotoxicity to normal cells. However, the acid-labile amide of C16-DCA could be hydrolyzed in the acidic microenvironment of cancer cells with pH 6.5-6.8,31 thus changing the headgroup charge from negative to positive. As a result, the cationic amphiphile (C16N) might display high anticancer bioactivity through disruption of cell membranes only in low pH cancer sites. In this way, distinguishing cancer cells


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from normal cells may be achieved by employing pH-sensitive anticancer charge-reversal amphiphiles.

Scheme 1. Chemical structure of the charge-reversal amphiphiles (C16N-DCA) and its proposed anticancer behavior induced by the acidic microenvironment of cancer cells.

EXPERIMENTAL SECTION

Materials. All reagents were purchased from commercial suppliers and used without further purification unless specified. Water used in this work was triple distilled. Synthesis of C4N-DCA. Butylamine (C4N, 7.3 mg, 0.1 mmol) was added into 10 mL anhydrous DCM. After C4N was dissolved, 1,2-dicarboxylic-cyclohexene anhydride (DCA, 15.2 mg, 0.1 mmol) was added. The mixture was stirred for 0.5 h at room temperature. When the solvent was


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evaporated under reduced pressure, the pure product (C4N-DCA) was obtained with a yield of 99%. Synthesis of C16N-DCA. Synthesis of C16N-DCA was performed similarly to the procedure of C4N-DCA. C16N (24.1 mg, 0.1 mmol) was added into 10 mL anhydrous DCM. After C16N was dissolved, 1,2-dicarboxylic-cyclohexene anhydride (DCA, 15.2 mg, 0.1 mmol) was added. The mixture was stirred for 0.5 h at room temperature. When the solvent was evaporated under reduced pressure, pure product (C16N-DCA) was got with a yield of 99%. Cell culture. The human hepatic cancer cells (HepG2) were cultured in the MEM medium containing 10% FBS and 1% penicillin/streptomycin under the humid atmosphere with 90% humidity and 5% CO2 at 37.0 °C. Human hepatic normal cells (L02) and lung normal cells (BEAS-2B) were cultured in the DMEM medium containing 10% FBS and 1% penicillin/streptomycin under same incubation conditions. Human lung cancer cells (A549) and doxorubicin resistant breast cancer cells (MCF-7/ADR) were cultured in the RPMI-1640 medium containing 10% FBS and 1% penicillin/streptomycin under same incubation conditions. Cell viability assay. We evaluated the cell viabilities of C4N-DCA and C16N-DCA by Cell Counting Kit-8 assay (CCK-8) in vitro. The cells were planted in 96-well plates at a density of 3 × 103 cells per well in 100 µL complete medium. After incubation for 24 h, they were incubated with C4N-DCA or C16N-DCA at certain concentrations for a period of time. The old medium was replaced with fresh medium containing CCK-8. After incubation for another 2 h, the


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absorbance at 450 nm was measured using a microplate reader (EnVision, PerkinElmer). Three independent experiments were performed. Confocal laser scanning microscopy (CLSM). HepG2 and L02 cells were planted in 35 mm plastic bottomed µ-dishes. After incubation for 24 h, the mediums of HepG2 cells were adjusted to pH 6.5 while the mediums of L02 cells were kept at pH 7.4. The cells were then incubated with 30 µM C16N-DCA for 24 h. The dishes were then washed with PBS three times and costained with 1 µM Calcein-AM and 2 µM PI for 15 min. After washing with PBS, the cells were observed under a confocal fluorescence microscope (A1, Nikon). Critical aggregation concentrations (CACs) measurements. CACs of C16N and C16N-DCA were evaluated by pyrene fluorescence probe method according to the previously published literature. 32 Lactate dehydrogenase (LDH) assay. The LDH assay was performed to monitor membrane leakage. According to the manufacturer’s instructions, HepG2 cells were seeded in 96-well plates at a density of 3 × 103 cells per well in 100 µL complete MEM medium. After incubation for 24 h, the old medium was replaced with pH 6.5 complete MEM medium, and the cells were incubated with different concentrations of C16N-DCA for another 24 h. Untreated cells were regarded as a control for background LDH release, and cells treated by Lysis Buffer were regarded as a control for maximal LDH release. To measure LDH release, 100 µL/well of Working Solution was added into each well and the plates were incubated for 20 min at room temperature. After the addition of 50 µL Stop Solution, the absorbance at 490 nm was recorded


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by a microplate reader (EnVision, PerkinElmer). The percentage of LDH release was calculated by the following equation: 33

LDH release (%) = (AbsorbanceC16N-DCA treated cells – Absorbanceuntreated cells)/(AbsorbanceLysis Buffer treated cells -

Absorbanceuntreated cells) × 100

Zeta potential measurements. HepG2 and L02 cells were planted in 6-well plates (5 × 105 cells/well) and cultured in complete DMEM medium for 24 h. The cells were harvested. HepG2 cells were re-suspended in complete DMEM medium in pH of 6.5 containing designed concentration of C16N. L02 cells were re-suspended in complete DMEM medium in pH of 7.4 containing designed concentration of C16N-DCA. After incubation for 0.5 h at 37°C, the cells were centrifuged and re-suspended in 1 mL H2O. The zeta potentials of cells were measured using a Malvern NanoZS90 apparatus. Human xenograft tumor model. Four- to six-week-old female BALB/c nude mice were inoculated with HepG2 cells via intravenous injection. When tumors reached 150-200 mm3, mice were randomized into three groups with eight animals. C16N was dissolved by acidic PBS and C16N-DCA was dissolved in Na2CO3/NaHCO3 solution (0.1 M Na2CO3/NaHCO3). The experimental group was injected with C16N or C16N-DCA solution at the dose of 4 mg kg-1 via the tail vein twice a week. Meanwhile, the vehicle control group was injected with the same volume of bicarbonate buffer. The tumor volumes were measured by digital vernier callipers three times a week. Tumor volumes were calculated using the formula V (mm3) = L (mm) × W


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(mm)2 × 0.5, where L represented the long diameter and W represented the short diameter measured by caliper. Tumor Volumn Change (%) = Vt / V0 × 100, where Vt represented the tumor volumn at t day and V0 represented the tumor volumn at 0 day. Body Weight Change (%) = BWt / BW0 × 100, where BWt represented the body weight at t day and BW0 represented the body weight at 0 day. Hemolytic test, pharmacokinetics experiment and histology analysis, were performed according to the previously published literature.33-35

RESULTS AND DISCUSSION

Hydrolysis of the acid-labile amide group in different pH conditions. We hypothesized that the charge reversal of β-carboxylic amide could be achieved in pH 6.5-6.8 while the βcarboxylic amide may remain intact at pH 7.4. To answer this question, we employed 1H NMR spectroscopy to investigate the hydrolysis process in different pH conditions. Considering that C16N-DCA has relatively poor water-solubility and possible aggregation in high concentration, a model molecule (C4N-DCA) was prepared and utilized (Figure 1a) for the 1H NMR experiments. Hydrolysis of C4N-DCA at pH 6.5, 6.8 and 7.4 was monitored by 1H NMR. As shown in Figure 1b, the proton peak ascribed to Hb of the hydrolyzed product increased much faster at pH 6.5 and 6.8 compared to that at pH 7.4.


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Figure 1. a) The synthesis and hydrolysis scheme for C4N-DCA. b) The hydrolysis of C4NDCA at pH 6.5, pH 6.8, pH 7.4 characterized by 1H NMR (phosphate buffer, D2O).

To make these data more quantitative, the percentage of hydrolyzed product was calculated based on the integrated ratio of the two peaks (Ha and Hb) (Figure S6a, b, c). Hydrolysis kinetics curves of the C4N-DCA at different pH conditions were obtained as shown in Figure 2, which indicate that the β-carboxylic amide is easily hydrolyzed at pH 6.5 and 6.8, but relatively stable at pH 7.4. Therefore, the pH-induced charge reversal of β-carboxylic amide from negative to positive can be achieved selectively and reliably in different pH conditions.


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Figure 2. The hydrolysis kinetics of C4N-DCA at pH 6.5, 6.8, and 7.4, respectively.

Low cytotoxicity to normal cells and high killing efficiency to cancer cells. To test if the distinctive charge reversal of C16N-DCA could lead to low cytotoxicity to normal cells and high killing efficiency to cancer cells, extensive cell culture experiments were performed. Human hepatic normal and cancer cell lines were selected as model demonstrations in vitro. The killing efficiency of C16N-DCA against hepatic cancer cells (HepG2) and cytotoxicity to hepatic normal cells (L02) in different pH conditions were evaluated after different incubation times with the Cell Counting Kit-8 (CCK-8) assay.32 As shown in Figure S7, C16N-DCA exhibited high killing bioactivity against both HepG2 cancer cells and L02 normal cells in acidic conditions (pH 6.5 and pH 6.8) for 24 h, 48 h, and 72 h, respectively. Meanwhile, almost no cytotoxicity of C16N-DCA against L02 normal cells and HepG2 cancer cells was observed in pH 7.4 conditions even after incubation for 72 h. The results above indicate that the killing bioactivity of C16N-DCA is hidden in physiological conditions while can be activated by acidic conditions effectively.


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Figure 3. a) The calculated IC50 of C16N-DCA (24 h) toward HepG2 and L02 cells. b) The calculated IC50 of C16N-DCA (24 h) toward A549 and BEAS-2B cells. All cell viabilities were estimated by CCK-8 assay. Data represent mean ±s.d., n=3.

Considering the different pH conditions in normal and cancer cell microenvironments, HepG2 cancer cells were cultured with different concentrations of C16N-DCA in acidic conditions (pH 6.5), while L02 normal cells were cultured with C16-DCA in physiological conditions (pH 7.4) to calculate the IC50 value. The IC50 value of C16N-DCA to HepG2 cancer cells was determined to be 7.9 µM, which was much lower than that for anticancer drug such as cisplatinum (IC50 33.7 µM, in pH 6.5 conditions, Figure S8). More importantly, the IC50 value of C16N-DCA to L02 normal cells was 74.9 µM, significantly higher than the IC50 value to HepG2 cancer cells (Figure 3a). To demonstrate the general pH-selective anticancer applicability of


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C16N-DCA, human lung normal cells (BEAS-2B) and cancer cells (A549) were further investigated. Alike with hepatic normal and cancer cells, the IC50 values of C16N-DCA to BEAS-2B normal cells (88.6 µM) was also much higher than the IC50 value to A549 cancer cells (18.6 µM) (Figure 3b). In addition, distinct growth inhibition effects were also directly demonstrated through a live/dead cell co-staining assay. In Figure 4a, nearly 100% of HepG2 cells were killed (red) through treatment with 30 µM C16N-DCA for 24 h. By contrast, viability of L02 cells remained (green) after the same treatment (Figure 4b). It should be noted that both the growth of HepG2 and L02 cells were inhibited by treatment with cationic amphiphiles (30 µM C16N), suggesting that the charge reversal is essential for distinguishing cancer cells from normal cells (Figure S9). Therefore, these results clearly indicate that the charge-reversal amphiphile is able to distinguish cancer cells from normal cells in cell culture.


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Figure 4. a) HepG2 and b) L02 cells co-stained by PI (red, dead cells) and calcein AM (green, live cells) after exposure to 30 µM C16N-DCA for 24 h.

To investigate the possible influence of aggregation of C16N-DCA on its anticancer cell activity, critical aggregation concentrations (CACs) of C16N and C16N-DCA were measured by the pyrene fluorescence probe method.32 As shown in Figure S10, the CAC values of C16N and C16N-DCA were calculated to be 378 µM and 136 µM, respectively. Since the concentrations in cell experiments were in the range of 0 - 120 µM, C16N and C16N-DCA would not be expected to aggregate in these conditions. Therefore, the possible influence of aggregation of C16N and C16N-DCA on their cancer cell killing performance is excluded. The mechanism of cell membranes disruption. We assumed that the mechanism behind the pH-dependent cancer cell killing performance of C16N-DCA could be ascribed to its disruption of cell membranes. To support this hypothesis, we firstly employed in situ differential interference contrast microscope (DIC) to monitor cell membrane disruption induced by C16NDCA. A series of images showing the process was obtained in Figure 5 and the video was also attached in Supplementary Movie 1. At the beginning (0 h), all HepG2 cells were alive with relatively clear edges. After adding C16N-DCA, the cell membranes of HepG2 cells were strongly disturbed by the charge-reversed form of C16N-DCA (C16N) as time evolved (0 ~ 7.2 h). Consequently, the cell membranes edges became unclear, and the cells also shrunk. Then, cytoplasm leaked from the cells (13.7 ~ 24 h) and the HepG2 cells shriveled, indicating that


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these cells were finally killed. Therefore, the above results suggest that disruption of cell membranes induced by C16N-DCA is responsible for its high cancer cell bioactivity.

Figure 5. In situ microscopy images of HepG2 cells with 100 µM C16N-DCA treatment for different times.

The membrane-disruption mechanism should rely on two points as follows: first, the chargereversed form of C16N-DCA (C16N) binds the surface of the cell membrane via electrostatic interactions. The second point is that the long alkyl chain of C16N inserts into the cell membrane and destabilizes it, thus inducing cytoplasm leakage and cell killing. To confirm the first point, we studied the interaction between C16N and HepG2 cells by zeta potential measurements.36 As shown in Figure 6a, the zeta potential values of HepG2 cells increased significantly by treating them with increasing amounts of C16N, indicating that C16N is capable of partitioning into the cell membranes. On the contrary, little change of zeta potential was observed for L02 normal cells treated by C16N-DCA under the same conditions. This result suggested weak binding


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capability of C16N-DCA and supported the low cytotoxicity of C16N-DCA against L02 normal cells. Next, we detected the leakage of the cytoplasmic enzyme, lactate dehydrogenase (LDH) by an in vitro assay to confirm the second point.37 As shown in Figure 6b, the amounts of LDH from the cytoplasm increased with increasing concentrations of C16N-DCA. More importantly, the LDH amounts were inversely related to cell viabilities obtained by cell incubation experiments treated with C16N-DCA. These data indicate that the insertion of C16N into the cell membrane can induce cytoplasmic leakage, thus killing the cells. In addition, the cytotoxicity of C4N-DCA, the control compound without the long alkyl chain, toward HepG2 evaluated by CCK-8 assay, was negligible up to a concentration of 150 µM, indicating the necessary intercalation of long alkyl chains to disrupt cell membranes (Figure S11). Therefore, the membrane-disruption mechanism is proposed as the reason behind the cancer cell killing of the charge-reversal amphiphile (Scheme 1).


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Figure 6. a) Zeta potential of HepG2 and L02 cells treated by different concentrations of C16N and C16N-DCA after 0.5 h incubation, respectively. b) Relative cell viabilities of HepG2 cells treated with C16N-DCA and the consequent LDH release caused by cell membrane leakage.

In vivo pharmacokinetics and tumor inhibitory effect. In vivo pharmacokinetics of C16N and C16N-DCA were studied using rat model. After a same dose of either C16N or C16N-DCA intravenously (i.v.), plasma levels of the parent compounds were monitored during period of 12 h (Figure 7a). Pharmacokinetic parameters of C16N and C16N-DCA were assessed and summarized (Table S1). In the beginning, C16N-DCA achieved a much higher initial concentration (C0) than C16N. What’s more, C16N-DCA had a longer half-life (t1/2) and lower clearance (CL) compared with C16N. As a whole, C16N-DCA showed a superior pharmacokinetic profile with high plasma compound levels, which suggested the better stability


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and lower adhesion to plasma proteins than C16N. In addition, the hemolytic activity of C16NDCA and C16N was tested on cony blood. As shown in Figure 7b, C16N-DCA exhibited much lower hemolytic activity than C16N, demonstrating the significance of charge-reversal amphiphile for safety improvement.

Figure 7. a) In vivo pharmacokinetics of C16N-DCA and C16N in rat. b) Hemolysis of C16NDCA and C16N tested on cony blood.

To further evaluate the inhibitory effect of C16N-DCA on tumor growth in vivo, HepG2 tumor-bearing nude mice were employed. As shown in Figure 8a, tumor volume in the vehicle control group and C16N group increased over 6-fold over the original state after 19 days. Very interestingly, the growth of HepG2 tumors was remarkably slowed by intravenous injection of


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C16N-DCA. As shown in Figure 8b, no apparent loss of body weight was evident for mice treated with C16N-DCA, indicating that no severe systemic toxicity was observed. In addition, the vehicle control group and treated mice were sacrificed to collect the tumor and major organs (including heart, liver, spleen, lung, and kidney) for hematoxylin and eosin (H&E) staining after 19 days of intravenous injections. Histology shows substantial dead cells with clear tumor destruction in the images of H&E-stained tumor tissue treated by C16N-DCA, further confirming the effective anticancer activity of C16N-DCA in vivo (Figure 8c).34 It is very inspiring that there is no clear sign of organ damage observed in the H&E-stained organ slices (Figure S12). Therefore, these results indicate that C16N-DCA has considerable anticancer efficacy and no significant side effect in vivo.

Figure 8. a) HepG2 tumor growth profiles of the mice intravenously injected with bicarbonate buffer (vehicle control), C16N and C16N-DCA at dose of 4 mg kg-1 (* p < 0.05). b) Body weight change of the mice treated with bicarbonate buffer (vehicle control), C16N and C16N-DCA. c)


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H&E-stained tumor slices collected from mice treated with bicarbonate buffer (vehicle control) and C16N-DCA treatment. Data represent mean ±S.E., n = 8.

CONCLUSION

In conclusion, we have developed a new chemotherapeutic strategy to selectively inhibit cancer cells from normal cells through the construction of pH-induced charge-reversal amphiphiles. One main advantage is that the charge-reversal amphiphiles can be easily synthesized with precise molecular structures and desired cancer efficacy in mouse models via disruption of cell membranes. In addition, this facile approach may be of general applicability to other kinds of clinical anticancer drugs for further improving their selectivity for cancer cells versus normal cells. Even though the charge-reversal amphiphiles slow tumor growth instead of halting growth, the rich structure and function of amphiphiles provide a wide range of options for drug candidates. It is anticipated that this line of research may open up a new route for cancer chemotherapeutic design.

ASSOCIATED CONTENT

Supporting Information. Figure S1-S12 and Table S1 were shown in Supporting Information. Movie shows cell membrane disruption (Movie S1).

AUTHOR INFORMATION Corresponding Author


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*E-mail: [email protected]

ORCID

Xi Zhang: 0000-0002-4823-9120

Author Contributions The manuscript was written through contributions of all the authors. All authors have given approval to the final version of the manuscript.

Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT

This work was financially supported by the Ministry of Science and Technology of China (2018YFA0208900) and National Postdoctoral Program for Innovative Talents (BX201600082). We thank Prof. David W. Grainger (The University of Utah) for polishing the English. We also thank Dr. Bin Yuan, Dr. Yuetong Kang and Qi Hao (Tsinghua University) for their helpful discussions.

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SYNOPSIS


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pH-Induced Charge-Reversal Amphiphile with Cancer Cell-Selective

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