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Light-Activatable Red Blood Cell Membrane-Camouflaged Dimeric Prodrug Nanoparticles for Synergistic Photodynamic/Chemotherapy Qing Pei, Xiuli Hu, Xiaohua Zheng, Shi Liu, Yawei Li, Xiabin Jing, and Zhigang Xie ACS Nano, Just Accepted Manuscript • DOI: 10.1021/acsnano.7b08219 • Publication Date (Web): 18 Jan 2018 Downloaded from http://pubs.acs.org on January 18, 2018

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Light-Activatable Red Blood Cell Membrane-Camouflaged Dimeric Prodrug Nanoparticles for Synergistic Photodynamic/Chemotherapy Qing Pei,†,‡ Xiuli Hu,*, † Xiaohua Zheng, Zhigang Xie*, †

†,‡

Shi Liu,

†

Yawei Li,

†

Xiabin Jing,† and

†

State Key Laboratory of Polymer Physics and Chemistry, Changchun Institute of

Applied Chemistry, Chinese Academy of Sciences,5625 Renmin Street, Changchun, Jilin 130022, P. R. China ‡

University of Science and Technology of China, Hefei, Anhui, 230026, P. R. China

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ABSTRACT. Biomimetic approach offers numerous opportunities to design therapeutic platforms with enhanced antitumor performance and biocompatibility. Herein

we

report

red

blood

cell

membrane-camouflaged

nanoparticles

(RBC(M(TPC-PTX))) for synergistic chemo- and photodynamic therapy (PDT). Specifically, the inner core is mainly constructed by reactive oxygen species (ROS)-responsive

PTX

dimer

(PTX2-TK)

and

photosensitizer

5,10,15,20-tetraphenylchlorin (TPC). In vitro experiments show that the prepared RBC(M(TPC-PTX)) is readily taken up into endosomes. Under appropriate light irradiation, the TPC can generate ROS, not only for PDT, but also for triggering PTX2-TK cleavage and on-demand PTX release for chemotherapy. In vivo results show that the coating of RBC membrane prolongs blood circulation and improves tumor accumulation. The combination of chemo- and photodynamic therapy enhances anticancer therapeutic activity and light-triggered drug release reduces systematic toxicity. All these characteristics render the described technology extremely promising for cancer treatment.

KEYWORDS: prodrug nanoparticles, controlled drug release, cell membrane biomimetic, light sensitive, synergistic cancer therapy


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Cancer remains one of the most life-threatening diseases. As the mainstay in clinical tumor treatment, chemotherapy is limited by its severe side effects primarily caused by nonspecific action and poor tumor selectivity.1 Extensive efforts have been devoted to develop diverse drug delivery systems (DDS) to minimize side effects and improve the therapeutic efficacy, ranging from polymer or protein conjugates,2-4 to nanoparticle (NP) systems such as micelles and liposomes,5,

6

and to gels and

capsules. Among these DDS, micelles and liposomes have been extensively explored because of their distinct nano-structural characteristics.7 Indeed, many NPs could alter the biodistribution and pharmcokinetics of small molecular drugs and have shown superior performances in vitro and in vivo, and some have been clinically approved.8-11 However, due to various biological barriers at the systemic, tissular, cellular, and subcellular levels after intravenous injection, only 0.7 % (median) of the administered dose is reported to be delivered to the tumor sites.12 The clinical applications of nanomedicines are extremely limited. The rapid blood clearance rate, the premature drug release during blood circulation, the nonspecific uptake by normal organs, and poor tumor tissue penetration are the main obstacles that lie between the administration site and the therapeutic action site within the cancer cells.13 Even entering the cancer cells, the spontaneous or stimulated drug resistance weakens the efficacy of chemotherapy. So it is a persistent and significant challenge for the rational design of advanced NPs to cross the physical and biological barriers and overcome the drug resistance.


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In the past few years, the biomimetic approach using natural cell membrane to camouflage NPs has attracted special attention for their intrinsic biocompatibility, biodegradability, and nonimmunogenicity.14 The emerging delivery platform combines the advantages of natural cellular entities for long circulation and targeting abilities and synthetic biomaterials for controlled drug retention and releases.15 So far, typical cell membranes investigated for functionalizing NPs include red blood cell (RBC),16-18 platelets,19, 20 macrophage cell,21 leukocyte (WBC),22 stem cell and cancer cell membranes.23, 24 Among the various types of cell membranes, RBC membrane represents the most abundant, biocompatible, and affordable biological carriers and has been investigated as biomimetic coating on artificial nanocarriers for distinct functionalities.14 The most popular inner core is composed of drug loaded polymeric NPs from poly(lactic-co-glycolic acid) (PLGA),16 poly(caprolactone) (PCL), pluronic copolymer25 or other inorganic materials.26 However, in most of these formulations, large quantities of excipient materials are required to encapsulate the drug payloads due to the low drug loading capacity (typically less than 10 %).27 It also remains concerning with regard to excipient-associated side toxicities. On the other hand, some suboptimal properties such as premature drug release and poor tumor tissue penetration also cause the reduction in the therapeutic efficacy. More advanced biomimetic DDS that can unify high drug loading and on-demand drug release properties,28 recognize the microenvironment, and release the cargoes in spatial-, temporal-, and dosage-controlled fashions are highly desirable.29, 30


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Moreover, the single chemotherapy can hardly eliminate tumors because of the heterogeneity of tumor. One alternative is the combination of multiple therapies with different mechanisms. Photodynamic therapy (PDT), an important clinically approved noninvasive therapeutic modality, utilizes photosensitizers (PS) to generate cytotoxic reactive oxygen species (ROS), especially highly reactive singlet oxygen (1O2), which interacts with adjacent biological macromolecules and thus leads to cell death.31 PDT is considered to be an ideal noninvasive stimulus that can precisely treat the tumor with facile, flexible, and spatiotemporal control.32

Herein, we constructed smart RBC membrane-camouflaged dimeric prodrug NPs with light-activatable drug release for synergistic photodynamic-chemotherapy. This NP platform is composed of two components: (i) paclitaxel (PTX) dimer and 5,10,15,20-tetraphenylchlorin (TPC) loaded NPs for drug retention and light-triggered amplified drug release; (ii) RBC membrane-based outer shell for long blood circulation. Different from the reported systems, the inner core is mainly composed of paclitaxel dimer (PTX2-TK). The dimeric drug strategy has been proved to be an effective method to improve the drug loading content by several groups.33-35 In our previous work, the drug loading content of PTX dimer in methoxypoly(ethylene glycol)-block-poly(D, L-lactide) (PEG-b-PDLLA) can be as high as 85 wt%.36-38 Considering the much higher ROS level in cancer cells (up to 10×10-6 M) than in normal tissue (10×10-9 M),39-41 ROS-responsive PTX dimer with thioketal (TK) as the linker, PTX2-TK, was designed and synthesized successfully. Furthermore, in view of the different ROS levels in different cancer cells and the short lifetime of ROS.42 5 ACS Paragon Plus Environment

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Photosensitizer TPC was also introduced for in situ generation of ROS to regulate on-demand drug release triggered by light.

As shown in Scheme 1, inner NPs were first prepared by co-encapsulation of PTX2-TK and TPC into copolymer PEG-b-PDLLA via the solvent exchange method. The obtained NPs were abbreviated as M(TPC-PTX). TPC was selected for its efficient PDT capability and π−π stacking interaction with PTX for its facile and high drug loading in the copolymer PEG-b-PDLLA.43 The RBC membrane vesicles (RBCVs) derived from RBC were then coated on the surface of M(TPC-PTX) to obtain RBC(M(TPC-PTX)). After intravenous (i.v.) injection, RBC(M(TPC-PTX)) is expected to bypass macrophage uptake and systemic clearance, and to accumulate in the tumor site by the enhanced permeability and retention effect for the nanostructure.44 With cellular internalization, the photosensitizer moieties embedded in the inner core will generate singlet oxygen (1O2) upon light irradiation, leading to the disruption of the endosome and induction of cell death. At the same time, the generated ROS would subsequently interact with the thioether moiety linker to activate PTX2-TK,45, 46 leading to a sequential and amplified PTX release. Finaly, the free PTX quickly difffuses into the microtubule for enhanced chemotherapy. This biomimetic nanoplatform integrates photodynamic and chemotherapy into a single platform with high drug loading and on-demand drug release and is promising to overcome the rapid blood clearance and premature drug release, thus maximizing the therapeutic efficacy while minimizing the side effects of each therapeutic agent.


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Scheme 1. Schematic illustration of RBC membrane-camouflaged dimeric prodrug NP (RBC(M(TPC-PTX))), their prolonged blood circulation, and light triggered on-demand drug release and combined photodynamic/chemotherapy.

RESULTS AND DISCUSSION

Synthesis and Characterization of RBC-membrane-coated NPs. The synthetic route for PTX dimer bridged with ROS-responsive thioketal linker (abbreviated as PTX2-TK) was shown in Figure S1. PTX2-TK was synthesized through a one pot reaction of PTX and ROS-cleavable thioketal linker in the presence of


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1-(3-dimethylaminopropyl)-3-ethylcarbodiimide

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hydrochloride

(EDC·HCl)

and

4-dimethylaminopyridine (DMAP). The obtained PTX2-TK was purified via a silica gel column with high yields (> 84%) and its chemical structure and purity were confirmed by 1H NMR and linear ion trap mass spectrometer (LTQ-MS). As shown in Figure S2, the disapperance of characteristic peak of 2’-hydroxyl group of PTX at 3.61 ppm and the shift of the resonance of 2’-CH of PTX from 4.8 to 5.5 ppm and the appearance of the peaks near 2.6-2.8 ppm (Figure S2) verified that the esterification reaction between TK and PTX was at the 2’-hydroxyl group of PTX. The particular peak values in mass spectra were consistent with that of theoretical calculation, indicating the successful synthesis of PTX2-TK as shown in Figure S3. The perfect purity of

PTX2-TK was further confirmed

by high

performance liquid

chromatography (HPLC) (Figure S4). Photosensitizer TPC was selected for its hydrophobic structure and π-π stacking interaction with PTX, which facilitated TPC encapsulation and NP stability.47, 48 All the protons could be clearly resolved in 1H NMR (Figure S5) and the peak at m/z 617.5 in the LTQ-MS spectra was the same to its theoretical molecular weight (Figure S6), confirming the successful synthesis and high purity of TPC.

The RBC-membrane-coated NPs were prepared via two steps: inner particle preparation and surface membrane coating. The inner core was first prepared by co-encapsulating PTX2-TK and TPC with PEG-b-PDLLA in aqueous solution through the solvent exchange methods.37 The obtained NPs were abbreviated as M(TPC-PTX). Single drug PTX or TPC loaded NP, M(PTX) or M(TPC), was used as 8 ACS Paragon Plus Environment

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the control. Next, RBC membrane coated M(TPC-PTX) (abbreviated as RBC(M(TPC-PTX))) was obtained by co-extruding RBC-membrane derived vesicles (RBCVs)16 and M(TPC-PTX) through 450-nm porous membranes. M(PTX), M(TPC), M(TPC-PTX), and RBC(M(TPC-PTX)) all displayed homogeneous spherical structure observed by transmission electron microscopy (TEM) (Figure 1a). After coating with RBCVs, the resulted RBC(M(TPC-PTX)) explicitly revealed an spherical core-shell structure, indicating the successful coating of RBCVs onto M(TPC-PTX) (Figure 1a). Compared with M(TPC-PTX), the average hydrodynamic diameters of RBC(M(TPC-PTX)) increased from 161 nm to 168 nm according to the dynamic light scattering (DLS) results in Figure 1b. The increased 7 nm in diameter was likely attributed to the thickness of RBC membranes generated from the lipid bilayer.49 The surface zeta potential increased from −24.0 mV to −28.5 mV upon fusing with the RBCs (Figure 1c). Both the increased size and increased surface negative charge further confirmed the successful coating of RBCs with M(TPC-PTX) (Figure 1 b and c).50, 51 The loading content and loading efficiency were 38 wt% and 82% for PTX2-TK, 13 wt% and 73% for TPC, respectively, as determined by HPLC and UV−vis absorption spectra. The optical properties of M(TPC-PTX) and RBC(M(TPC-PTX)) were investigated by UV−vis absorption and photoluminescence spectra. Compared with the small molecule TPC, the maximum absorption of M(TPC-PTX) and RBC(M(TPC-PTX)) both had a red-shift of about 20 nm from 417 nm to 437 nm (Figure S7a), ascribing to the formation of aggregation structure in NPs.52 Due to dexter-type excitonic migration between porphyrins molecules in the 9 ACS Paragon Plus Environment

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aggregation state,53 tremendous fluorescence quenching was observed for all NP formulations (Figure S7b). However, the fluorescence of TPC in NPs could still be used for the following confocal laser scanning microscopy (CLSM) and flow cytometry (FCM) examination.

The Stability of RBC(M(TPC-PTX)). The stability of M(TPC-PTX) and RBC(M(TPC-PTX)) were tested by monitoring their size and size distribution in PBS (pH 7.4) solution containing with 10% fetal bovine serum (FBS). As shown in Figure 1d, M(TPC-PTX) and RBC(M(TPC-PTX)) dispersions remained stable for up to 24 h in the FBS containing solutions, indicating their superior serum stability. Additionally, M(TPC-PTX) and RBC(M(TPC-PTX)) dispersions remained stable and monodispersed for up to 18 days in aqueous solution at 4 oC (Figure S8), guaranteeing the feasibility of long time storage.

Characterization of the membrane proteins on RBC(M(TPC-PTX)). The complicated membrane proteins on RBC membranes played a vital role on immune-evasive function.22 Following, the successful transfer of complicate the membrane proteins onto the surface of RBC(M(TPC-PTX)) was examined by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE). Emptied RBC membrane and RBCVs were used as parallel controls. Figure 1e showed that the three protein brands displayed the same stripes, indicating the membrane proteins were reserved during the preparation of RBCVs and RBC(M(TPC-PTX)). Since being equipped with large numbers of “self-recognized” proteins of RBC membrane, 10 ACS Paragon Plus Environment

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RBC(M(TPC-PTX)) was expected to escape from the reticuloendothelial system (RES) capture and realize long blood circulation.16,

54

Then we examined the

bioactivity of membrane proteins present on the surface of RBC(M(TPC-PTX)) by testing its ability to escape from the capture of macrophage phagocytosis. M(TPC-PTX) and RBC(M(TPC-PTX)) were incubated with the mouse macrophage RAW264.7 cells and the cells were examined by CLSM and FCM. As shown in Figure 1f, the conspicuous lower red fluorescence in RAW264.7 cells incubated with RBC(M(TPC-PTX)) than that in cells incubated with M(TPC-PTX) suggested that coating of RBC membranes significantly suppressed the cellular uptake of RBC(M(TPC-PTX)). The cellular uptake was further quantified by FCM as shown in Figure S9, which agreed well with the CLSM results in Figure 1f. All the above results demonstrated that RBC membrane had been successfully cloaked onto the surface of M(TPC-PTX) and the bioactivity of membrane proteins were reserved.


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Figure 1. Biophysical characterization of RBC(M(TPC-PTX)). (a) TEM and enlarged TEM images (top right inset) of M(PTX), M(TPC), M(TPC-PTX) and RBC(M(TPC-PTX)). (b) Size distribution and (c) zeta-potential of M(PTX), M(TPC), M(TPC-PTX) and RBC(M(TPC-PTX)). (d) Size changes of M(PTX), M(TPC), M(TPC-PTX) and RBC(M(TPC-PTX)) in PBS (pH 7.4) solution containing with 10% fetal bovine serum (FBS); (e) SDS-PAGE protein analysis of empty RBCs, RBCVs and RBC(M(TPC-PTX)). (f) CLSM images of RAW264.7 cells cultured with M(TPC-PTX) and RBC(M(TPC-PTX)) at 37 oC for 6 h, respectively. Scale bar, 20 µm.

The cleavage of thioketal linker in PTX2-TK by H2O2. The thioketal linker in the prodrug PTX2-TK was expected to be cleaved by ROS upon irradiation for on-demand drug release.42, 46 So it is crucial to test the efficiency of thioketal linker 12 ACS Paragon Plus Environment

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response to ROS. HPLC was used to monitor the cleavage of thioketal and hydrolysis of PTX precursor from PTX2-TK in the presence of 10 mM H2O2. PTX2-TK exhibited a monodispersed peak at an elution time of 6.3 min (Figure 2a). During the first 7 h, the peak at elution time of 6.3 min decreased dramatically and completely disappeared within 16 h. At the same time, a new peak at elution time of 5.1 min, belonging to PTX-SO3H according to the literature,55, 56 appeared within 1 h and increased gradually. After 7 h, the new peak at elution time of 3.7 min, belonging to PTX, emerged and increased over time, while the peak belonging to PTX-SO3H decreased gradually. LTQ-MS was further used to characterize the structure of the degradation products. As shown in Figure 2b, the peaks at m/z 1958.6 belonging to PTX2-TK disappeared after H2O2 treatment, and peaks at m/z 988.4 and 888.3 belonging to PTX-SO3H and PTX, respectively, were observed. While for the control group, the peak at m/z 1958.6 belonging to PTX2-TK did not change after treatment for 120 h without H2O2 (Figure S10), which agreed well with the HPLC results (Figure 2a). The possible H2O2-triggered degradation mechanism of PTX2-TK was illustrated in Figure 2c and PTX2-TK degradation triggered by H2O2 occured in the following three steps: (i) cleaving thioketal linker into two sulfydyl; (ii) oxidation of the sulfydyl to hydrophilic sulfoacid;57,

58

(iii) release of active PTX molecule.

Considering the upper limit of intracellular H2O2 concentration in normal cells could up to be 10×10-6 M,40, 41 the degradation of PTX2-TK at biological dose ROS (100 µM) was also tested. As shown in Figure S11, there was no peak belonging to PTX appeared even after 96 h. This indicated that the low H2O2 concentration in normal 13 ACS Paragon Plus Environment

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cells was not enough to cleave PTX2-TK and limited active drug would be released during blood circulation, thus diminishing the side effects.

Figure 2. The degradation of PTX2-TK triggered by H2O2. (a) The degradation of PTX2-TK in the presence of 10 mM H2O2 monitored by HPLC; (b) LTQ-MS of PTX2-TK after 120 h treatment in the presence of 10 mM H2O2; (c) The proposed degradation mechanism of PTX2-TK triggered by H2O2. ROS

Generation

in

RBC(M(TPC-PTX))

upon

Irradiation.

Although

intracellular ROS could be as an indicator of cancer, however, considering their short lifetime (< 0.1 ms) and the heterogeneity of cancer, it is necessary and more effective to generate ROS in situ to trigger the degradation of PTX2-TK and induce specific on-demand PTX release from RBC(M(TPC-PTX)). Therefore, we investigated the ROS, mainly 1O2, production of TPC in RBC(M(TPC-PTX)) upon irradiation. ROS 14 ACS Paragon Plus Environment

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generation was determined by the UV−vis absorption spectra of ROS capturer, indocyanine green (ICG).59, 60 Upon the irradiation with a 638 nm laser lamp (100 mW/cm2) for 7 min, the maximum absorption band of ICG at 779 nm decreased rapidly in M(TPC), M(TPC-PTX) and RBC(M(TPC-PTX)) solutions (Figure 3a-c), indicating the effective generation of ROS, while the maximum absorption band of the pure ICG solution decreased gently with light irradiation (Figure S12). M(TPC) and M(TPC-PTX) had similar ROS generation efficacy (Figure 3c), confirming the addition of PTX2-TK did not impact the ROS production. Compared with M(TPC-PTX), the presence of RBC(M(TPC-PTX)) decreased ICG absorption more slowly. This may be ascribed to the delayed ROS diffusion from the inner core to the solution due to the hindrance of RBC membrane in RBC(M(TPC-PTX)). When vitamin C (VC), another ROS capturer,61 was added, the decrease of maximum absorption peak of ICG in both RBC(M(TPC-PTX)) and M(TPC-PTX) solutions was significantly inhibited, further confirming ROS generation upon irradiation (Figure S13). Additionally, the photostability of all nanoformulations, including M(TPC), M(TPC-PTX), and RBC(M(TPC-PTX)), was confirmed by negligible change of the maximum absorption wavelength (Figure S14).

The cleavage of thioketal linker in PTX2-TK by ROS. Then, HPLC was further used to test if the generated ROS could break thioketal linker in RBC(M(TPC-PTX)) and trigger PTX release. We irradiated RBC(M(TPC-PTX)) with a 638 nm laser lamp (100 mW/cm2) for different times before monitoring the hydrolysis of PTX2-TK using 15 ACS Paragon Plus Environment

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HPLC. Figure 3d illustrated the decreased HPLC intensity corresponding to PTX2-TK at elution time of 6.3~6.8 min over time under light irradiation, confirming the cleavage of PTX2-TK triggered by ROS. As shown in Figure 3e, almost 96% % of PTX2-TK disappeared in 2 h. In sharp contrast, alsmot no PTX2-TK degradated without irradiation during the same time period, indicating that the generated ROS could break thioketal linker in PTX2-TK. When the light switched periodically between on and off for 0.5 h each time, a pulsatile degradation was observed (Figure 3f), indicating the sensitivity of thioketal linker response to light irradiation. The smaller spherical nanoparticles observed by TEM images of RBC(M(TPC-PTX) upon light irradiation indicated that ROS could break the integrity of RBC membranes and released its internal contents (Figure S15). Thus, the present formulation could reduce the premature drug release during blood circulation and control drug release upon light irradiation at desired site, which is promising for reducing systemic toxicity and enhancing theraputic effect.

ROS generation in Hela cells. Following, we investigated the ROS generation inside the cancer cells by incubating M(TPC-PTX) and RBC(M(TPC-PTX)) in cervical cancer (HeLa) cells, using PBS as the control group. ROS generation was detected by a cell-permeable ROS-sensitive fluorescent probe, 2′,7′-dichlorofluorescin diacetate (DCFH-DA). DCFH-DA is nonfluorescent, but it could be rapidly oxidized to a fluorescent molecule (dichlorofluorescein, DCF) by ROS62. As shown in Figure 3g, negligible fluorescence was observed when the cells were incubated with RBC(M(TPC-PTX)) without light irradiation, similar to that of the control groups 16 ACS Paragon Plus Environment

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(PBS and PBS (L+)). However, obvious green fluorescence was seen when the cells were cultured with RBC(M(TPC-PTX)) plus light irradiation (RBC(M(TPC-PTX)) (L+)). When VC was added, the green fluorescence signal of DCF decreased significantly, which further confirmed the ROS genenration inside the cancer cells during light irradiation. The results were in accordance with the above results detected by ROS capturer ICG (Figure 3a-c).

Figure 3. Singlet oxygen generation ability of as-prepared NPs. Time-dependent UV absorption spectra of ICG in (a) M(TPC-PTX) and (b) RBC(M(TPC-PTX)) solutions after irradiation with a 638 nm laser lamp (100 mW/cm2) for 7 min. (c) The decay rate of ICG at 779 nm in M(TPC), M(TPC-PTX) and RBC(M(TPC-PTX)) solutions with light irradiation. (d) HPLC intensity changes corresponding to PTX2-TK at elution time of 6.3~6.8 min over time under light irradiation. (e) The degradation of 17 ACS Paragon Plus Environment

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PTX2-TK in RBC(M(TPC-PTX)) over time upon irradiated by 638 nm laser lamp (100

mW/cm2) tested

by

HPLC.

(f) The

degradation

of

PTX2-TK

in

RBC(M(TPC-PTX)) triggered by light (638 nm laser lamp, 100 mW/cm2) switched periodically between on and off for 0.5 h each time. (g) The generation of intracellular ROS indicated by the fluorescence of DCF in HeLa cells incubated with PBS, PBS (L+), RBC(M(TPC-PTX)), RBC(M(TPC-PTX) (L+), and RBC(M(TPC-PTX) (L+) (VC+), respectively. Scale bar, 20µm.

Intracellular uptake and distribution of NPs. The intracellular uptake and distribution of M(TPC-PTX) and RBC(M(TPC-PTX)) were investigated in HeLa cells. As shown in Figure 4a, the RBC(M(TPC-PTX)) group exhibited comparable intracellular green fluorescence intensity in HeLa cells to that of M(TPC-PTX), indicating that coating of RBC membranes negligibly influenced the internalization of M(TPC-PTX) in the cancer cells.63, 64 The increased green fluorescence signals as a function of time from 2 h to 6 h suggested the efficient internalization of both M(TPC-PTX) and RBC(M(TPC-PTX)) by cells in a time-dependent manner, and the fluorescence mainly located in the cytoplasm of cells in both groups (Figure S16). Additionally, the green fluorescence of TPC in RBC(M(TPC-PTX)) colocalized perfectly with the red fluorescence from Lyso-Tracker, indicating the accumulation of RBC(M(TPC-PTX)) in endolysosome (Figure S17). With light irradiation, the red fluorescence intensity of lysosome tracker decreased, which was attributed to the damage of lysosome escape caused by 1O2 pump (Figure S18).65


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In vitro cytotoxicity evaluation. The in vitro cytotoxicity was evaluated against HeLa cells by using the 3-(4,5-dimethyl thiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay. As shown in Figure 4b, both M(TPC) and RBC(M(TPC)) exhibited negligible cytotoxicity, indicating the biocompatibility of RBC membrane and low dark toxicity of photosensitizer TPC. Upon light irradiation, increased cytotoxicity

was

observed

for

RBC(M(TPC))

(L+),

M(TPC-PTX)

(L+),

RBC(M(TPC-PTX)) (L+) compared with their corresponding NPs without light irradiation. For RBC(M(TPC)) (L+) group, the cell cytotoxicity of TPC was limited at the present dosage, because the cell viability was still higher than 50% even at TPC doses at 13.75 µM. By contrast, the cell cytotoxicity of PTX-containing NPs, including M(TPC-PTX) (L+) and RBC(M(TPC-PTX)) (L+), have been greatly improved upon irradiation. These results verified the synergetic antitumor efficacy of PDT and chemotherapy. Annexin-V FITC and propidium iodide (PI) double staining assay was applied to quantitatively examine the cell apoptosis induced by PTX and TPC in RBC(M(TPC-PTX)) by FCM analysis. As shown in Figure 4c, the light irradiation had no effect on cell apoptosis for there was no difference between cells incubated with PBS and PBS with irradiation. The late apoptosis increased from 6.55% to 12% for cells treated with M(TPC-PTX) without irradiation and M(TPC-PTX) (L+). The most significant late apoptosis (48.1%) was observed in cells treated with RBC(M(TPC-PTX)) (L+). Both the CLSM result in Figure 4b and FCM result in Figure 4c confirmed the enhanced PDT and chemotherapy upon light irradiation.


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In order to visually evaluate the in vitro therapeutic effect, the cells were stained with calcein-AM and PI to identify live and dead/late apoptotic cells. Figure 4d showed that most cells treated with PBS or PBS (L+) were alive with strong green fluorescence. Much more cell death was observed for RBC(M(TPC-PTX)) (L+) than that for M(TPC-PTX) and M(TPC-PTX) (L+) (Figure 4d), which was consistent with the FCM results (Figure 4c). The immunofluorescence staining of tubulin was further applied to examine acting mechanism of PTX that released from RBC(M(TPC-PTX)) triggered by ROS. As reported, the action site of PTX is the microtubules and PTX can be combined with the specific site of microtubule protein, which can promote the polymerization of microtubules and stabilize them, thus interfering with the mitosis of cells.66 As shown in Figure 4e, the microtubule network of HeLa cells was well organized in the control group (PBS and PBS (L+)). On the contrary, variable degrees of microtubule bundle could be visualized for all PTX-containing groups, including M(TPC-PTX), M(TPC-PTX) (L+), RBC(M(TPC-PTX)), and RBC(M(TPC-PTX)) (L+). Under irradiation, much more microtubule bundle was observed for M(TPC-PTX) (L+) and RBC(M(TPC-PTX)) (L+) compared with those without irradiation (M(TPC-PTX) and RBC(M(TPC-PTX)). Notably, RBC(M(TPC-PTX)) (L+) group exhibited the most microtubule bundle, indicating that ROS could cleave PTX2-TK in RBC(M(TPC-PTX)) and release potent PTX. Collectively, these results validated the efficient therapeutic effect enabled by combination of PDT from TPC and chemotherapy from PTX.


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Figure 4. Cellular uptake of M(TPC-PTX) and RBC(M(TPC-PTX)) detected by CLSM (left) and FCM (right). Scale bar, 20 µm. (b) Cell cytotoxicity of M(TPC), RBC(M(TPC)),

RBC(M(TPC))

(L+),

M(TPC-PTX),

M(TPC-PTX)

(L+),

RBC(M(TPC-PTX)) in dark or under irradiation with 638 nm laser lamp (100 mW/cm2) for 5 min against HeLa cells after incubation for 48 h. (c) Flow cytometry analysis of early and late apoptosis of HeLa cells incubated with PBS, PBS (L+), M(TPC-PTX), M(TPC-PTX) (L+), RBC(M(TPC-PTX)), and RBC(M(TPC-PTX)) (L+) for 24 h at TPC and PTX concentration of 13.75 µM and 0.325 µM, respectively, and then treated with or without light irradiation (638 nm laser lamp, 100 mW/cm2) for 5 min. The four quadrants indicate the four status of cells: necrotic (Q1), late-stage apoptotic (Q2), early apoptotic (Q3), and live (Q4). (d) Fluorescence microscopic 21 ACS Paragon Plus Environment

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images of HeLa cells co-stained with calcein AM (green, live cells) and propidium iodide (red, dead cells) after being incubated with NP formulations at TPC and PTX concentration of 13.75 µM and 0.325 µM, respectively, with or without light irradiation (638 nm laser lamp, 100 mW/cm2) for 5 min. Scale bars, 100 µm. (e) Microtubule bundle formation induced by the prepared NPs with or without irradiation for 15 min against HeLa cells. Scale bar, 20µm.

In vivo circulation and biodistribution of NPs. Then we compared the in vivo circulation and biodistribution of M(TPC-PTX) and RBC(M(TPC-PTX)). After a single i.v. injection (equivalent PTX2-TK dose of 17.4 mg/kg body weight, n=4), blood samples were taken out at different time intervals and the concentration of total plasma PTX2-TK was determined as a function of time using HPLC. As shown in Figure 5a, RBC(M(TPC-PTX)) group displayed longer blood circulation than naked M(TPC-PTX) NP, which could be ascribed to the external RBC membrane.16, 67, 68 Additionally, the biodistribution of both NPs in the main organs such as liver and tumor at different time points (0 h, 1 h, 3 h, 6 h, 23 h, 47 h) were also examined. As shown in Figure 5b, the coating of RBC membranes reduced the capture by liver, indicating that RBC(M(TPC-PTX)) contained the “self-recognized” protein of RBC membrane that could reduce reticuloendothelial system (RES) uptake in the liver.54 Figure 5c displayed the PTX2-TK concentration in the tumor tissue at different time points after injection of M(TPC-PTX) and RBC(M(TPC-PTX)), respectively. The RBC(M(TPC-PTX))

group

showed

higher

PTX2-TK

concentration

than

M(TPC-PTX) over time. The M(TPC-PTX) group and RBC(M(TPC-PTX)) group 22 ACS Paragon Plus Environment

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displayed their maximum PTX2-TK concentrations at 3 h and 23 h after drug administration, respectively. At 23 h post treatment, the PTX2-TK concentration in tumor a was 4.6-fold higher in RBC(M(TPC-PTX)) group than that in M(TPC-PTX) group, further demonstrating the importance of RBC membrane coating on the surface. The immunofluorescence staining of the tumorous vasculature by blood vessel marker CD31 was used to investigate the distribution of RBC(M(TPC-PTX)) at the tumor region before and after light irradiation. As shown in Figure S19, obvious light-induced intratumoral extravasation and deep tumorous penetration were observed, verifying the high accumulation of RBC(M(TPC-PTX)) at tumor region and the light triggered drug release upon irradiation and subsequent deep penetration.

In vivo antitumor efficacy. The in vivo antitumor efficacy was then evaluated using human cervical carcinoma HeLa tumor-bearing nude mice. Tumor-bearing mice were randomly divided into seven groups with six mice in each group: PBS, PBS (L+),

M(PTX),

M(TPC)

(L+),

M(TPC-PTX),

M(TPC-PTX)

(L+),

RBC(M(TPC-PTX)) and RBC(M(TPC-PTX)) (L+). When the tumor grew to a size of 50 mm3, 5 days after inoculation of the cancer cells, different formulations with equivalent dose of PTX (30 mg/kg) and TPC (10 mg/kg) were given via tail i.v. injection. At 6 h post treatment, the tumor site of mice in irradiation group was irradiated by 638 nm laser lamp (200 mW/cm2) for 15 min. Following, the tumor size was measured every other day. Figure 5d showed the tumor size as a function of time. The growth of the tumor was inhibited in a certain extent after the treatment with M(TPC) (L+), M(PTX), M(TPC-PTX), M(TPC-PTX) (L+), RBC(M(TPC-PTX)) and 23 ACS Paragon Plus Environment

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RBC(M(TPC-PTX)) (L+), respectively, compared with the PBS control group and PBS (L+) group. The similar tumor volume growth profiles of PBS (L+) group and PBS group indicated light irradiation had no effect on tumor inhibition. The slight tumor inhibition for M(PTX) and M(TPC-PTX) groups indicated that the single chemotherapy could not completely eliminate the tumor. The tumor treated with M(TPC-PTX) (L+) showed remarkably smaller volume than that treated with M(TPC) (L+) and M(PTX) groups (Figure 5d and 5e), indicating the synergetic antitumor efficacy enabled by the combination of PDT and PTX. More importantly, the strongest antitumor effect was achieved by RBC(M(TPC-PTX)) (L+) due to the long blood circulation and preferential tumor accumulation. The tumor volume and weight of the excised tumor (Figure 5e and 5g) agreed well with that measured in living mice (Figure 5d). Meanwhile, the body weight of mice remained stable during the treatment, suggesting no systemic toxicity for all formulations (Figure 5f). The histologic images of the tumor section stained by the hematoxylin and eosin (H&E) showed the highest level of tumor cell nuclear ablation after treatment with RBC(M(TPC-PTX)) (L+) as shown in Figure 5h. H & E staining also showed that all formulations had no distinct damage to main organs, including heart, liver, spleen, lung and kidney as shown in Figure S20. Furthermore, serum biochemical analysis of kidney and liver function parameters showed no difference with those in control group (Figure S21). These results demonstrated that RBC(M(TPC-PTX)) (L+) possessed excellent therapeutic effect but reduced systemic toxicity. We believe that the combination of ROS-responsive PTX dimer and photosensitizer, and subsequently 24 ACS Paragon Plus Environment

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coating them with RBC membranes would promisingly serve as the distinguished candidates for cancer treatment.

Figure 5. In vivo treatment. (a) In vivo PTX2-TK plasma concentration-time profiles of M(TPC-PTX) and RBC(M(TPC-PTX)) after i.v. administration to mice (n =4) at a PTX equivalent dose of 15 mg kg-1. Quantitative analysis of PTX2-TK concentration in liver (b) and tumor (c) at different time after i.v. administration to mice (n =4) with M(TPC-PTX) and RBC(M(TPC-PTX)) at a PTX equivalent dose of 15 mg kg-1. (d) Change of tumor volumes of subcutaneous xenografts in nude mice treated with PBS, PBS

(L+),

M(PTX),

M(TPC)

(L+),

M(TPC-PTX),

M(TPC-PTX)

(L+), 25

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RBC(M(TPC-PTX)) and RBC(M(TPC-PTX)) (L+). (e) Quantitative analysis of tumor weight of PBS, PBS (L+), M(PTX), M(TPC) (L+), M(TPC-PTX), M(TPC-PTX) (L+), RBC(M(TPC-PTX)) and RBC(M(TPC-PTX)) (L+) groups. (f) Body weight of tumor bearing mice after systemic injection. (g) Photos of excised tumors. From top to down: PBS, PBS(L+), M(PTX), M(TPC) (L+), M(TPC-PTX), M(TPC-PTX) (L+), RBC(M(TPC-PTX)) and RBC(M(TPC-PTX)) (L+). (h) Representative H&E-stained tumor slice images of mice after various treatment. Scale bar, 200 µm. All data (d-g) represented as mean ±SEM (n= 6). The levels of significance were set at probabilities of *p< 0.05, **p< 0.01, and ***p

Light-Activatable Red Blood Cell Membrane ... - ACS Publications

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