Plasma membrane activatable polymeric nanotheranostics with self-enhanced light-triggered photosensitizer cellular influx for photodynamic cancer therapy

Accepted Manuscript Plasma membrane activatable polymeric nanotheranostics with self-enhanced light-triggered photosensitizer cell influx for photodynamic cancer therapy

Hao-Ran Jia, Yao-Wen Jiang, Ya-Xuan Zhu, Yan-Hong Li, HongYin Wang, Xiaofeng Han, Zhi-Wu Yu, Ning Gu, Peidang Liu, Zhan Chen, Fu-Gen Wu PII: DOI: Reference:

S0168-3659(17)30077-9 doi: 10.1016/j.jconrel.2017.04.030 COREL 8775

To appear in:

Journal of Controlled Release

Received date: Accepted date:

14 February 2017 22 April 2017

Please cite this article as: Hao-Ran Jia, Yao-Wen Jiang, Ya-Xuan Zhu, Yan-Hong Li, Hong-Yin Wang, Xiaofeng Han, Zhi-Wu Yu, Ning Gu, Peidang Liu, Zhan Chen, FuGen Wu , Plasma membrane activatable polymeric nanotheranostics with self-enhanced light-triggered photosensitizer cell influx for photodynamic cancer therapy. The address for the corresponding author was captured as affiliation for all authors. Please check if appropriate. Corel(2017), doi: 10.1016/j.jconrel.2017.04.030

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ACCEPTED MANUSCRIPT Plasma membrane activatable polymeric nanotheranostics with self-enhanced light-triggered photosensitizer cell influx for photodynamic cancer therapy

Hao-Ran Jia a, Yao-Wen Jiang a, Ya-Xuan Zhu a, Yan-Hong Li a, Hong-Yin Wang a, Xiaofeng b,

**, and Fu-Gen Wu a, *

State Key Laboratory of Bioelectronics, School of Biological Science and Medical Engineering,

Southeast University, Nanjing 210096, P. R. China

Department of Chemistry, University of Michigan, 930 North University Avenue, Ann A rbor,

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b

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Michigan 48109, United States c

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a

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Han a, Zhi-Wu Yu c, Ning Gu a, Peidang Liu d, Zhan Chen

Key Laboratory of Bioorganic Phosphorous Chemistry and Chemical Biology (Ministry of Education),

School of Medicine, Southeast University, Nanjing 210009, P. R. China

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d

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Department of Chemistry, Tsinghua University, Beijing 100084, P. R. China

* Corresponding author. State Key Laboratory of Bioelectronics, School of Biological Science and Medical Engineering, Southeast University, Nanjing 210096, P. R. China.

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** Corresponding author. Department of Chemistry, Univers ity of Michigan, 930 North Univers ity Avenue, Ann Arbor, Michigan 48109, United States. E-mail addresses: [email protected] (F.G. Wu), [email protected] (Z. Chen).

ACCEPTED MANUSCRIPT ABSTRACT To address the issue of low cellular uptake of photosensitizers by cancer cells in photodynamic therapy (PDT), we designed a smart plasma membrane-activatable polymeric nanodrug by conjugating the photosensitizer protoporphyrin IX (PpIX) and polyethylene glycol (PEG) with glycol chitosan (GC). The as-prepared GC-PEG-PpIX can self-assemble into core-shell nanoparticles (NPs) in aqueous solution and the fluorescence of PpIX moieties

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in the inner core is highly quenched due to strong π–π stacking. Interestingly, when

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encountering plasma membranes, the GC-PEG-PpIX NPs can disassemble and stably attach to plasma membranes due to the membrane affinity of PpIX moieties, which effectively

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suppresses the self-quenching of PpIX, leading to significantly enhanced fluorescence and singlet oxygen (1 O2 ) production upon laser irradiation. The massively produced 1 O 2 can

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compromise the integrity of the plasma membrane, enabling the influx of extracellular nanoagents into cells to promote cell death upon further laser irradiation. Through local

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injection, the membrane anchored GC-PEG-PpIX enables strong physical association with tumor cells and exhibits highly enhanced in vivo fluorescence at the tumor site. Besides,

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excellent tumor accumulation and prolonged tumor retention of GC-PEG-PpIX were realized after intravenous injection, which ensured its effective imaging- guided PDT.

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Keywords:

Responsive polymeric nanoparticle Cell surface engineering

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Glycol chitosan

Wash-free membrane imaging Photodynamic therapy

ACCEPTED MANUSCRIPT 1. Introduction Photodynamic therapy (PDT), a noninvasive and highly selective theranostic modality, has great potential as a treatment for numerous malignant and nonmalignant diseases [1–3]. It has been applied to destruct selected tumors with clinical approval and several studies have

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confirmed that PDT is a feasible therapeutic option against early-stage cancer and

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co-therapeutic approach in late-stage cancer [4]. Typically, PDT involves a photosensitizer

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(PS), which can transfer energy to surrounding molecular oxygen to generate reactive oxygen species (ROS), primarily singlet oxygen (1 O2 ), which can consequently induce cell apoptosis

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or necrosis and tissue destruction [5]. Nonetheless, PDT still serves as a fringe cancer

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treatment option due to many limitations, especially a lack of an ideal clinical PDT formulation [6]. One crucial issue is that most classic PS molecules are hydrophobic and

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prone to aggregate in aqueous condition, leading to a severe self-quenching effect, and

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therefore drastically reducing 1 O2 generation as well as the therapeutic efficacy [7,8]. Besides, problems including difficult intravenous delivery [6] and low selectivity toward tumor areas remain poorly solved [9].

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To circumvent these drawbacks of classic PSs, many studies have reported that incorporating PS molecules into nanoparticles (NPs) can improve their properties and achieve better cancer theranostic effects [10]. These PS-containing NPs include inorganic NPs [11– 20], quantum dots [21–23], liposomes [24–26], micelles [27–30], protein-based NPs [31–36], nucleic acid-based NPs [37,38], and porphysomes [39,40]. Besides, many stimuli-responsive polymeric drug carriers have been developed to achieve controlled release of PS molecules and activatable PDT strategy [41–43]. Typically, most of these PDT nanoconstructs require

ACCEPTED MANUSCRIPT cellular uptake to exert lethal photodynamic damage to cancer cells. Hence, the PDT efficacy depends largely on cellular uptake efficiency and subcellular localization of PS molecules. Currently, low drug efficacy in cancer treatment is due, at least partially, to the inadequate cellular uptake and/or the improper efflux of drug molecules by cancer cells. As a functional

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barrier, the plasma membrane plays a pivotal role in controlling the transport of substances.

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To enhance the cellular uptake of PS nanocarriers, modifying the surface of NPs with

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targeting ligands has been a commonly used strategy to strongly interact with corresponding receptors on plasma membranes [44]. However, the cellular uptake efficiency is still limited

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depending on the type of ligand–receptor interaction and the density of surface ligands [45].

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On the other hand, the subcellular distribution of PS molecules is diverse; they can be in lysosome, mitochondrion, cell membrane, Golgi apparatus, and nucleus [1,46]. Among these

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locations, plasma membrane which protects cells from outside environment and also plays an

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important role in many biological events, serves as an extremely sensitive target for PDT. Only mild PDT treatment can lead to membrane dysfunction and ultimately cell death [1]. To the best of our knowledge, aside from a few relevant investigations [25,47,48], plasma

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membrane-targeted PDT remains largely unexplored. In this work, we rationally designed a “smart” plasma membrane-activated PDT construct which achieves a self-enhanced PS influx by actively increasing plasma membrane permeability upon laser irradiation. Generally, this construct is composed by three components: glycol chitosan (GC), polyethylene glycol (PEG), and protoporphyrin IX (PpIX). Through chemical conjugation, PEG segments and PpIX molecules were linked to the GC backbone to form the final construct GC-PEG-PpIX (Fig. 1a). When dispersed in aqueous

ACCEPTED MANUSCRIPT solution, the amphiphilic GC-PEG-PpIX molecules self-assemble into core-shell structured NPs (Fig. 1b), where the hydrophobic PpIX moieties form the inner core due to the π–π interaction between neighboring PpIX moieties, while the hydrophilic PEG chains attached to the GC backbones form the outer shell. PpIX moieties in this state rarely generate 1 O2 due to intramolecular and

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the fluorescence resonance energy transfer (FRET) mediated

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intermolecular quenching (self-quenching). When approaching plasma membranes, however,

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the PpIX moieties insert into lipid bilayers in a multisite anchoring manner and are separated from each other, resulting in the disassembly of GC-PEG-PpIX NPs. The large distances

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between neighboring PpIX moieties significantly prevent the self-quenching effect of the PS 1

O2

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molecules, leading to enhanced 1 O 2 generation efficiency. Under laser irradiation,

oxidation severely damages plasma membrane structures and causes increased membrane

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permeability of the amphiphilic compounds [49], enabling influx of extracellular

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GC-PEG-PpIX into cells to promote cell death. In addition, the development of this new amphiphilic construct of GC-PEG-PpIX also promotes the advancement of cell surface engineering, which is a recently emerged field that

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can modify cell membranes and endow the cells with new properties and functions [50–58]. Due to the suppressed self-quenching effect of PpIX, the GC-PEG-PpIX construct not only enables enhanced

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O2 production upon laser irradiation, but also realizes wash-free

fluorescence imaging for plasma membranes. The intrinsic fluorescence property of PpIX can also be used for monitoring the subcellular localization of the PDT agent after endocytosis. Through local injection, the membrane anchored GC-PEG-PpIX enables strong physical association with tumor cells and exhibits highly enhanced in vivo fluorescence at the tumor

ACCEPTED MANUSCRIPT site. Besides, excellent tumor accumulation and prolonged tumor retention of GC-PEG-PpIX were realized after intravenous injection, which verified its feasibility of imaging-guided PDT. In vivo study also demonstrates effective tumor elimination, negligible systemic toxicity, and good hemocompatibility of GC-PEG-PpIX, showing great potential for future clinical

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application.

Fig. 1. Schematic illustration of plasma membrane imaging-guided PDT based on cell surface engineering. (a) Chemical structure of GC-PEG-PpIX. (b) Proposed membrane-anchoring mechanism for GC-PEG-PpIX and the corresponding plasma membrane-activated PDT.

2. Materials and methods

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2.1. Materials

Glycol chitosan (GC) was purchased from Sigma-Aldrich (St. Louis, MO) and its molecular

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weight and degree of deacetylation were reported to be 67 kDa and 88%, respectively [50]. Methoxyl

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PEG2000 succinimidyl ester (NHS-PEG2000-OMe) was bought from Nanocs, Inc. (New York, NY).

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Protoporphyrin IX (PpIX), N-hydroxysuccinimide (NHS), and dimethyl sulfoxide (DMSO) were obtained from Aladdin Chemistry Co. Ltd. 1-Ethyl-3-(3-(dimethylamino)propyl)carbodimide (EDC)

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was purchased from Sigma-Aldrich. Dialysis membranes with a molecular weight cut-off (MWCO) of 10 kDa (Spectra/Por®6 Dialysis membranes, Regenerated Cellulose) were ordered from Spectrumlabs.

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2.2. Synthesis of GC-PEG-PpIX

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Deionized water (18.2 MΩ·cm) was obtained from a Milli-Q system (Millipore, Billerica, MA).

To obtain GC-PEG, 5 mg GC and 10 mg NHS-PEG2000-OMe were separately dissolved in 1 mL

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PBS solution (50 mM, pH 7.4), and were mixed together for 4 h under stirring at room temperature. Then the mixture solution was dialyzed (MWCO = 10 kDa) against deionized water for 3 days and GC-PEG was obtained after lyophilization. Then, 0.54 mg PpIX (1.0 mg/mL in DMSO), 1.49 mg EDC (1.0 mg/mL in DMSO), and 1.66 mg NHS (1.0 mg/mL in DMSO) were mixed together for 0.5 h at room temperature. The activated PpIX was added to the above obtained GC-PEG (dissolved in 1 mL H2 O) to react under stirring overnight. The mixture was dialyzed (MWCO = 10 kDa) for 3 days against DMSO and 1 day against H 2O, respectively. Finally, GC-PEG-PpIX was obtained after

ACCEPTED MANUSCRIPT lyophilization. For control experiments, PpIX-conjugated GC (GC-PpIX) was also synthesized following the similar synthetic procedure. Before experiments, a suspension of GC-PpIX in water was sonicated for a total of 3 min-pulse period using a probe-type sonicator at 45 W in 5 s-intervals (3

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s-pulse application; 2 s-off) due to the extremely poor solubility of the reagent.

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2.3. Characterization

The morphology and size of GC-PEG-PpIX NPs were characterized by transmission electron

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microscopy (TEM). The GC-PEG-PpIX solution (100 μg/mL) was deposited onto a glow-discharged

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carbon-coated grid, followed by staining with 2% phosphotungstic acid. After being dried at room temperature, the sample was imaged using a transmission electron microscope (JEM-2100, JEOL Ltd,

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Japan). The hydrodynamic diameters of GC-PEG-PpIX NPs were characterized by dynamic light scattering (DLS) using a Zetasizer instrument (Nano ZS, Malvern Instruments, United Kingdom). To

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measure the degree of chemical conjugation of PpIX in GC-PpIX and GC-PEG-PpIX, samples were dispersed in 2 mL of water and UV–vis absorbance spectra were recorded using a UV-2600

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spectrophotometer (Shimadzu). According to the standard concentration curve of free PpIX, the concentration of PpIX in these samples was quantified by observing the absorbance at 540 nm. Fluorescence spectra of these samples were recorded using a spectrofluorophotometer (RF-5301PC, Shimadzu, Japan).

2.4. Preparation of Supported Lipid Bilayer and Giant Unilamellar Vesicle

ACCEPTED MANUSCRIPT The vesicle fusion method was adopted to prepare supported lipid bilayers (SLBs). First, liposomes

were

prepared

by

the

extrusion

1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine

method.

(POPC)

Typically, and

1-palmitoyl-2-{12-[(7-nitro-2-1,3-benzoxadiazol-4-yl)amino]dodecanoyl}-sn-glycero-3-phosphocholi

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ne (NBD-PC) were dissolved in chloroform, respectively, and mixed together at a molar ratio of

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99.5:0.5. Then, chloroform was evaporated under a stream of nitrogen gas and further dried under

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vacuum overnight. The thus-formed dry lipid films were hydrated in deionized water and mechanically extruded through a 100 nm polycarbonate filter for 21 times. Next, suspension of

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liposomes was pipetted onto an ultraclean glass substrate (pretreated with oxygen plasma), followed

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by incubation with a buffer solution (400 mM KCl, 4 mM Tris, 2 mM CaCl2 , pH 8.0) for 30 min, during which the liposomes fused together on the substrate to form the final supported bilayer. Finally,

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the SLB was rinsed with PBS solution repeatedly to remove excess lipids. POPC with 0.5 mol% of NBD-PC was dissolved in chloroform to make a lipid solution with a

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final concentration of 4 mg/mL. Then we used spin-coating to produce a thin, homogeneous film which was dried under vacuum overnight to evaporate the residual chloroform. Next, a common ly

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used electroformation method was adopted to form GUVs [59]. In our experiment, the electroformation chamber between two indium tin oxide (ITO) glasses was filled with 0.23 M sucrose. The electroformation process lasted for 2 h with a 3 V, 10 Hz ac field at 60o C. Subsequently, the solution was gently removed from the chamber. After being diluted 3 times with an iso-osmolar solution of 0.23 M glucose, GUVs were deposited onto the coverslip due to the density difference between sucrose (inner pool) and glucose (external medium) followed by observation under an inverted confocal laser scanning microscope TCS SP8 (Leica, Germany).

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2.5. Cellular experiments

Adenocarcinomic human alveolar epithelial cells (A549) were cultured in Dulbecco’s modified

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Eagle’s medium (DMEM), supplemented with 10% fetal bovine serum (FBS) and 100 IU/mL

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penicillin-streptomycin at 37o C in a humid atmosphere with 5% CO 2. For confocal imaging, A549

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cells were seeded at a density of 3 × 104 cells per well in 35 mm glass dishes and incubated at 37o C for 24 h. Then cells were treated with free PpIX, GC-PpIX, and GC-PEG-PpIX (5 μg/mL of free PpIX)

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for a certain period of time at 37o C. Confocal fluorescence images were taken with an inverted

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confocal laser scanning microscope TCS SP8 (Leica, Germany) with a 63× oil immersion objective. To quantify the fluorescence intensity, A549 cells were incubated with GC-PEG-PpIX (5 μg/mL of

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free PpIX) for different time periods. Then the cells were washed by PBS, digested with trypsin, harvested in fresh culture medium and analyzed by a flow cytometer (ACEA Bioscience, NovoCyte,

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SD). Channel used for analyses was PE Texas Red with the excitation at 488 nm. To observe the morphological changes in the plasma membrane, A549 cells were incubated with

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GC-PEG-PpIX (5 μg/mL of PpIX). After 15 min of incubation, cells were irradiated with a laser at the power of 14 mW/cm2 . After irradiation, cells were observed immediately under a confocal microscope with a 100× oil immersion objective. To evaluate the cytotoxicity of PpIX, GC-PpIX, and GC-PEG-PpIX, cells were seeded into 96-well plates at a density of 5 × 103 cells per well in 100 μL complete culture medium. When growing to 80% confluence in each well, cells were incubated with various concentrations of free PpIX, GC-PpIX, and GC-PEG-PpIX for 36 h. Then, cell viabilities were determined by a cell

ACCEPTED MANUSCRIPT counting kit-8 (CCK-8). Typically, media were carefully aspirated and 100 μL DMEM containing CCK-8 (10%) was added to each well. After incubation for an additional 2 h at 37o C, the absorbance at 450 nm was measured against a background control using a microplate reader (Multiskan FC, Thermo-Scientific, USA).

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To study the phototoxicity of GC-PEG-PpIX in vitro, A549 cells were seeded into a 96-well plate

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and incubated with free PpIX, GC-PpIX, and GC-PEG-PpIX (5 μg/mL of free PpIX) for 15 min.

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Afterwards, cells were illuminated with a laser (635 nm, 14 mW/cm2 ) for pre-determined irradiation time. After incubation for another 4 h, CCK-8 assay was carried out as described above to determine

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2.6. In vivo fluorescence imaging

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the cell viabilities.

Athymic female nude mice, purchased from Yangzhou University Medical Centre (Yangzhou,

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China), were used for animal experiments under protocols approved by Southeast University Laboratory Animal Center. To establish the subcutaneous tumor model, 2 × 106 murine cervical U14

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carcinoma cells were inoculated onto the back of each mouse. The mice were used when tumor volumes reached approximately 50 mm3 . Before in vivo imaging, mice were fully anesthetized by inhalation of a mixture of oxygen with isoflurane (5%) under general anesthesia. For intratumoral administration, 100 μL of PpIX, GC-PpIX, and GC-PEG-PpIX with a 48 μg/mL equivalent concentration was adopted. For intravenous administration, 100 μL of PpIX, GC-PpIX, and GC-PEG-PpIX at the dose of 5 mg/kg PpIX was adopted. In vivo fluorescence imaging was performed on a Cri Maestroin and PerkinElmer in vivo imaging system with excitation wavelength of

ACCEPTED MANUSCRIPT 540 nm and emission wavelength of 670 nm. CRi Maestro Image software was required to conduct the ROI (regions of interest) analysis.

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2.7. In vivo PDT

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When U14 tumors grew to about 40 mm3 in volume, nude mice were injected intravenously with

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100 μL of saline, free PpIX, GC-PpIX, and GC-PEG-PpIX (dose: PpIX 5mg/kg), respectively. Tumors were irradiated with a 635 nm laser (30 mW/cm2 , 18 h after injection) for 20 min every other

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day for a total of three treatments. The tumor sizes were monitored by a caliper every 2 days for 22

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days and the tumor volumes were calculated as width2 × length/2.

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2.8. Ex vivo histological staining

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To investigate the PDT-induced damage to the tumor cells, representative mice from different groups were sacrificed at 22nd day after treatment and tumors were dissected and then fixed in 4%

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formaldehyde solutions, followed by making paraffin sections. Next, the paraffin sections were performed with hematoxylin and eosin (H&E) following standard protocol. All sections were observed using a Leica TCS SP8 microscope. To evaluate the systemic toxicity of GC-PEG-PpIX, the mice injected with saline (control) and GC-PEG-PpIX were sacrificed 22 days after treatment, and the organs including heart, liver, spleen, lung, and kidneys were taken out to conduct H&E staining following the same process described above.

ACCEPTED MANUSCRIPT 2.9. Hemolysis experiment

Blood from mouse was collected in tubes containing sodium citrate and used immediately. Red blood cells (RBCs) were isolated by centrifugation (1000 rpm, 5 min), washed several times with PBS

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solution and suspended in PBS solution. Then, 0.5 mL of the RBC suspension was mixed with 0.5 mL

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suspension containing different concentration of GC-PEG-PpIX at 200, 500, and 1000 μg/mL in PBS

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solution to give a final concentration of 100, 250, and 500 μg/mL, respectively. RBCs incubated with ultrapure water and PBS solution were set as positive and negative control, respectively. The mixture

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was incubated at room temperature for 2 h and then centrifuged at 5000 rpm for 5 min. The

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percentage of hemolysis was calculated as follows:

Hemolysis% = (sample absorbance – negative control absorbance) / (positive control absorbance –

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negative control absorbance) × 100%.

3. Results and discussion

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3.1. Design and characterization of GC-PEG-PpIX NPs

GC is a highly water-soluble, biocompatible, and biodegradable polymer, which has been broadly applied in drug delivery [60–63] due to its ease of chemical modification (via amine groups) and excellent drug-loading capacity. Besides, GC is positively charged at physiological pH and serves as an excellent choice for cell surface modification [50,64]. PEG has been approved by the US Food and Drug Administration (FDA) as a safe hydrophilic

ACCEPTED MANUSCRIPT polymer for drug delivery to increase stability and pro long blood circulation time in vivo. PpIX, a derivative of porphyrin, is also very safe since it is the key component of hemoglobin in red blood cells. Porphyrin has been widely explored in multimodal imaging and phototherapy [28,65,66], and the porphyrin-based PS molecules are known as the earliest and

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most useful PS molecules in clinical trials. Through chemical conjugation, the as-prepared

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GC-PEG-PpIX showed excellent water solubility, which is due to the presence of PEG

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segments. This was confirmed by the poor water solubility of the PEG- free compound, GC-PpIX, which could only be soluble after tip sonication but still easily formed

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agglomerates within 1 day after dissolution. Transmission electronic microscopy (TEM)

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result revealed that GC-PEG-PpIX self-assembled into spherical NPs of approximately 80 nm in aqueous solution, and such observation was in good agreement with the hydrodynamic

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diameter (82.2 ± 22.1 nm) measured by dynamic light scattering (DLS) (Fig. 2a). In contrast,

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GC-PpIX showed a smaller size at ~51 nm (Supplementary Fig. 1). The larger size of GC-PEG-PpIX as compared to that of GC-PpIX implied a relatively more loose nanostructure formed with the presence of PEG segments. Besides, the chemical conjugation

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degree of PpIX molecules was calculated to be ~4% of the total repeat units of glycol chitosan, determined by the UV–vis spectrum of GC-PEG-PpIX.

3.2. Increased fluorescence emission and 1 O2 generation of GC-PEG-PpIX upon membrane binding

It can be envisaged that, if the hydrophobic PpIX moieties of GC-PEG-PpIX can anchor

ACCEPTED MANUSCRIPT on plasma membranes, the self-assembled GC-PEG-PpIX nanostructure may be destroyed. It has been reported that nanoparticles bearing hydrophobic moieties are readily disassembled in the presence of surfactants such as sodium dodecyl sulfate (SDS) [63,67]. We believe that it is reasonable to exploit this phenomenon to mimic the interaction between GC-PEG-PpIX

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and cell membranes. We measured fluorescence spectra of PpIX and GC-PEG-PpIX in

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deionized water and SDS solution, respectively. As shown in Fig. 2b, the fluorescence

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intensity of GC-PEG-PpIX in water was very weak due to the self-quenching effect of PpIX moieties in the core. Nevertheless, the fluorescence intensity of GC-PEG-PpIX increased

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dramatically (by almost 5- fold) after being treated with 0.5 wt% SDS for 30 min. In

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comparison, free PpIX showed only a negligible fluorescence increase in the presence of SDS, indicating that the aggregates of free PpIX are so condense that they could not be dispersed

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by SDS.

The 1 O2 generation of different samples were then measured using the singlet oxygen

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sensor green (SOSG) kit. SOSG is a 1 O2 -sensitive fluorescence probe and the oxidation of SOSG leads to increased fluorescence emission. We compared the 1 O2 production efficiency

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of PpIX, GC-PpIX, and GC-PEG-PpIX in deionized water and SDS solution upon laser irradiation at 635 nm, respectively (Fig. 2c). As expected, all these samples in water exhibited negligible increase of SOSG’s fluorescence signals, indicating that the 1 O2 generation of PpIX was severely suppressed in this state. After treatment with 0.5 wt% SDS solution, however, the 1 O2 generation of all samples increased as a function of irradiation time due to the weakened self-quenching effect of PpIX molecules. Compared with PpIX and GC-PpIX groups, GC-PEG-PpIX displayed the utmost enhancement of 1 O2 production. This result

ACCEPTED MANUSCRIPT suggested that GC-PEG-PpIX was much easier to disassemble in an SDS solution and achieved a turn-on response of 1 O2 generation. We thus hypothesized that the as-prepared GC-PEG-PpIX NPs would generate activatable and enhanced phototoxicity toward plasma membranes, similar to that in the membrane- mimic SDS environment. Besides, the low 1 O2

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generation of the NPs in water prevented unfavorable side effect during systemic circulation,

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which ensured the safety of the nanoagent.

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According to the results described above, PpIX moieties of GC-PEG-PpIX in unquenched state (treated with SDS solution) showed simultaneous enhancements in fluorescence

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emission and 1 O2 generation upon laser irradiation. As displayed in Fig. 2d, the energy of

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PpIX molecules in the excited singlet state can be dissipated through three typical pathways: (1) FRET mediated self-quenching, (2) fluorescence emission, and (3) 1 O 2 production after

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intersystem crossing. When hydrophobic PpIX moieties form aggregates, the energy in

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excited state is dissipated dominantly by self-quenching. However, when PpIX molecules are separated from each other, the self-quenching effect is significantly suppressed and more excited molecules return to the ground state through the other two pathways, resulting in

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increased fluorescence emission and 1 O2 generation. In this case, fluorescence quantum yield is highly correlated with 1 O2 quantum yield [2]. Therefore, fluorescence as a more convenient and detectable parameter can be utilized to estimate 1 O2 generation. To investigate the anchoring ability of GC-PEG-PpIX on cell membranes, we employed supported lipid bilayer (SLB) and giant unilamellar vesicle (GUV) as model cell membranes. SLB

was

liposomes

prepared

from 1-palmitoyl-2-oleoyl-sn- glycero-3-phosphocholine

labeling

with

0.5%

green

fluorescent

(POPC) lipid

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2e, middle), suggesting GC-PEG-PpIX was strongly adsorbed on the model membrane. We

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hypothesized that the hydrophobic PpIX moieties of GC-PEG-PpIX would anchor on the

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lipid bilayer via hydrophobic interaction. To verify this idea, we further invest igated interactions between GC-PEG-PpIX and GUVs which had the same constituents as SLBs.

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GC-PEG-PpIX was dissolved in an iso-osmolar solution of 0.23 M glucose and gently mixed

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with GUVs at a final concentration of 100 μg/mL. As shown in Fig. 2f, GUVs were also labeled with red fluorescence. In contrast, the fluorescence signal of background was

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considerably lower. This result indicated that GC-PEG-PpIX showed greatly enhanced

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fluorescence when interacting with membranes, implying that the hydrophobic insertion of

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PpIX units into lipid bilayers suppressed the self-quenching effect of the PS molecules.

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Fig. 2. Characterization of GC-PEG-PpIX NPs. (a) Hydrodynamic diameter of GC-PEG-PpIX

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measured by DLS. Insert: TEM image of GC-PEG-PpIX NPs. (b) Fluorescence spectra of free PpIX and GC-PEG-PpIX dispersed in deionized water and 0.5 wt% SDS solution. (c) Singlet oxygen generation of PpIX, GC-PpIX, and GC-PEG-PpIX in water and SDS solution. The concentration of

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PpIX in these samples was fixed at 5 μg/mL. (d) Simplified schematic diagram indicating that the energy of excited PpIX molecules can be dissipated through different pathways. Confocal

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fluorescence images of SLB (e) and GUV (f) were acquired at NBD-PC channel (Ex: 488 nm, Em: 500–550 nm) and PpIX channel (Ex: 552 nm, Em: 605–655 nm), respectively. For SLB, before

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imaging, the green fluorescence-labeled SLB was incubated with GC-PEG-PpIX for 30 min followed by PBS washing.

3.3. In vitro study for plasma membrane-activated PDT

After the model cell membrane studies, we further studied the interactions between GC-PEG-PpIX and cell membranes of live cells. Here confocal fluorescence imaging experiments were carried out (Fig. 3a) for A549 cells treated with GC-PEG-PpIX solutions, while PpIX and GC-PpIX solutions were also studied for comparison purposes (To facilitate

ACCEPTED MANUSCRIPT the comparison, the concentration of PpIX in all the three solution samples was fixed at 5 μg/mL). Different incubation time periods (15 min, 1 h, and 2 h) were used for cell incubation with the above three solutions, and before imaging, no washing treatments were conducted for the reagent treated cells. At 15 min, the cells incubated with GC-PEG-PpIX

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exhibited strong red fluorescence mainly at the cell periphery, clearly outlining the plasma

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membranes. Such an observation is consistent with the above results obtained from studies

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using model membranes. As the incubation time increased, GC-PEG-PpIX can still stably attach to the cell plasma membranes with uniformly labeled red fluorescence. In comparison,

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cells treated with free PpIX and GC-PpIX solutions only displayed weak red fluorescence

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throughout the cytoplasm. To better understand the binding efficiency of GC-PEG-PpIX, we quantified the fluorescence intensity of GC-PEG-PpIX-treated cells at different incubation

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time points by flow cytometry. The result showed that the fluorescence intensity reached its

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maximum within 30 min, which unambiguously confirmed the rapid cell membrane binding of the nanoparticle (Supplementary Fig. 2). Therefore, GC-PEG-PpIX enables stable and specific fluorescence imaging for plasma membranes, which may possibly be attributed to the

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hydrophobic insertion of PpIX into the plasma membrane and the electrostatic attraction interaction between the positively charged GC and the negatively charged plasma membrane. It has been reported that many PS molecules especially lipophilic porphyrins can preferentially partition into lipid bilayers including plasma membranes and intracellular organelle membranes [68,69]. Compared to free PpIX molecules, the GC-PEG-PpIX molecules are more conformationally flexible assisted by the bulky hydrophilic PEG chains, enabling their excellent membrane anchoring affinity. On the other hand, the good membrane

ACCEPTED MANUSCRIPT anchoring effect of GC-PEG-PpIX can be attributed to the steric hindrance of PEG segments which have been reported to be resistant to cellular uptake [70,71]. On the basis of our previous experimental data, the increased fluorescence emission of GC-PEG-PpIX is always accompanied with enhanced 1 O2 production under laser irradiation. Hence, the stable and

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specific fluorescence imaging cannot only display the outline of cells, but also imply

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effective photodynamic damage toward plasma membranes upon laser irradiation.

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It has been reported that PS molecules associated with plasma membranes mainly induce cell necrosis[2] which is considered to be an unprogrammed cell death involving cytoplasm

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swelling, devastation of organelles [65,72], and necrotic blebbing [73]. Benefiting from the

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specific plasma membrane imaging of GC-PEG-PpIX, we then managed to dynamically trace the morphological changes of membranes under continuous laser irradiation. In this

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experiment, GC-PEG-PpIX treated A549 cells were continuously exposed to a 635 nm laser

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at 14 mW/cm2 and observed under a confocal microscope at different irradiation time points (0, 1, 5, 10, and 15 min). As shown in Fig. 3b, slight cytoplasm swelling could be recognized in the bright field after 1 min of laser irradiation, confirming a rapid cellular response to the

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photoinduced injury. Besides, small-sized but clearly visible membrane blebs in the PpIX fluorescence channel were observed (marked by arrows), which could hardly be seen in the bright field. As the irradiation time increased to 5 min, the degree of cell swelling increased with more small blebs around the plasma membrane. Strikingly, evident red fluorescence signals were detected in the cytoplasm, demonstrating the influx of GC-PEG-PpIX into cells. Since the nanoagents were impermeable to non- irradiated plasma membranes for the same incubation time, such an observation implied that the membrane integrity was reduced due to

ACCEPTED MANUSCRIPT photodynamic damage. With the increasing irradiation time, micrometer-sized membrane blebs and highly vesiculated structures emerged in cytoplasm, indicating the significantly enhanced cellular influx of the nanoagents. Based on the above- mentioned cellular fate, we believed that GC-PEG-PpIX induced necrosis- like cell death upon laser irradiation.

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Nevertheless, these results demonstrated that the cell death was not solely due to the damages

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caused by plasma membrane- located PS molecules. Initially, plasma membrane-anchored

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GC-PEG-PpIX molecules activated photoinduced cell membrane damage, causing compromised membrane integrity. According to the literature, ROS can cause peroxidation of

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membrane lipids which consequently leads to increased membrane permeability and even

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pore formation [74]. Therefore, extracellular GC-PEG-PpIX NPs were readily internalized into cells and interacted with cytoplasmic membrane-containing organelles to promote cell

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destruction upon irradiation, which further led to self-enhanced cell influx of PDT agents.

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Collectively, it is reasonable to deduce that the nanoagents achieve photodynamic damage in a synergistic manner: loss of plasma membrane integrity and subsequent intracellular organelle destruction by the PS influx.

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We then evaluated the phototoxicity of GC-PEG-PpIX (5 μg/mL of PpIX) in A549 cells after irradiated with a laser (635 nm, 14 mW/cm2 ) for different irradiation time periods (Fig. 3c). After incubation the cells with GC-PEG-PpIX for another 4 h, the cell counting kit-8 (CCK-8) assay was conducted to measure relative cell viabilities. As expected, GC-PEG-PpIX presented much better cancer cell killing efficiency as compared to free PpIX and GC-PpIX. After 10 min, almost all the cancer cells were killed by the combined use of GC-PEG-PpIX and laser. Furthermore, the cytotoxicities of free PpIX, GC-PpIX, and

ACCEPTED MANUSCRIPT GC-PEG-PpIX without laser irradiation were also determined by the CCK-8 assay, showing negligible cytotoxicity toward A549 cells (Supplementary Fig. 3). In addition, the cell killing effect was evaluated using trypan blue staining. After 5 min of laser irradiation, GC-PEG-PpIX treated cells were stained with trypan blue immediately. Fig. 3d displayed a

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blue circular region that matched the laser spot size, while the cells outside the laser spot still

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remained viable. Considering the fact that trypan blue can only penetrate the damaged plasma

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membrane, the above observation provided another piece of evidence for the photoinduced loss of membrane integrity. Collectively, GC-PEG-PpIX exhibited negligible cytotoxicity and

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remarkable PDT efficacy in vitro, which is promising for in vivo tissue-specific PDT therapy

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without noticeable adverse effects on normal tissues.

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Fig. 3. Confocal fluorescence images of A549 cells and the in vitro PDT results. (a) Confocal fluorescence images of A549 cells incubated with free PpIX, GC-PpIX, and GC-PEG-PpIX (5 μg/mL of PpIX) at different time points, respectively (Scale bar = 25 μm). (b) Real time monitoring of A549

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cells treated with GC-PEG-PpIX (5 μg/mL of PpIX) upon continuous laser irradiation. The white arrows indicate membrane blebs and the white dotted circle shows vesiculated structure in cytoplasm (Scale bar = 10 μm). (c) Viability of treated A549 cells in the dark (0 min) and after laser irradiation for different periods of time (1, 2, 4, 6, and 10 min). Before irradiation, cells were incubated with free PpIX, GC-PpIX, and GC-PEG-PpIX (5 μg/mL of PpIX in all samples) for 15 min. The results were expressed as the mean ± s.d. *P < 0.05, one-way analysis of variance (ANOVA). (d) Trypan blue staining image of A549 cells treated with GC-PEG-PpIX after laser irradiation. The red dotted circle indicates the size and position of the laser spot.

3.4. In vivo imaging-guided PDT for subcutaneous tumors

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Encouraged by the in vitro results, we then tested the feasibility of GC-PEG-PpIX for in vivo fluorescence imaging. Nude mice bearing U14 tumors were intratumorally injected with free PpIX, GC-PpIX, and GC-PEG-PpIX (with the same PpIX content of 4.8 μg),

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respectively, and then imaged by an animal imaging system. The fluorescence intensities of

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GC-PEG-PpIX in the tumor region showed a sharp increase during the first 2 h and a

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decreasing trend after 3 h post- injection (pi) until no detectable signal at 15 h pi (Supplementary, Figs. 4a−b). In contrast, GC-PpIX exhibited relatively low fluorescence

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intensity and fast fluorescence decay, whereas free PpIX only emitted weak fluorescence

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signal. We believed that a direct fluorescence comparison would be reasonable for evaluating local pharmacokinetics since all mice were treated/imaged under the identical condition [75].

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Thus, GC-PEG-PpIX could realize enhanced in vivo fluorescence imaging and prolonged retention time at the tumor site. Inspired by our in vitro confocal imaging results, we

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hypothesized that this advantage of GC-PEG-PpIX might share the similar membrane anchoring mechanism that this nanoagent has strong physical association with plasma

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membranes and emit greatly increased fluorescence. As a proof-of-concept, nude mice were intratumorally injected with free PpIX, GC-PpIX, and GC-PEG-PpIX, and the tumors were homogenized after 2 h to prepare single cell suspensions, followed by confocal imaging immediately (Supplementary, Fig. 5). As expected, GC-PEG-PpIX-treated tumor cells were labeled with observable red fluorescence mainly on plasma membranes, while free PpIX and GC-PpIX groups showed weak fluorescence. This result demonstrated the excellent membrane-anchoring ability of GC-PEG-PpIX at the in vivo level, which was advantageous

ACCEPTED MANUSCRIPT in high-contrast tumor fluorescence imaging. To testify if GC-PEG-PpIX could be used as in vivo theranostics, we also investigated its feasibility for imaging- guided PDT. Nude mice bearing subcutaneous U14 xenograft tumors were intravenously (i.v.) injected with free PpIX, GC-PpIX, and GC-PEG-PpIX (doses: PpIX

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5 mg/kg) and imaged by an animal imaging system at different time points. As for

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GC-PEG-PpIX group, apparent tumor accumulation was observed at 12 h pi and the

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fluorescence signal at the tumor area reached the maximum at 18 h pi (Figs. 4a−b). Although GC-PpIX could also be accumulated in the tumor site, its tumor fluorescence was much

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lower than that of GC-PEG-PpIX. No detectable signal was observed in the free PpIX group.

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In addition, nude mice i.v. injected with GC-PEG-PpIX and GC-PpIX were sacrificed at 3 d and 7 d pi, respectively. Ex vivo imaging revealed that GC-PEG-PpIX distributed mainly in

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tumor and partially in liver at 3 d pi and still retained in tumor at 7 d pi without other organ

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distributions (Fig. 4c). In contrast, GC-PpIX showed a high uptake by the liver and kidneys with relatively poor tumor accumulation. The fluorescence intensities of organs and tumors were quantified in Fig. 4d. The above results unambiguously demonstrated that

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GC-PEG-PpIX achieved excellent tumor accumulation with enhanced fluorescence signal and prolonged retention in tumor site. Besides, the absence of fluorescence signal in other main organs at 7 d indicated that the nanoagent was very safe for in vivo applications since it was not accumulated in these organs. The improved tumor retention performance of GC-PEG-PpIX can be mainly attributed to the presence of PEG chains (which have been proved to reduce the retention of NPs in reticuloendothelial system and prolong the blood circulation time) and its proper size (~82 nm), which enables the enhanced permeability and

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retention (EPR) effect of the drug.

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Fig. 4. In vivo and ex vivo fluorescence imaging. (a) Real-time in vivo fluorescence images of nude mice after i.v. injection of free PpIX, GC-PpIX, and GC-PEG-PpIX, respectively, at different time points. GC-PEG-PpIX shows statistically significant fluorescence increase as compared to PpIX, *P <

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0.05. (b) Average fluorescence intensities of tumor areas from different groups at different time points based on in vivo fluorescence images shown in (a). (c) Ex vivo fluorescence images of major organs and tumor tissues excised from mice i.v. injected with GC-PpIX and GC-PEG-PpIX at 3 d and 7 d pi, respectively. From top to bottom: heart (H), liver (Li), spleen (S), lung (Lu), kidneys (K), and tumor (T). (d) Semi-quantitative biodistribution of GC-PpIX and GC-PEG-PpIX in major organs and tumors at 3 d and 7 d pi determined by the fluorescence intensities in (c).

Motivated by the excellent therapeutic effect in vitro and the high tumor accumulation of GC-PEG-PpIX, we then conducted the in vivo imaging-guided PDT on the U14 subcutaneous

ACCEPTED MANUSCRIPT tumor model. On the basis of aforementioned results, 18 h pi was detected as the suitable time point to carry out PDT. After i.v. injection of saline, free PpIX, GC-PpIX, or GC-PEG-PpIX (dose: PpIX 5 mg/kg), all mice were irradiated with a 635 nm laser (30 mW/cm2 ) for 20 min every other day for a total of three treatments and the tumor volumes

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were measured using a caliper every other day. Remarkably, the tumors on mice treated with

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GC-PEG-PpIX were effectively eliminated after PDT without noticeable regrowth within the

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period of our observation (22 days). In contrast, GC-PpIX revealed a tumor inhibition effect only in the early 10 days, while free PpIX exhibited little therapeutic efficacy (Fig. 5a).

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Hematoxylin and eosin (H&E) staining of tumor slices from GC-PEG-PpIX-treated group

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showed that most cells were severely damaged after treatment, but the tumors of the other groups were only partially or scarcely damaged (Fig. 5c). In addition, no evident body weight

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loss was observed in GC-PEG-PpIX group, suggesting the little toxic effect of this nanoagent

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to animals (Fig. 5b). Systemic toxicity was further evaluated by analyzing the tissue slices from mice administered with GC-PEG-PpIX (Fig. 5d). Compared with the control group, no obvious inflammation, cell necrosis, or apoptosis were observed in the heart, liver, spleen,

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lung, and kidney, indicating the absence of appreciable toxic side effects.

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Fig. 5. In vivo photodynamic therapy. (a) Tumor growth curves of different groups after PDT treatment. U14 tumor-bearing mice were i.v. injected with saline, free PpIX, GC-PpIX, and GC-PEG-PpIX, respectively, and irradiated by a laser (635 nm, 30 mW/cm2 ) for 20 min every other day for a total of three treatments. *P < 0.05, **P < 0.01, ***P < 0.001, one-way ANOVA. (b) Relative body weight curve of different groups of mice after various treatments indicated above. (c) Representative tumor images from different groups taken at the 22nd day (the upper row) and corresponding H&E-stained tumor slices (the bottom row) (Scale bar = 100 μm). (d) H&E-stained tissue slices of major organs in mice treated with saline (control) and GC-PEG-PpIX after 22 days of

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To assess the biocompatibility of the nanoagents more comprehensively, we have also evaluated the hematological toxicity of GC-PEG-PpIX by observing the morphological

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changes of red blood cells (RBCs) and the hemolysis of the RBCs. The optical images of RBCs treated with different concentrations of GC-PEG-PpIX are shown in Fig. 6a. RBCs

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maintained their normal discoid shapes below 250 μg/mL GC-PEG-PpIX, and a portion of

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RBCs changed into echinocytes at 500 μg/mL GC-PEG-PpIX. Such an observation was in

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good agreement with the hemolysis results, which showed good hemocompatibility of

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GC-PEG-PpIX (Fig. 6b).

Fig. 6. Hematological toxicity assessment. (a) Optical images for RBC cells in the presence of various

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concentrations of GC-PEG-PpIX. (b) Hemolysis percentages of RBC cells at different GC-PEG-PpIX concentrations. *P < 0.05, **P < 0.01, ***P < 0.001.

4. Conclusions In summary, an innovative and potent PDT nanoagent based on cell surface engineering was successfully designed and synthesized for imaging- guided cancer treatment. A series of model membrane (SLB and GUV), live cell (A549 cancer cells), and subcutaneous U14 tumor model experiments have demonstrated that the PpIX moieties in GC-PEG-PpIX have

ACCEPTED MANUSCRIPT excellent membrane-anchoring ability and can shift from the initial stacked state in bulk solution to the membrane-bound segregated state. The membrane insertion of PpIX moieties in GC-PEG-PpIX greatly suppressed their self-quenching effect, resulting in significantly increased fluorescence and 1 O2 generation efficiency. It was found that GC-PEG-PpIX could

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rapidly bind to plasma membranes and severely damage the membrane integrity upon

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irradiation, leading to the drastic cell influx of the NPs. Furthermore, we carefully

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investigated the in vivo performance of GC-PEG-PpIX through intratumoral as well as intravenous injection, and the results revealed that the nanoagent achieved high-contrast

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tumor fluorescence imaging, enhanced tumor accumulation, and prolonged tumor retention,

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which facilitated further imaging- guided PDT to effectively ablate subcutaneous tumors. Collectively, this innovative PDT nanoagent has the following advantages: (1) the preparation

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method is very simple and only requires a two-step synthesis from three commercial reagents;

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(2) covalent binding of PpIX to the polymeric construct prevents premature release of PS molecules during systemic circulation; besides, the outer PEG shell ensures the nanoconstruct excellent water solubility/dispersibility under aqueous conditions, which can pre vent protein

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adsorption and increase in vivo circulation time; (3) GC-PEG-PpIX can rapidly attach on cell membranes and achieve wash-free fluorescence imaging for the plasma membrane; (4) the plasma membrane-based PDT can actively increase membrane permeab ility and achieve self-enhanced cell influx of PSs upon laser irradiation, which does not involve either inadequate cellular uptake or the lysosomal escape problem faced by most of the nanodrugs, and can therefore avoid the drug-resistance problem encountered by other conventional cancer therapies; (5) GC-PEG-PpIX with enhanced fluorescence signals at tumor sites and

ACCEPTED MANUSCRIPT prolonged tumor accumulation is advantageous in realizing high-contrast fluorescence imaging assisted PDT which enables effective tumor ablation without regrowth in 3 weeks; (6) Consisting of three safe components (GC, PEG, and PpIX), the GC-PEG-PpIX shows negligible systemic toxicity and good hemocompatibility, enabling its potential clinical

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applications. We believe that this multifunctional PDT nanoagent has a great potential for

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cancer treatment.

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The authors declare no conflict of interest.

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Disclosures

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Acknowledgements

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This work was supported by grants from the National Key Basic Research Program of China (973 Program) (2013CB933904), National Natural Science Foundation of China (21273130 and 21673037),

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Fundamental Research Funds for the Central Universities (2242015R30016), Six Talents Peak Project in Jiangsu Province (2015-SWYY-003), and Scientific Research Foundation for the Returned Overseas Chinese Scholars, State Education Ministry. Z.C. acknowledges the support from the University of Michigan for his sabbatical.

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Graphical abstract