Self-templated, Green-synthetic, Size-controlled Protein

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Biological and Medical Applications of Materials and Interfaces

Self-templated, Green-synthetic, Size-controlled Protein Nanoassembly as Robust Nanoplatform for Biomedical Application Ya Wen, Haiqing Dong, Kun Wang, Yan Li, and Yongyong Li ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.7b19201 • Publication Date (Web): 06 Mar 2018 Downloaded from http://pubs.acs.org on March 6, 2018

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ACS Applied Materials & Interfaces

Self-templated, Green-synthetic, Size-controlled Protein Nanoassembly as Robust Nanoplatform for Biomedical Application Ya Wen, Haiqing Dong, Kun Wang, Yan Li and Yongyong Li * Shanghai East Hospital, The Institute for Biomedical Engineering & Nano Science (iNANO), Tongji University School of Medicine, Shanghai 200092, P.R. China. KEYWORDS:

Protein

nanoparticles;

Self-templated;

Green-synthetic;

Nanoplatform;

Biomedical application; Drug delivery; Immune effect. ABSTRACT: Despite of inherent advantages over synthetic polymer based counterparts, protein nanoparticles remain unsatisfactory in fabrication owing to low size control, usage of toxic crosslinker or organic solvents. This partially contributes the marginal benefits of Abraxane® in clinic. Herein, a green synthetic, size controlled approach was developed to generate stable albumin nanoparticles. Physically packed ovalbumin (OVA) nanoclusters were temporally formed in heat, which was then used as the template to form protein nanoparticle chemically stabilized by intermolecular disulfide network. Exposure of the embedded free thiols within hydrophobic albumin structure and oxidation of them into disulfides (2-3 fold reduction of thiols groups in this process) was identified the key during the process. Fact of structure stable in SDS treatment (hydrophobic destroyer) while fast disassemble in reduction condition (to cleave disulfide)

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validated the disulfide cross-linked mechanism. The developed approach is facile and reproducible with precision size control (from tens to hundreds of nanometers). The approach can be extended to other protein like bovine serum albumin (BSA), underscoring the potentially universal applicability. Further study demonstrated that the resultant protein nanoparticles can be robust nanoplatform for extensive biomedical applications including drug delivery (DOX encapsulation of 5.7%), target bioconjugation or robust immune adjuvant effect.

1. Introduction The increasing attention in utilizing nanotechnology to pharmaceutical research is mainly attributed to its attractive features for drug delivery, bio-detection, imaging, vaccine formulations and the inherent therapeutic nature of some nanomaterials 1-2. Several therapeutic nanoparticle platforms like inorganic nanoparticles, polymeric micelles, liposomes as well as albumin nanoparticles have been applied for cancer treatment 3; other nanotherapies with definite results expected in near future also show great promise in clinical translation 1, 4. For either basic research or clinical potential, nanoparticle platforms have revealed significant interests in numerous medical applications 5-8. Among the various types of nano-sized carriers, polymer-based nanoparticles are especially interesting because of their high flexibility in design and easily chemical modification 9. In particular, proteins have attracted wide attention as material for the formation of nanoparticle 10. They hold several advantages over synthetic polymers in aspects of biocompatibility, toxicity, immunogenicity, stability as well as accessibility. For easy accessibility and excellent biocompatibility, albumin represents the most studied protein nanoparticle for biomedical application particularly for therapeutic delivery. The approaches


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employed for albumin nanoassembly can be divided into two major categories: chemical crosslinking and physical aggregation. In this regard, many techniques were developed, including 1112

: emulsification, desolvation, and thermal gelation. Despite the wide applicability, the particle

fabrication processes of these methods remained unsatisfactory owing to their inherent defects (usage of toxic reagents, poor size control, insufficient stability, complicated fabrication process etc.) 12-16. The established techniques using chemical cross-linkers such as glutaraldehyde represents the most employed approach to yield protein nanoparticle. However, it usually suffers from toxic chemical-crosslinker, e.g. aldehyde, causing complications for potential clinical translation. Moreover, the random cross-linking among albumin macromolecules easily leads to large aggregates and poor size control particularly for sub-100 nm nanoparticle. Additionally, other methods based on heat-induced aggregation usually generate protein nanoasssembly with low stability for weakly hydrophobicity-driven force 17. The emergence of Nab-technology represents a significant step to protein nanoparticle fabrication with clinical importance, based on which the formulation of paclitaxel bound to albumin (Abraxane®) was approved by FDA 18. Yet concerns on stability and size control of this system remained. It was demonstrated by recent experimental results on Abraxane® indicating insufficient stability 19. These limitations may be associated with marginal benefits of Abraxane® in clinical cancer treatment. Furthermore, the complicated and labor intensive fabrication process impedes the broader clinical impact of Nab-technology. Therefore, in spite of the various established techniques, green-synthetic, size-controlled, convenient and stable protein nanoparticle synthesis remains urgently needed. The disulfide bond occurred in intramolecular protein has long been explicitly revealed by protein chemistry, which lays the foundation in maintaining and stabilizing the spatial structure


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of proteins critical for their inherent biological activities. Taking full advantage of this function, it is highly desired to achieve stable protein nanoassembly if one can delicately drive the formation of intermolecular disulfide network within protein nanoparticles. Chemical disulfide cross-linking is expected to largely reinforce the nanoparticle structure and is able to address the critical problems aforementioned, particularly in regards to the mild condition for disulfide formulation. In addition, capability of selective cleavage of disulfide bond inside the cell and hypoxia tumor condition would facilitate bioresponsive properties of protein nanoparticles for tumor selective delivery 20. Bearing in mind the fact that heat-mediated denaturation could expose free thiols of protein by destroying their spatial structure 21, we proposed a self-templated approach for preparation of protein-based nanoparticles. OVA with four free thiols was used as model protein, heat was firstly employed to form temporal protein nanoclusters in which process the embedded thiol groups were properly exposed. After stopping the growth of protein nanocluster by immediate ice water bath, stable protein nanoparticle supported by disulfide network can be generated by following a mild H2O2 oxidation process without using any organic or toxic cross-linker. The underlying mechanism of the nanoassembly was investigated through stability resistance to various protein-perturbing reagents, surface hydrophobicity and quantification of sulfhydryl (SH) group, the sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) as well as circular dichroism spectra. The further biomedical applications were explored in drug carrying capacity, functional modification property and immune effect. The above approach was also extended to other protein like reduced BSA, verifying the potential universal applicability for the as-developed protein-based nanoparticle synthesis. 2. Materials and methods


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2.1. Materials 2-morpholinoethanesulfonic acid (MES), Ovalbumin (OVA), and 3-(2-Pyridyldithio) propionic acid N-hydroxysuccinimide ester (SPDP) were purchased from Sigma-Aldrich. Sodium dodecyl sulfate (SDS), dithiothreitol (DTT), hydrogen peroxide solution (H2O2, AR, 30 wt% in H2O) and 8-Anilino-1-naphthalenesulfonic acid ammonium salt (ANS) were purchased from Aladdin Chemistry Co. Ltd. (Shanghai). RGD-SH was obtained from GL Biochem (Shanghai) Ltd. Dulbecco’s modified eagle’s medium (DMEM), fetal bovine serum (FBS), phosphate buffered saline (PBS, 10 mM, pH 7.4), trypsin, penicillin-streptomycin and 3-(4, 5-dimethlthiazol-2-yl)-2, 5-diphenyltetrazolium bromide (MTT) were obtained from Gibco Invitrogen Corp. Bodipy 650/665-X NHS ester was purchased from Life Technologies. 4,6-diamidino-2-phenylindole (DAPI) was obtained from Beyotime Biotech. Co. Ltd. PE-anti-CD 86, FITC-anti-CD 80, FITCanti-MHC class II monoclonal antibodies were purchased form eBioscience. 2.2. Synthesis of OVA nanoclusters and nanoparticles fixed by intermolecular disulfide network (FNPs) OVA was firstly dissolved with deionized water to obtain OVA stock solution (10 mg/mL), which was then diluted with MES (0.05 M, pH 6) to prepare the working solution (1 mg/mL). The tube was subject to rigorous stirring and sealed. OVA working solution (5 mL) was heated at 70 oC for synthesis of OVA nanoclusters. Size of the nanoclusters was controlled by duration of heat. Heat times of 130 s, 140 s, 160 s, 180 s, 200 s, 210 s and 225 s were employed to obtain 50 nm, 60 nm, 100 nm, 150 nm, 180 nm, 200 nm, 250 nm OVA nanoclusters, respectively. The growth of nanoclusters was stopped by immediately sinking in ice water bath. This process is critical for keeping a particular size with specific heat time, otherwise, aggregation or even


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precipitation will occur due to longer heat treatment. The obtained nanocluster suspension was then slowly introduced into equivalent volume of H2O2 solution with the final concentration in mixture of 1.67 µM. The reaction was conducted under rigorous stirring, RT. Size of the nanoparticles was closely monitored in the whole process. 20 h later, the reaction solution was sufficiently dialyzed (10 mL reaction mixture dialyzed against 5 L deionized water, 4 – 5 times dialysis) to remove MES and excess H2O2. The final suspension was freeze dried (- 60 oC, 0.3 mba) to obtain the OVA FNPs. 2.3. Targeting functionalization by bioconjugation To investigate the functionalized property of the platform, RGD-SH was employed to confer the synthesized FNP (60 nm) targeting ability. 2.18 mg SPDP (stock solution: 20 mM) was introduced into PBS containing 15 mg OVA FNPs under continuous stirring. 0.5 h later, the excess SPDP was removed through ultrafiltration (MWCO: 3000) with PBS. Then 4.94 mg RGD-SH dissolved in PBS was added into above resultant mixture. After 16 h reaction under gentle stirring at 4 oC, the final mixture was subjected to ultrafiltration (MWCO: 3000), and RGD-OVA FNPs were finally obtained. 2.4. Characterizations The particle size and distribution of OVA nanoclusters and FNPs were measured by Dynamic Light Scattering (DLS) (Zeta Sizer Nano ZS, ZEN 3690, Malvern). The morphology was observed by transmission electron microscopy (TEM, Tecnai-12 Bio-Twin, FEI, Netherlands). The conjugation of RGD and formation of disulfide bond in FNPs were measured with X-ray photoelectron spectroscopy (XPS) (PHI5300, 250 W, 14 kV). The conformational change of the


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protein induced by heat treatment and following oxidation was investigated with circular dichroism spectrometer (CD, J-810, JASCON CO., LTD, Japan). 2.5. The biological stability of OVA nanoclusters and FNPs To evaluate the stability of OVA nanoclusters and FNPs, the size change of OVA particles (1 mg/mL) in DMEM with the presence of 10% FBS were measured by DLS at predetermined time points. 2.6. Size alterations under different treatments To explore the formation mechanism of the nanoparticle structure, size of the nanoclusters and FNPs in the presence of 1 % SDS, and 30 mM DTT, alone or in combinations, was measured respectively. All reagents were dissolved in PBS (10 mM, pH 7.4). 2.7. Surface hydrophobicity determination Surface hydrophobicity of OVA molecules, nanoclusters and FNPs was determined with fluorescence spectrophotometer utilizing ANS as the probe referring to previously reported method 22. Protein dispersion was diluted with PBS. 2.8. Quantification of sulfhydryl group and confirmation of intermolecular disulfide bond Sulfhydryl group content of OVA molecules and resultant OVA nanoclusters and FNPs was determined using Ellman’s reagent 23. OVA molecules and resultant nanoclusters and FNPs were suspended in PBS (2 mg/mL) containing 1 % SDS or along with 30 mM DTT for two hours. The mixture was then subjected to


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reducing SDS-PAGE referring to reported method 23 to verify the existence of intermolecular disulfide bonds. 2.9. Investigation on universal applicability To verify the feasibility of the self-templated strategy to other proteins, we transferred this approach to BSA which contains 17 disulfide bonds and 1 sulfhydryl group. Briefly, BSA dissolved in deionized water (40 mg/mL) was incubated with DTT (0.74 mg/mL) at 37 oC for 2 h under slow stirring. Then the solution was taken out to dialyze against the deionized water for 24 h. The reduced BSA was therefore obtained and diluted to 1 mg/mL with MES for the synthesis of BSA nanoparticles referring to the procedure of OVA nanoparticle synthesis. 2.10. The in vitro and in vivo toxicity evaluation of OVA FNPs HeLa cells cultured in 96-well plates at 5 × 103 cells per well were incubated with free OVA FNPs at predetermined concentrations. After 24 h incubation, the MTT assay was conducted to characterize the in vitro cytotoxicity of nanoparticles. For the in vivo toxicity assessment, nine male and female SD rats and nine female HeLa tumorbearing nude mice (tumor volume of 40 – 60 mm3) were divided into 3 groups with three animals of each group, namely PBS group, OVA FNPs with high dosage (28.5 mg/kg to rats, 57 mg/kg to mice) group and low dosage (5 mg/kg to rats, 10 mg/kg to mice) group. The PBS and FNPs were intravenously injected into the animals’ tails every 3 days with 5 medications to rats and 3 medications to mice during the whole treatment process. The body weights were recorded after each medication. 3 days after the final medication, the serum biochemistry assays including the indicators of kidney function (blood urea nitrogen (BUN) and creatinine (CRE)) and hepatic


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function (alanine transaminase (ALT) and aspartate transaminase (AST)) were performed and analyzed. Then the organs of animals were excised to calculate the organ coefficient (weight ratio of certain organ to body) following with the hematoxylin and eosin (H&E) staining to evaluate the possible toxicity of OVA FNPs. 2.11. Drug loading and release behavior from the synthesized FNPs in vitro Drug loading: OVA FNPs (60 nm) were prepared as described above. After interacting for 20 h with H2O2, the reaction mixture was subjected to ultrafiltration (MWCO: 3000) with PBS to remove MES and excess H2O2. DOX·HCL (2 mg/mL, 0.5 mL), as a model anticancer drug, was slowly added into 10 mL freshly synthesized OVA FNPs (0.5 mg/mL) in PBS under constant stirring for 3 h. The final suspension was exhaustedly dialyzed and lyophilized to obtain the DOX@OVA FNPs. Drug release: To investigate the DOX release behavior, lyophilized DOX@OVA FNP was introduced into 15 mL PBS. After homogeneously dispersed, the suspension was divided into three portions in parallel against 100 mL PBS respectively with constant stirring at 150 rpm, 37 o

C. At specific time points, the released DOX in recipient PBS was taken out and measured with

UV spectra at 480 nm. The DOX release profile was obtained with time versus cumulative DOX release. 2.12. Cytotoxicity of DOX loaded OVA FNPs HeLa cells cultured in 96-well plates at 5 × 103 cells per well were incubated with free DOX and DOX@OVA FNPs at predetermined concentrations, respectively. After 24 h incubation, the MTT assay was conducted to determine the cytotoxicity of DOX loaded OVA FNPs.


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2.13. Cellular internalization To examine cellular internalization of free FNPs (60 nm) (RGD-OVA FNPs) and DOX@OVA FNPs, HeLa cells were seeded into a six-well-plate at 1.5 × 105 cells per well for 24 h, and then incubated with bodipy labeled OVA FNPs, RGD-OVA FNPs (the final concentration of OVA of 0.3 mg/mL) and DOX@OVA FNPs (the final concentration of DOX of 3.6 µg/mL) respectively at 37 oC for 4 h. The bodipy labeling was made according to the protocol provided by the manufacture. Briefly, bodipy stock solution of 10 mg/mL was firstly prepared in DMSO, then the reaction was carried out in 0.1 M NaHCO3 solution at room temperature for 1 h with molar ratio 1:1 of OVA (OVA FNP) to bodipy. Free DOX was used as control. The cells were then washed three times with PBS, then fixed with 4 % paraformaldehyde for 10 min, stained with DAPI for 3 min and finally imaged by confocal laser scanning microscopy (CLSM, Nikon, A1R). The cells were pre-incubated 1 h with 60 µg/mL RGD peptide to observe the competition effect of peptide on the RGD-OVA FNPs internalization. To quantitatively determine the internalization of OVA FNPs (60 nm) (RGD-OVA FNPs), HeLa cells were seeded into a six-well-plate at 1.5 × 105 cells per well for 24 h. The cells were incubated with fresh medium containing bodipy labeled OVA FNPs, RGD-OVA FNPs with equivalent OVA concentration of 0.3 mg/mL. After incubation for 4 h at 37 oC, the cells were trypsinized, collected and then resuspended in 0.4 mL PBS. For the competition effect analysis, 60 µg/mL of RGD peptide was pre-incubated for 1 h before introducing RGD-OVA FNPs. Analysis by flow cytometer were then performed. 2.14. Body distribution of OVA FNPs and RGD-OVA FNPs after intravenous injection in vivo


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All animal experiments were conducted complying relevant laws and guidelines. Female nude mice were inoculated subcutaneously with HeLa cells (6 × 106) in the right flank. Two weeks later, when tumors grew to 40 – 60 mm3, the tumor bearing mice were intravenously injected with OVA molecules, OVA FNPs (60 nm) and RGD-OVA FNPs (70 nm) labeled with fluorescent bodipy (0.92 mg/kg of bodipy, 57 mg/kg of OVA) and subsequently tracked by the Maestro IN-VIVO fluorescent imaging system (excitation at 630 nm, emission at 780 nm, CRI, MA) at 0.5 h, 1 h, 2 h, 3h post injection. When the fluorescent intensity of tumor region became rich, the tumors as well as organs were excised and imaged. 2.15. The maturation of bone marrow-derived dendritic cells (BMDCs) under stimulation of OVA FNPs BMDCs were harvested from C57Bl/6 mice as described 24 and incubated with OVA molecules, nanoclusters and FNPs (150 µg/mL) for 24 h on day 8. For the degree of maturation analysis, 1 × 10 6 BMDC were stained at 4 oC for 30 min using PE-anti-CD 86, FITC-anti-CD 80 or FITCanti-MHC class II monoclonal antibodies. Cells were then washed with PBS and analyzed using flow cytometer. 3. Results and discussion 3.1. Synthesis and characterization of nanoparticles fixed by intermolecular disulfide network (FNPs)


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Figure 1. (a) Hydrodynamic diameter of OVA nanoclusters assembled by hydrophobic interaction and (b) FNPs with various sizes resulted by an oxidized “fixation” process. (c) Size alterations upon treatment by 1% SDS (destroyer of hydrophobic interaction). (d) Visual appearance of OVA nanoclusters with various size. OVA, the main protein of hen egg white that contains one internal disulfide bond and four free thiols 17, was employed as model protein. OVA was firstly dissolved in MES and then subjected to heat treatment at 70 oC. Very quickly in the solution, OVA gradually formed nanoclusters during the assembly process within tens of seconds to several minutes as shown in Figure 1d, as shown by the opalescence of the reaction solution. Dynamic light scattering (DLS) measurements showed a size of 50 ~ 250 nm (nanoclusters) dependent on the reaction time (Figure 1a). As expected, the stability analysis demonstrated that nanoclusters, though with


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well-defined size distribution, were mainly stabilized with hydrophobic interaction since SDS (destroyer of hydrophobic interaction) could lead to fast disassembly of nanostructures as demonstrated by the sharp decrease of particle size (Figure 1c). Enlightened by the crucial effect of disulfide in the maintenance of spatial structure of protein, we sought to stabilize nanoparticles through the formation of intermolecular disulfides between proteins via sulfhydryl-disulfide exchange or sulfhydryl oxidation. An optimization strategy was proposed based on the fact that heat-mediated denaturation could expose free thiols of protein 25. H2O2, a clean oxidant for its nontoxic decomposition byproducts, was therefore chosen to oxidize OVA nanoclusters to promote formation of intermolecular disulfide network at ambient temperature. It is interesting to find that, with sufficient oxidation time, OVA nanoparticles with robust stability (FNPs) were structurally retained even after interaction with SDS (Figure 1c). Compared with nanoclusters, the size of FNPs are slightly increased (Figure 1b) (the size volume distribution and polydispersity indexes of nanoclusters and FNPs shown in Figure S1 and Table S1, respectively). It indicates that disulfide ‘fixation’ within nanoclusters had little impact on size change during the gentle oxidation process. FNPs (60 and 180 nm) (DLS data shown in Figure S2) were chosen for further study. 3.2. Mechanistic study for FNP formation


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Figure 2. Stability study treating by destroyers of hydrophobicity (SDS) or disulfide (DTT): (a) Effects of 1 % SDS and 30 mM DTT on the hydrodynamic diameter of OVA nanoclusters and (b) FNPs. (c) Surface hydrophobicity (S0) and sulfhydryl (SH) content of OVA molecules, nanoclusters and FNPs. The data are presented as means ± SD (n = 3), ** p < 0.01. (d) SDS-


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PAGE electrophoresis gels. Lane M: marker proteins. Line a, b and c represent OVA molecules, nanocluster - 60 nm and 180 nm treated with 1 % SDS for 2 h. Lane d, e and f represent OVA molecules, FNP- 60 nm and 180 nm treated with 1 % SDS, and lane d’, e’ and f’ represent treatment of 1 % SDS in combination with 30 mM DTT for 2 h. (e) CD spectra of OVA molecules, nanoclusters and FNPs (0.5 mg/mL). To elucidate the disulfide formation in fixing or stabilizing the nanoparticles, FNPs as well as nanoclusters were treated by DTT, an effective destroyer to disulfide bonds, as well as by hydrophobic destroyer (SDS). As shown in Figure 2b, SDS or DTT alone could not dissociate the nanoparticles since size of both FNPs (60 nm and 180 nm) were well retained after treatment. However, if being co-treated by DTT in combination with SDS, there appeared a complete dissociation with significant decrease in size. This implies the co-existence of synergistic physical and chemical stabilizing forces. For comparison, nanoclusters formed after heat treatment could disassemble in SDS alone (Figure 2a), while DTT exerted no obvious size effect for the absence of disulfide bond. It was reported most of thiol groups located in hydrophobic region. Surface hydrophobicity value was therefore assessed by classic ANS method. Hydrophobicity value of OVA nanoclusters (Figure 2c) is significantly increased compared with that of OVA molecules, indicating the exposure of protein hydrophobic residues upon heat treatment. This was favorable for the intermolecular hydrophobic interaction necessary for nano-assembly. In contrast, the value of FNP decreased probably due to reduced exposure of hydrophobic region for disulfide formation. Size showed pronounced effect on hydrophobicity value for nanoclusters but not so obvious for FNPs. Using Ellman’s reagent, oxidation of nanoclusters significantly reduced the content of sulfhydryl group, suggestive of the transformation of sulfhydryl group into disulfide bond.


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SDS-PAGE assay provided further evidence to support the presence of intermolecular disulfide bonds inside the FNPs. As shown in Figure 2d, the nanoclusters were easily dissociated into monomeric molecules in the presence of SDS. In contrast, OVA FNPs only subjected to DTT treatment dissociated into monomeric molecules as the native OVA molecules, while non-DTT treatment group showed very weak migration band of OVA molecules particularly for FNP 60 nm. Therefore, the dissociation behavior in accordance with the presence of DTT gave further evidence of the presence of intermolecular disulfide bonds in FNPs. In addition, XPS was also employed to verify the formation of disulfide bonds. As shown in Figure S3, the S 2p3 peak located at 163 eV was associated to the presence of disulfide specie, confirming the formation of disulfide bonds in FNPs. The conformational alteration of protein was determined by CD spectra (Figure 2e). Two characteristic peaks located at 208 nm and 222 nm were recognized as typical signals of α-helix structure, the content of which could be employed to inspect the conformational alteration of the protein 26. The CD spectra of OVA nanoclusters and FNPs of different size were similar to that of free OVA molecules, indicating the dominance of α-helix structure in OVA is preserved. 3.3. Size distribution and morphology observation


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Figure 3. TEM images and hydrodynamic diameters determined with DLS of OVA nanoclusters (a and c) and corresponding FNPs (b and d). As shown in TEM micrograph (Figure 3), both OVA nanoclusters and FNPs, the size of which were well maintained after disulfide ‘fixation’, exhibited spherical morphology with uniform size distribution. The precision size control of nanoparticle synthesis in a large size range is of great importance for meeting different applications. Such self-templated new strategy of engineering protein based nanoparticles confers the protein nanoassembly platform with versatile medical application potential. 3.4. Stability analysis


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Figure 4. Stability of OVA nanoclusters (a and c) in PBS and 10 % FBS containing DMEM, and FNPs (b and d) in DMEM containing 10 % FBS at 37 oC. Since synthesized polymer-based nanoparticles have been reported to bring about coagulation or gelation in vivo, it is imperative to determine the stability of the synthesized nanoparticles at 37 o

C in circumstance of 10 % FBS 27, mimicking the physiological environment. As shown in

Figure 4, the OVA nanoclusters were fast disassembled in DMEM containing 10 % FBS as demonstrated by the disappearance of the size peaks (Figure 4a and c) while the size distribution of the FNPs (Figure 4b and d) showed negligible variation after 96 h deposit, implying the more stable property of nanoparticles obtained after oxidation.


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3.5. Method extended to BSA protein

Figure 5. (a) Schematic illustration of preparation for BSA FNP with the developed approach. (b) Hydrodynamic diameter variation of BSA FNPs in the presence of 1 % SDS and 30 mM DTT alone or in combinations. We also transferred this nanoparticle preparation approach to BSA to verify the potential universal applicability. BSA was chosen for it contains 17 disulfide bonds while only 1 sulfhydryl group, which is very different from that of OVA. Therefore, DTT, a reducing agent, was firstly employed to interact with BSA to expose sufficient sulfhydryl groups. The reducing BSA was subsequently used to prepare BSA nanoparticles in accordance with the method described above (Figure 5a). The self-templated, green synthetic approach can also generate well-defined size of BSA nanoparticles with desired stabilization capability (Figure 5b), suggesting the universal applicability of this protein nanoassembly method. 3.6. Toxicity assessment


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Figure 6. Cytotoxicity of synthesized OVA FNPs against HeLa cells. The in vitro cytotoxicity of OVA FNPs with different size was assessed employing MTT assay against HeLa cells. As shown in Figure 6, the viabilities were all over 90 % and even 100 % at high concentrations. The systemic toxicity of synthesized OVA FNPs was further investigated using healthy SD rats and HeLa tumor-bearing nude mice. Serum biochemistry indicators, organ coefficient (weight ratio of certain organ to body) as well as the body weight variation of the animals injected with different dosages of FNPs were measured. In addition, a histological analysis of organs was also carried out to appraise possible damage of tissue, inflammation, or lesions by FNPs. The results demonstrated the negligible toxicity of OVA FNPs (shown in Table S2, S3, S4, S5 and Figure S4, S5), indicating the great potential in biomedical application. 3.7. OVA FNP as the nanoplatform for drug vehicle


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Figure 7. (a) Cumulative DOX release behavior of DOX@OVA FNPs at 37 oC in PBS of pH 7.4. (b) Cytotoxicity of DOX@OVA FNPs against HeLa cells. (c) CLSM images of HeLa cells at 4 h post-incubation with bodipy labeled DOX@OVA FNPs. For potential application of OVA FNP as a drug carrier, insoluble DOX was selected as a model anticancer drug. The resultant DOX@OVA FNP was about 70 nm in size with DOX loading content of 5.7 %. DOX loading content is determined by the amount of DOX in DOX@OVA FNPs versus the feed amount of DOX@OVA FNPs. The relative low DOX encapsulation efficiency is presumably due to the DOX loading after the nanoparticle synthesis in which process drug can be only loaded through passive diffusion into the inner nanoparticle. Hopefully, loading degree could be improved if it is able to load the drug during the process nanoparticle synthesis which will be explored in our further study. The DOX release behavior was measured


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in PBS at 37 oC as exhibited in Figure 7a. The cumulative DOX release ratio elevated during the initial 12 h, kept a steady rise in the following 60 h, and eventually leveled off. Furthermore, the cytotoxicity of DOX@OVA FNPs was evaluated using the MTT assay. As shown in Figure 7 b, the DOX@OVA FNPs exhibited enhanced inhibitory effect parallel with that of free DOX against HeLa cells with the increase of DOX concentration owing to the efficient cellular internalization (Figure 7c, the internalization of free DOX was shown in Figure S6). The controlled drug release behavior and efficient cancer cell killing ability of drug loaded nanoparticles demonstrated OVA FNP a promising candidate as drug vehicle. 3.8. OVA FNP as the nanoplatform for targeting imaging via bioconjugation


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Figure 8. (a) CLSM images of HeLa cells at 4 h post-incubation with bodipy labeled nanoparticles of OVA FNP (60 nm), RGD-OVA FNP (70 nm) and RGD-OVA FNP preincubated with RGD for 1 h. (b) Flow cytometry analysis showing profiles of different formulations in HeLa cells. (c) In vivo fluorescent images of xenograft mice administrated with bodipy labeled OVA molecules, OVA FNP and RGD-OVA FNP. Red circles indicate tumor sites. (d) Fluorescent images of excised organs and tumor tissues 1 h after the injection of OVA molecules and FNP. To examine the targeting modification property of the synthesized OVA FNPs, RGD, a short peptide that specifically recognizes and interacts with integrin αvβ3 receptors expressed in HeLa cells, was conjugated to the protein nanoparticles (RGD-OVA FNPs) followed by labeling with bodipy. The negligible variation of fluorescence intensity of bodipy in PBS in 3 days demonstrated the stability of fluorescence property of labeling (shown in Figure S7). The conjugation of RGD was verified with XPS, as shown in Figure S8, the peak value of 284.6 ± 0.1 eV is attributed to the C 1s of C=C, and the increase of the intensity of C 1s in RGD-OVA FNPs could be ascribed to the integration of phenylalanine, indicating the successful conjugation of RGD. The targeting effect was estimated using CLSM. As shown in Figure 8 a, compared with OVA nanoparticles, HeLa cells incubated with RGD-OVA FNPs exhibited obvious increase of red fluorescence. However, after free RGD pre-incubation for 1 h, the bodipy fluorescence of RGD-OVA FNPs dramatically decreased. These observations were also quantitatively measured by flow cytometry (Figure 8b), where the mean fluorescence intensity of RGD-OVA FNPs was much higher than that of OVA FNPs and could be reduced by pre-incubation with free RGD, implying the modification of RGD could efficiently enhance the cellular uptake. The efficient


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uptake of RGD-OVA FNPs was mainly ascribed to the specific binding of RGD to integrin receptors expressed on HeLa cells. To investigate the in vivo tumor targeting property of RGD functionalized nanoparticles, time dependent in vivo biodistribution of OVA and RGD-OVA FNPs was visualized employing bodipy fluorescence signal in tumor-bearing mice. As exhibited in Figure 8c, significant fluorescence accumulation at tumor site was found at 0.5 h after the injection of RGD-OVA FNPs, reaching the maximum value at 1 h, and then suffering a sluggish decrease over 2 h. While for the OVA FNPs, the accumulation process at tumor region was relatively slow and weak in 1 h although the utmost fluorescence intensity was also achieved. Furthermore, the elimination profile was relatively fast. These results indicated that the synthesized OVA FNPs could be efficiently targeted to tumor site owing to the desired particle size (sub-100 nm) which helps to effectively repel the uptake of reticuloendothelial system (RES). The targeting property was further intensified by the assistance of integrin-mediated endocytosis because of RGD modification. In vitro excised tumor from RGD-OVA FNPs treated mice (Figure 8d) exhibited much stronger fluorescence intensity than that of OVA FNPs (the quantitative fluorescence intensity was shown in Figure S9), further confirming the passive tumor targeting and integrinmediated active targeting property of OVA FNPs. In contrast, tumor selective accumulation of OVA molecules was weak. Therefore, the desired targeting modification properties could be achieved with the synthesized OVA FNPs, showing the promise of the protein nanoassembly platform in target imaging applications. 3.9. FNP as the nanoplatform for immune adjuvant


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Figure 9. Effect of OVA molecule, nanocluster and FNP with different size on the expression of maturation markers CD 86 (a), CD 80 (b) and MHC II (c) of BMDC in vitro by flow cytometry. (d) The median fluorescence intensity (MFI) of maturation markers after different treatment. All data were presented as mean ± SD (n = 3). It is well-known that dendritic cells (DC) as professional antigen presenting cells play pivotal roles in the initiation and regulation of innate and adaptive immunities. The activation and maturation of DCs is an essential process to trigger effective immune responses 28. We therefore evaluated the immune potency of OVA FNPs, as OVA were broadly utilized as model antigen in molecules, through promoting the maturation of BMDCs by maturation markers: CD86, CD80 and MHCII. The results showed both OVA nanoclusters and FNPs could in some extent upregulate the expression of CD86, CD80 and MHCII (Figure 9), boosting the maturation of


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BMDCs. Given the superior biological stability, OVA FNPs may be an ideal candidate for immunology applications. Above results can also inspire one to choose tumor-associated proteins to form nanoclusters or FNPs, which constitute only tumor associated antigen with considerably high antigen density due to absence of exterior nanocarrier. This work is currently underway to explore novel formulations of nanovaccine. 4. Conclusions A novel approach was developed for the fabrication of protein-based drug delivery systems. Exposure to oxidizing reagent is employed to facilitate the formation of intermolecular disulfide network of heat-mediated protein nanocluster, resulting in the stable protein nanoassembly. Different from previous reported templated nanoparticle synthesis approaches and stabilization methods, we explored a self-templated protein nanoassembly technique followed by disulfide fixation. The preparation process is totally green, without the use of any organic solvents and requiring no cross-linking agents or additional surfactants to form stable nanoparticles. It was demonstrated in our model system that OVA-based FNPs can successfully deliver DOX with a controlled release profile, be functionally modified with RGD, facilitate obvious targeting property, and act as useful immune adjuvant. Moreover, the approach is not limited to OVA as only particle material, revealing the potential to be universally applied to proteins with certain amount of sulfhydryl groups. This study would help to open up new technological and pharmaceutical innovations for the delivery of drugs. ASSOCIATED CONTENT Supporting Information


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Size volume distribution of OVA nanoclusters and FNPs; the polydispersity indexes of OVA nanoclusters and FNPs with different sizes; XPS S 2p spectra of nanocluster and FNP; blood chemistry results for rats and female HeLa tumor-bearing nude mice injected with OVA FNP; organ coefficient of rats and female HeLa tumor-bearing nude mice injected with OVA FNP; in vivo toxicity evaluation of OVA FNP, including variations of weight and H&E stained images of major organs from rats and female HeLa tumor-bearing nude mice treated with OVA FNP; CLSM images of HeLa cells at 4 h post-incubation with free DOX; stability of fluorescence property of bodipy labeling in PBS; XPS C 1s spectra of OVA FNP and RGD-OVA FNP; quantitative measurements of the body distribution of particles in tumors and different organs. AUTHOR INFORMATION Corresponding Author * E-mail: [email protected]. Notes The authors declare no competing financial interest. Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. ACKNOWLEDGMENT This work was financially supported by research grants from the National Natural Science Foundation of China (NSFC51773154, 31771090, 51473124 and 81571801), Shanghai Natural Science Foundation (17ZR1432100) and Young Hundred-Talent Program of Tongji University.


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Table of Contents artwork


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Self-templated, Green-synthetic, Size-controlled Protein

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