Serum Albumin

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Bioprosthesis of Core-Shell Gold Nanorod-Serum Albumin Nanoimitation: A Half-Native and Half-Artificial Nanohybrid for Cancer Theranostics Hsien-Ting Chiu, Chung-Hao Chen, Meng-Lin Li, ChengKuan Su, Yuh-Chang Sun, Chi-Shiun Chiang, and Yu-Fen Huang Chem. Mater., Just Accepted Manuscript • DOI: 10.1021/acs.chemmater.7b04127 • Publication Date (Web): 13 Dec 2017 Downloaded from http://pubs.acs.org on December 13, 2017

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Chemistry of Materials

Bioprosthesis of Core-Shell Gold Nanorod-Serum Albumin Nanoimitation: A Half-Native and Half-Artificial Nanohybrid for Cancer Theranostics Hsien-Ting Chiu, † Chung-Hao Chen, ‡ Meng-Lin Li, ‡ Cheng-Kuan Su,§ Yuh-Chang Sun, † Chi-Shiun Chiang, † and Yu-Fen Huang*,† †

Department of Biomedical Engineering and Environmental Sciences, National Tsing Hua University, Hsinchu 30013, Taiwan, R.O.C.

‡

Department of Electrical Engineering, National Tsing Hua University, Hsinchu 30013, Taiwan, R.O.C. §

Department of Bioscience and Biotechnology, National Taiwan Ocean University, Keelung, 20224, Taiwan, R.O.C. Keywords: biomimetic nanomaterials, protein corona, native and artificial properties, cellular uptake, high drug loading capacity, photoacoustic nanoamplifiers, combinational therapy.

ABSTRACT High concentrations of aldehyde-based crosslinkers have been commonly used for protein immobilization to facilitate microscale and nanoscale observations. This fixation maintains cell morphology and partial protein activity. In this study, a facile one-step strategy based on a similar concept was first developed for the bioprosthesis of a uniform core-shell gold nanorod/serum albumin (NR@SA) nanoplatform. The resultant albumin shell preserved half of its native form, leading to decreased free SA adsorption, and even these adsorbed proteins were close to their native form. This strategy efficiently prevents subsequent adsorption cascades of other proteins and has a remarkable influence on cellular uptake (of macrophages and tumor cells). Furthermore, the other,

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artificial part endowed NR@SAs with higher drug loading capacity and enhanced photoacoustic signal intensity for cancer theranostics compared with its pristine counterpart. These findings suggested that preserved fidelity and artificial characterizations provide a new perspective for biomimetic nanomaterial design.

INTRODUCTION Albumin and gold nanorods (Au NRs) have emerged as a versatile carrier for the systematic delivery of therapeutic and diagnostic agents. The former one is one of the naturally occurring, principal transport plasma proteins that is nontoxic, biocompatible, and biodegradable. An increasing number of therapeutically active drugs are currently designed possess a high binding affinity for albumin to improve their pharmacokinetic profiles with a prolonged stability and half-life in body circulation for solid tumor targeting.1-3 The latter one, gold nanorods (Au NRs) has exhibited great popularity and success in tumor regression in mouse models through photothermal therapy (PTT).4-5 In addition, the large surface area of Au NRs also makes them a versatile scaffold for efficient delivery of cancer chemotherapeutics. The combination of hyperthermia and chemotherapy usually enhances anti-tumor activities with results that are greater than the sum of individual treatments alone.6-7 Moreover, photon energies that have been absorbed by Au NRs under short laser pulses can be efficiently converted into heat in a localized volume, leading to rapid thermal expansion and contraction and the generation of pressure transients. The outgoing thermoacoustic wave can be 2


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detected with an ultrasound transducer and then used to reconstruct photoacoustic (PA) images. Few studies have further reported that Au NRs modified with silica8 or graphene oxide9 as core-shell nanoparticles exhibit greater enhancement in photoacoustic signals than their pristine counterparts, leading to a better imaging capability for in vivo studies. Physical adsorption and desolvation/crosslinking are two common strategies used to successfully combine Au NRs with serum albumin (SA). By using these methods, serum proteins or pure serum albumin can be loaded onto a gold surface to form a large corona agglomerate containing several Au NRs.10-14 Generally, a small concentration of crosslinker, glutaraldehyde, is required in desolvation methods to stabilize nanoparticles through the crosslinking of resoluble SA.15 The soft corona influences the resultant particle size, stability, and drug release because of rapid replacement by the bioenvironment.10-11 However, to achieve a desirable therapeutic outcome with in vivo animal studies, high-quality control of products with homogeneity in particle size, shape, and composition is highly demanded.16 Similarly, these requirements have been reported to be difficult in many studies on the bioformulation of nanoparticles. The selected biomimetic materials range from one single type of protein to complex microorganisms.17 Another unpredictable concern is that the protein corona is expected to be surrounded by these nanoparticles on encountering a biological fluid. The interplay of the protein corona and nanoparticles and the resultant characterizations of the protein corona (e.g. masking effects and immune system activation) strongly determine the final in vitro and in vivo outcomes.16, 18-19 3



Pegylation and the design of biomimetic materials (including albumin-based nanoparticles) are the most common strategies for incorporation within drug delivery systems to enhance the efficacy of nanoparticle accumulation in tumors.20-22 However, the delivery mechanism between the pegylation strategy and biomimetic-based delivery system is different because biomimetic materials compared with pegylated nanoparticles should have no or less natural interaction with biological fluids or cells, suggesting the distinctive behaviors of biomimetic materials that have specific relations with biological fluids, thereby reducing nonspecific immune cellular uptake (e.g. macrophage uptake) and enhancing tumor accumulation.23-24 However, the mechanism underlying the interplay between biomimetic nanomaterials and biological fluids or cells remains poorly understood. Furthermore, every artificial imitation has always inevitably had some flaws. Moreover, each mimicked part cannot be exactly the same as its native counterpart, which may help explain the incomplete adjustment of these systems to in vivo environments with less notice by immune system.17, 23, 25 The development of additional strategies for rectifying artificial defects or the potential use of these defects to convert disadvantages into advantages may provide these materials worthy of further investigation. This study developed a facile and controllable synthesis method for the successful fabrication of a homogeneous core-shell Au NR–SA nanoparticles (NR@SA, GTA system). Compared with the previous albumin-based nanoformulation,15 the present formulation used a relatively high ratio of glutaraldehyde to protein in order to directly immobilize albumin and form a bioprosthetic shell 4


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outside the Au NRs without requiring a predenaturing process. The fabricated NR@SAs were compared with other denaturing-based NR@SAs (EM system) with regard to their interaction with high protein concentrations in biological fluids and cellular response induction. Notably, the fabricated NR@SAs (GTA) retained more native characteristics, resulting in a low amount of protein adsorption. The composition of the protein shell and adsorbed protein corona influenced the distinct outcomes of macrophage and tumor cell uptake. Moreover, further investigation of the in vivo adsorption of three blood proteins—bovine SA, transferrin (Tf), and fibrinogen (Fib)—revealed that albumin is the determining factor that cooperates with the adsorption of Tf and Fib. Our results indicated that the bioprosthetic shell of the fabricated NR@SAs (GTA) that has less initial interaction with SA can further reduce nonspecific protein adsorption, leading to less protein corona formation. In general, biomimetic materials extracted from an original source are complicated, and the amount of product that can be extracted is very small.24 Therefore, the present study performed simple one-step fabrication of the bioprosthetic SA shell to provide another perspective of biomimetic nanomaterial design. Furthermore, although the artificial crosslinking process might have been responsible for the loss of the native characteristics of the SA shell, it achieved high drug loading capacity, maintained colloidal stability for elevated drug release efficiency, and enhanced the photoacoustic signal intensity compared with its pristine counterpart. Finally, the drug-loaded NR@SAs achieved successful in vivo tumor growth inhibition through combined photothermal therapy and 5



chemotherapy followed by near-infrared (NIR) light exposure. An improved survival rate and less systemic side effects were observed in prostate tumor-bearing mice who received combined therapy compared with those who received conventional drugs or individual treatment.

RESULTS AND DISCUSSIONS Cross-linked effect: One-step synthesis and characterizations of NR@SA nanoplatform In the beginning, to construct a homogenous protein shell, a serial concentration of the cross-linker GTA (0–0.1 M) was acquired for the preparation of NR@SAs to gain more insight into the effect of cross-linking on the formation of core-shell nanocomposites (GTA:SA = 0-33300). The UV-Vis studies (Figure S1A) displayed colloid stability of the resulting nanoparticles at a relatively high GTA concentration, while the plasmon band of the raw Au NRs was completely lost due to aggregation (a pellet is observed). This result indicates that a sufficient amount of cross-linker (0.1 M) was required for rapid cross-linking between SA molecules and that consequent steric hindrance was responsible for the dispersibility of NR@SAs. An analogous result was also obtained with NR@SAs when the CTAB concentration of the dispersions was adjusted from 40 µM to 13.3 µM prior to SA coating (Figure S1B). However, at a lower CTAB concentration, the NR@SAs (line 0.1 M GTA in Figure S1B) were relatively unstable. This suggests that CTAB surfactant plays a crucial role in adequate protein adsorption and consequently determines the colloidal stability of NR@SAs. This result was further confirmed by fluorescence labeled SA (Figure S1C), showing that a 6


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decreasing CTAB concentration with low GTA concentration leads to a reduced level of protein adsorption; Even under high concentration of GTA was benefited for SA loading, the lower CTAB concentration of Au NRs was inefficient to concentrate SAs on the surface, indicating the more random cross-linking resulted in unsatisfied colloidal stability of nanoparticles. To ensure long-term colloidal stability, GTA and CTAB concentration of 0.1 M and 40 µM were selected for NR@SA fabrication and subsequent experiments (Figure S2). Besides, successful surface passivation was detected through a slight red-shift and band broadening of both the transverse and longitudinal surface plasmon resonance bands of NR@SAs over the CTAB-coated NRs in UV-Vis spectra (Figure 1A). The dynamic light scattering (DLS) experiments revealed an obvious increase in the hydrodynamic size of final product, NR@SAs over CTAB-coated NRs (Figure 1B). The formation of a protein corona at the surfaces of NRs was further confirmed by the surface charge changes; the surface zeta potential of CTAB-coated NRs and NR@SAs was +21.5 (± 0.8) mV to −20.1 (± 2.9) mV, respectively (Figure 1C). In addition, negative staining transmission electron microscopy (TEM) images of the resulting NR@SAs also confirmed the core-shell morphologies with an average SA corona thickness of 11.5 nm (Figure 1D). Meanwhile, another core-shell gold nanorods-albumin nanoparticles by desolvation methods that was introduced from the previous study was also fabricated.26 The similar protein loading amount, size, surface charge and morphology of nanoparticles was a good candidate served as a negative control for the following experiments (Figure 1). 7



Bioprosthetic albumin shell reduced phagocytosis by macrophages but enhanced tumor cell uptake After contact with a biological milieu, a nanoparticle surface undergoes biotransformation with protein corona formation before the cells see. Therefore, we first investigated protein adsorption under various concentrations of fetal bovine serum (FBS). Sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) revealed that NR@SAs (GTA) in a 10% FBS environment (Dulbecco’s Modified Eagle Medium [DMEM] + 10% FBS + 1% PS, DFP) adsorbed relatively fewer multiple proteins (Figure 2A). To quantitatively analyze the FBS bound to nanoparticles, FBS was covalently linked to Cy 5.5 prior to the adsorption tests. After the analysis of each collected supernatant, the GTA system was found to adsorb less FBS at FBS concentrations of 10%–50% than the EM system (Figure S3A). These results were further validated using microscale thermophoresis (MST), suggesting that NR@SAs (GTA) are less prone to interact with FBS than NR@SAs (EM) (Figure S3B). To further demonstrate the mechanism of FBS adsorption on the two nanosystems, DLS was used to measure the corresponding size profile (Figure 2B).27 Although both types of nanoparticles were stable after FBS adsorption, the results showed that NR@SAs (EM) with large protein corona formation had a 50.1 ± 13.7 and 3.8 ± 3.1 nm increase in the first and second distribution of DLS profile, respectively, whereas NR@SAs (GTA) had only a 22.6 ± 7.7 and −1.1 ± 2.1 nm increase (Figure 2C). 8


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We hypothesized that the composition of adsorbed FBS and the SA shell of NR@SAs have a distinctive influence on intracellular uptake (Figure 2D). Therefore, both types of NR@SAs (GTA and EM) were used to explore the relationships among the constructed SA shell, adsorbed FBS, and cells. Two cell types, RAW 264.7 (macrophages) and Tramp-C1 (prostate cancer) cells, were selected as the model cells because although the designed nanoparticles are designed to target nonphagocytic cancer cells, they are mainly removed by phagocytic cells from the reticuloendothelial system (RES) before they come into contact with cancer cells. Prior to the in vitro experiments, both nanoparticle types were immersed in different biological buffers for 24 h to examine whether the constructed shell would be replaced by other external proteins because this phenomenon could have influenced cellular uptake.28 The results (Figure S4A) revealed that the crosslinked SA shell of the NR@SAs (GTA) acted as a hard protein corona that was not replaced, whereas the denatured NR@SAs (EM) had a soft protein corona, in which approximately 25.8% ± 2.5% of the SA shell was replaced with FBS. Nevertheless, colloidal stability was not affected by this protein replacement (Figure S4B), and the level of replacement was not sufficient for possible cellular uptake.28 Dark-field microscopic images and inductively coupled plasma mass spectrometry (ICP-MS) were employed to analyze intracellular uptake. More NR@SAs (EM) were internalized by macrophages than were NR@SAs (GTA) (Figure 2E and 2F). Preincubation with phagocytosis inhibitor (cytochalasin D) reduced the cellular uptake of NR@SAs (EM) by approximately 50%; however, the uptake of NR@SAs (GTA) under the same incubation conditions was not affected by 9



the inhibitor. Furthermore, a competition test with free SA, implemented through the microscopic analysis of intracellular fluorescence, demonstrated that both types of NR@SAs (GTA and EM) substantially competed with tetramethylrhodamine (TRITC)-labeled SA for cellular uptake; however, more severe competition was exhibited by NR@SAs (GTA) (Figure S5, 1.2 nM nanoparticles). This result demonstrated that the mechanism of intracellular uptake of NR@SAs (GTA) by macrophages is mainly mediated by the increased levels of the clathrin-dependent receptor, indicating that the fabricated SA shell after GTA crosslinking or adsorbed SA from FBS can retain native SA characteristics. By contrast, the higher FBS adsorption of NR@SAs (EM), which possibly contained only a few native forms of adsorbed SA, allowed nanoparticle internalization by macrophages through endocytosis, and mostly covered denatured SAs induced the phagocytosis pathway. With regard to tumor cell uptake (Figure 2E and 2G), both types of NR@SAs (GTA and EM) exhibited a comparable level of competition with free SA (Figure S5), indicating that both nanoparticles have a similar interaction with the targeted receptor, SPARC receptor.29 However, considering the SA load (surface-modified SA shell plus SA adsorbed from FBS, Figure S4A) on both types of nanoparticles, the bioprosthetic shell of the NR@SAs (GTA) was more efficient for tumor cell uptake. Although denatured SA nanoparticles could target SPARC and gp 60 receptor for drug delivery in cancer therapy,22 the SPARC receptor is more sensitive to native SA than conformationally altered SA,30-31 even if NR@SAs (EM) were docked on a SPARC receptor, conformational changes in the SA structure reduced the rate of subsequent cell internalization, 10


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thereby reducing the uptake of NR@SAs (EM). By contrast, NR@SAs (GTA) possibly preserved more native SA and was therefore more efficient for tumor cell uptake. Notably, surface functional groups are another possible key factor that determines the protein corona composition and resultant cellular uptake.32 NR@SAs (EM) preserved more amine/carboxyl groups than NR@SAs (GTA) after fabrication (Figure S6, w/o wash). Unavoidable protein adsorption on both nanoparticles during purification could balance the loss of functional groups during fabrication (Figure S6, w/ wash); some of the proteins adsorbed during the fabrication process or during the incubation with culture medium were attached to the amine groups on NR@SAs (EM) and were possibly closer to their denatured form, whereas adsorbed proteins on NR@SAs (GTA) were closer to their native form.32 To determine the structural integrity of the constructed protein shell and adsorbed protein, Raman spectroscopy, which assesses single molecular sensitivity, was used to identify their secondary structure. The few obvious peaks of native albumin referred to amide III (1239 and 1274 cm−1), amide I (941 cm−1) and CH (1320–1340 and 1450 cm−1) deformations, and aromatic amino acids (1004 cm−1; Figure 3A).33 The crowded effect caused an obvious red shift in the amide I (1663 cm−1) peak, which should have been at 1650–1655 cm−1, because of the high SA concentration under dried conditions.34 By contrast, the strong enhanced SERS peaks related to the Phe plus Tyr regions within 1585–1620 cm−1 and the CH structure (1340 cm−1) in addition to the largely decreased amide I signal were considered hot spots because of interaction with the hydrophilic CTAB-coated Au NRs, inducing protein unfolding and partial or complete conformational changes.35 After fitting the Raman 11



spectra, the stimulated peak for the NR@SAs (GTA) and NR@SAs (EM) at approximately 1674 cm−1 indicated that the modified SA shells of both nanoparticle types were denatured.36 However, the constructed shell (GTA) retained some of its native characteristics related to the amide III (1222 and 1269 cm−1) and partial amide I (941 cm−1) structures, including an α helix and β sheet (red and blue arrows), whereas the fabricated shell (EM) only exhibited the signal of an α helix (1265 cm−1), indicating prevalent β sheet aggregation in the shell. These results were further confirmed through the circular dichroism (CD) spectra (Figure 3B), which demonstrated that the GTA shell with its partial α helix and β sheet structure was similar to that of previously reported albumin physically adsorbed on Au NRs.37 By contrast, NR@SAs (EM) retained relatively less of the native structure of SA. To demonstrate the protein structure of the resulting protein corona on both types of nanoparticles after further FBS incubation, two constructed shells were incubated with 0.6 wt% SA that comprised approximately equal amounts of the total protein in 10% FBS for 6 h. Two protein-adsorbed nanoparticles were obtained after centrifugation to remove the unbounded free protein and were suspended in PBS buffer. The constructed shell (GTA) not only still retained its amide III structure (1235 and 1270 cm−1) and partial amide I structure (941 cm−1; Figure 4A) but also the adsorbed SA further contributed to the loss of amide I structure at approximately 1665 cm−1 (Figure 3A), suggesting that the adsorbed SA had more resemblance to its native form. By contrast, despite a large amount of SA adsorption, the denatured SA shell (EM) did not exhibit a signal peak for β sheet and 12


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remain amide III structure at approximately 1674 cm−1, indicating that the adsorbed SA was present in a denatured state. Similar results were obtained from CD spectra analysis, which demonstrated that more native SA was bound to NR@SAs (GTA), whereas more denatured SA was adsorbed onto NR@SAs (EM) (Figure 3B). Although according to the concept of protein corona and protein structure analysis (Figure 3A and 3B), the first adsorbed SA on NR@SAs (EM) should be denatured and then covered with more native SA, the centrifugation process possibly removes the outer layers of weakly bound native SA, and only more strongly bound denatured SA would remain in the inside layers. Overall, the biomimetic shell induced less protein corona formation, and the formed protein corona was partially maintained in its native state, which reduced nonspecific phagocytosis by macrophages and enhanced tumor cell uptake efficiency (Figure 3C). By contrast, protein corona formation did not, as expected, mask the entire denatured protein shell (EM). Instead, the shell induced conformational changes, leading to the formation of a denatured adsorbed protein that enhanced macrophage uptake. Despite supposedly having weakly bound native SA in the outside layers, most of the denatured protein reduced the efficacy of nanoparticles for tumor cell internalization (Figures 2F and 2G and S5).

Bioprosthetic albumin shell reduced protein corona formation by blood proteins

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Three blood proteins—bovine albumin (SA), transferrin (Tf) and fibrinogen (Fib)—were modeled for the evaluation of nanoparticle–protein interaction because they are predominant plasma proteins.38-39 Protein adsorption was first investigated using a single protein model for the identification of distinct molecular recognition events. The prepared SA, Tf, and Fib protein solutions were individually mixed with NR@SAs (GTA or EM) and measured through MST analysis under a thermal gradient (Figure 4 A and 4B). As expected, the bioprosthetic shell (GTA), which is closer to the native albumin form, exhibited a relatively higher dissociation constant (KD) for free SA adsorption. Furthermore, the Tf adsorption curve revealed two binding sites for asymmetric Tf binding on NR@SAs (GTA),40 indicating that the bioprosthetic shell generally contained two types of SA (closer to its denatured and native states), supporting the results presented in Figure 3. The native SA was possibly responsible for a lower KD (KD1), which is consistent with previous findings that supported the slight interaction between SA and Tf.41-42 By contrast, free SA had two binding sites with a lower KD for the interaction with NR@SAs (EM),43 suggesting a strong interaction between native SA and the denatured SA shell. Moreover, the mostly covered denatured SA shell remained only one Tf binding site. Nevertheless, only the nonspecific binding of Fib on the two nanosystems suggested that both types of NR@SAs maintained satisfactory hydrophilicity and had a low tendency for hydrophobic interaction.44 To obtain detailed information on the behaviors of NR@SAs under in vivo conditions, series concentrations (including in vivo conditions) of single or mixed protein solution were incubated with 14


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NR@SAs (Figure 4C–4E) and the loading amount was then quantified. For the single protein adsorption profile (Figure 4C–4E, black line), NR@SAs (EM) exhibited a larger amount of SA and Tf than NR@SAs (GTA), particularly at high protein concentrations. No significant differences were observed between the two nanoparticle types with respect to Fib adsorption. However, after incubation in the mixed protein solution (SA + Tf + Fib = in vivo mimicking STF solution), NR@SAs (EM) had more abundant adsorption of each protein. This result was confirmed using SDS-PAGE (Figure S7), suggesting that NR@SAs (GTA) exhibited less nonspecific protein adsorption in multiple protein solution than NR@SAs (EM) after 1 h STF incubation. Notably, the adsorption profiles of the mixed and single protein solutions were not same, indicating that complex protein–protein interaction, particularly at higher protein concentrations (e.g. in vivo conditions), can be cooperative or repulsive. Protein adsorption on the nanoparticle surface can easily lead to changes in the subsequent protein–protein interaction. For instance, when both nanoparticle types were incubated with Tf and SA, the efficacy of SA loading was reduced (Figure 4C) because compared with free SA, Tf was more intimate with the constructed SA shells of both nanoparticles (Figure 4B). After both Tf and SA were mixed with Fib, Tf and SA interacted rapidly with the nanoparticles, effectively reducing Fib adsorption on both types of NR@SAs (EM and GTA; Figure 4E).39 Because SA and Tf adsorption on the NR@SA (EM) surface was more abundant (Figure 4C and 4D, blue line and red arrow), single protein adsorption of either SA or Tf could reduce the amount of Fib adsorption, whereas Fib adsorption on NR@SAs (GTA) was reduced only with the participation of 15



both SA and Tf. However, the adsorbed Fib could be blocked or replaced with mostly covered Tf and SA,39 allowing other free protein adsorption, possibly because Fib is known to be denatured, which can provide additional hydrophobic regions for protein adsorption on nanoparticles.45-47 Notably, additional incubation with Fib compared with Tf would have the benefit of increasing SA adsorption rather than Tf binding (Figure 4C, blue and red lines), suggesting the existence of a certain interaction between some of the adsorbed SA and Fib. This interaction could be explained by the results in Figure 3A and 3B, which revealed that the adsorbed SA was present in the denatured state and therefore had abundant hydrophobic regions that interacted with bound Fib, which was also denatured, with more abundant hydrophobic areas. After the successful adsorption of Tf, Tf binding on hydrophobic materials retains its native characteristics,38 which can prevent further protein interaction. To demonstrate the interactions between the three proteins, the informative approach of fluorescence resonance energy transfer (FRET) was used to identify the relationships between SA (TRITC), Tf (Alexa 633), and Fib (Alexa 488 or Cy 5.5) (Figures 5A and S8A). Free proteins that were equivalent to the loading amount of NR@SAs (GTA and EM) were considered as control groups. Compared with the corresponding free protein solution, more obvious energy transfer from SA (TRITC) to Tf (Alexa 633) and from Fib (Alexa 488) to SA (TRITC) was observed in the fluorescence spectra of both nanoparticle types; however, no particular energy transfer was observed from Tf (Alexa 633) to Fib (Cy 5.5) (Figures 5B and S8B). Notably, energy was transferred from SA 16


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(TRITC) to Fib (Cy 5.5). These results suggested that SA acts as the connector between Tf and Fib, which was mainly responsible for further protein adsorption. Moreover, the fluorescent ratio (650 nm/710 nm) of NR@SAs (EM) was more intense than that of the free protein, suggesting that more Tf was adsorbed onto the nanoparticle surface (Figure 4D). A higher density of Tf than that of the free protein prevented nonspecific interaction with Fib, which is consistent with the results of Fib adsorption (Figure 4E). The protein adsorption of NR@SAs (GTA and EM) in STF was explored using DLS measurements (Figure 5C and 5D). As expected, NR@SAs (EM) had a 29.1 ± 7.1 and 10.8 ± 2.2 nm increase in respective size distribution, whereas NR@SAs (GTA) had only a 10.8 ± 4.3 nm and 4.7 ± 1.6 nm increase, indicating that the low protein adsorption of NR@SAs (GTA) compared with that of NR@SAs (EM) resulted in a smaller size increase. Based on the MST analysis, protein loading magnitude, FRET data analysis, and possible protein structure analysis (Figures 4 and 5), we hypothesized the protein adsorption mechanisms for the two systems, which are illustrated in Figure 6. Tf, which was more intimate with both the constructed shells than other proteins, covered most of the nanoparticle surface, and some free SA was predominantly adsorbed in the rest of the denatured site of the constructed shells. Slow-diffusing Fib can block or substitute for mostly covered Tf and some SA. After Fib was adsorbed onto nanoparticles, the induced secondary structure changes that created more hydrophobic regions facilitated new protein adsorption, particularly SA binding. The protein adsorption reaction was assumed to achieve equilibrium once the surface was blocked with more Tf because Tf tends to 17



adsorb onto the hydrophobic parts and remains to be nondenatured.38 This assumption was supported by the results presented in Figure 4C and 4E, which revealed that additional incubation with Tf reduced the adsorption of other proteins. The more native constructed shell (GTA) efficiently reduced protein adsorption (SA + Tf), further reducing the interaction with Fib or other free proteins. By contrast, the denatured protein shell (EM), which had higher protein adsorption (SA + Tf), triggered more protein–protein interaction. The more denatured form of the adsorbed proteins SA and Fib caused severe deviations from the expected equilibrium behavior. Generally, these denatured proteins attributed to interparticle bridging to enable further protein adsorption.48 The large protein corona masked the SA shell and altered the in vitro or in vivo features of the nanoparticles.

Artificial characteristics: Crosslinking effect for high drug loading capacity of NR@SAs (GTA) To investigate the potential role of the crosslinked SA shell as a drug depot, serial concentrations of DOX solutions ranging from 0 to 512 µM were premixed with SA (3 µM), after which the aforementioned process was employed to prepare NR@DOX:SAs. The absorption band of NR@DOX:SAs was broad and had lower intensity than that of the non-DOX-loaded nanoparticles, and the hydrodynamic diameter increased as the DOX feeding concentration was increased (Figure S9A). This finding is consistent with previous research, which demonstrated that the positive charge of DOX can minimize electrostatic repulsion between SA layers and affect nanoparticle synthesis.10, 18


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49

The rapid crosslinking of SA by using GTA provided an adequate steric hindrance to maintain the

colloidal stability of NR@DOX:SAs, even at high drug loading concentrations. After nanoparticles were immersed in biological fluid (e.g. culture medium), the surface-bound DOX was grabbed by other free proteins ( 12 h), NR@SAs (GTA) exhibited similar DOX fluorescence intensity to free DOX, suggesting that long-lasting and sustained drug release can be efficiently activated by endolysosomes and that the rapid fluorescence decay of free DOX was due to its rapid interaction with the cell nucleus. To assess the interarrival variation of DOX fluorescence changes 20


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between free DOX and NR@DOX:SAs (GTA), the incubation was prolonged to 12 h and the nanoparticle concentration was halved (Figure S12B). As expected, although free DOX diffused rapidly and interacted efficiently with the cell nucleus, activity loss was observed for the remaining cells after 12 h of recovery. In addition, slow and efficient release of significantly increased DOX fluorescence from NR@DOX:SAs (GTA) was observed. The initial DOX and TRITC signals of NR@SAs (EM) were much lower than those of NR@SAs (GTA) because of inefficient cellular uptake. After the incubation period, the TRITC signal of NR@SAs (EM) decayed faster than that of NR@SAs (GTA; Figure 7C) due to more efficient surface replacement (Figure S4A) and the rapid degradation of the denatured protein shell.30 Figure 7E provides further evidence of the acid organelle-mediated protein degradation and drug release; both nanoparticle types had larger lysosomal fluorescence signal intensity than the free molecules (free DOX and SA) after 4 h of recovery; however, NR@SAs (GTA) maintained their lysosomal signal for a longer recovery time (t = 16 h), demonstrating that delayed degradation of drug release was due to the crosslinking. The dose-dependent cytotoxicity of NR@DOX:SAs (GTA) was assessed using an AlamarBlue assay. Different concentrations of DOX (64, 16, 4, 1, and 0.25 µM; corresponding to 0.6 nM Au NRs; Figure 7F) were used for the fabrication of various drug nanoagents. The half-maximal inhibitory concentration (IC50) of each type of NR@DOX:SA was determined to be 0.36, 0.57, 0.89, 1.13, and 3.56 µM, respectively. NR@DOX64:SAs had IC50 values similar to those of free DOX (0.32 µM), demonstrating that NR@DOX:SAs with a higher drug content had higher 21



antiproliferation activity toward Tramp-C1 cells. The large signal from Au NRs was internalized (Figure S13), and the DOX signals colocalized with both the endolysosomal pathway marker, transferrin-Alexa 633 (white arrow) and DAPI nuclear staining, which confirmed the long-lasting and sustained drug release of intracellular NR@DOX:SAs (Figure 7G). Furthermore, fewer NR@SAs are desirable to deliver an equivalent drug dose while causing the same level of toxicity. In addition, the photothermal properties of NR@DOX:SAs were evaluated to improve their therapeutic efficacy through combined photothermal therapy and chemotherapy (Figure S14A). The photothermal damage to cancer cells caused by NR@SA-based therapeutic nanoagents was further confirmed using live/dead double staining assays. The cells exposed to NR@DOX:SAs and NR@SAs were mainly stained red with propidium iodide (PI) after NIR irradiation, whereas most of the nontreated cells remained alive (in green, calcein-AM) under similar conditions (Figure S14B).

Hard and soft protein shells for penetrated drug delivery to tumor spheroids through NIR laser activation To further investigate the influence of the constructed SA shell on the NIR-laser-induced photothermal effect and triggered drug release against tumor cells, the induced temperature changes in both nanoparticle types were measured (Figure 8A) upon two bouts of NIR laser irradiation (808 nm, 2.65 W/cm2, 10 min). Both nanoparticle types exerted similar photothermal effects on the PBS buffer; however, the effects were different upon immersion in the lysosomal microenvironment (pH 22


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= 5). Similar temperature increases were observed in the nanoparticles, NR@SAs (GTA) and NR@SAs (EM), within a few minutes of the first NIR laser irradiation. The DLS and UV-Vis spectra (Figure S15) demonstrate that both types of nanoparticles aggregated in the acidic environment because of decreased electrical repulsion (SA, isoelectric point = ~4.5–5); however, the hard protein shell was able to preserve the optical properties to a higher degree than the soft protein shell, indicating that the hard protein shell protects the Au NR core from severe aggregation (Figure 8B). The results also reflected a variable efficacy of photothermally triggered drug release. NR@SAs (GTA) had twofold higher efficacy of photothermally triggered drug release than NR@SAs (EM; Figure 8C). Particularly, no significant drug release from NR@SAs (EM) after the second bout of NIR laser irradiation indicated that the induced aggregation resulted in the loss of the optical properties of nanoparticles that were less sensitive to NIR laser irradiation. Notably, compared with the PBS control (Figure 8B), both types of NR@SAs (GTA and EM) that did not undergo NIR laser irradiation had, after 2 h of incubation, released 10.7% and 19.3% of the drug, demonstrating that both shells were gradually biodegraded by lysosomal enzymes. The effects of differently constructed protein shells on multicellular tumor spheroids were then investigated. The highly dense tumor cells located on the peripheral edges of tumor spheroids impede the entry of drug-loaded nanoparticles. A solid structure with a diameter of approximately 250 µm was observed under a confocal microscope. The bright field image (Figure 8D) demonstrated that the abundant Au NRs from NR@SAs (GTA) surrounded the peripheral edges of tumor 23



spheroids, suggesting that the biomimetic shell exhibited high intimation with tumor cells, which is consistent with the in vitro results (Figure 2E and 2F). Notably, most DOX was localized at the tumor boundary; very few DOX molecules from NR@SAs (GTA) and NR@SAs (EM) without NIR laser irradiation were present in the central region of tumor spheroids. After 10 min of NIR laser irradiation, a considerable portion of encapsulated DOX from NR@DOX:SAs (GTA) was liberated and spread throughout the entire tumor spheroid. The fluorescent intensity of DOX was similar to that of the free-DOX-treated tumor spheroid (Figure S16). The colocalization results for the fluorescence performance in the x–z and y–z planes and the projection image (x–y plane) revealed that encapsulated DOX before NIR laser irradiation was predominantly present in the cytoplasm and some DOX molecules were possibly located in the small gaps between cells. After NIR laser irradiation, DOX from NR@SAs (GTA) was homogenously distributed in the cytoplasm and nuclei. By contrast, a small amount of DOX was released by NR@SAs (EM); however, a few DOX molecules were delivered to the central region (red arrow). This result indicated that the higher uptake of NR@SAs produced more heat, facilitating the homogenous delivery of DOX to the deeper regions of the tumor spheroids.

Cross-linked effect: NR@SAs nanoplatform (GTA) as an advanced photoacoustic amplifier Next, to verify the impact of constructed SA shell on photoacoustic (PA) effect, PA signals originated from both types of NR@SAs and Au NRs were acquired respectively using a 24


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photoacoustic microscopy system (Figure S17). Prior to the measurement, the optical density of individual suspensions was adjusted to be the same at the maximum absorption wavelength. (Figure 9A). The ns-pulse laser was set at an average fluence of 1.2 J/cm2 to avoid any spectral changes for both NR@SAs and CTAB-capped Au NRs.10, 55 The PA signal of NR@SAs displayed in Figure 9B was about 3.5-fold (10.9 dB) higher than that of CTAB-capped Au NRs in background medium (10% FBS). This result demonstrates that the constructed NR@SAs was a more powerful PA contrast agent compared to traditional Au NRs. To gain more insight into the cross-linking effect of SA shell on PA signal generation, NR@SA (EM) was obtained for further comparison. As shown in Figure 9B, the PA signal from suspension of NR@SAs (EM) of the same optical density was only 1.7-fold (4.4 dB) higher than that of CTAB-capped Au NRs. Since the albumin amount constructed for each type of NR@SAs was approximately the same (2.7 µM SA for 0.6 nM Au NRs), it is conceivable that the SA shell featured with high rigidity and cross-linked network would lead to a PA signal amplification. While protein adsorption was also an inevitable phenomenon for the measurement of PA signals in 10% FBS, the influence was monitored using 0.6 wt% fluorescence-labeled SA (SA-TRITC). The similar extent of protein adsorption found on both NR@SAs (9.3%) and CTAB-capped Au NRs (8.5%) again indicates that the compact and cross-linked SA shell plays a crucial role in PA signal amplification (Figure 9C). Compared to the albumin proteins that are loosely bound onto the surface of Au NRs through simple adsorption (Figure S18A), the appearance of the dark and thick shell 25



surrounding each gold core in negative staining TEM images (Figure 1D) confirms the dense and multilayered corona structures of NR@SAs after cross-linking. Additionally, above 80% of SA-TRITC adsorbed on both NR@SAs and CTAB-capped Au NRs could be readily removed using 1 mM glutathione; only minute protein detachment (< 5%) was detected for fluorescence-labeled NR@SAs under similar incubation conditions (Figure S18B). This result suggests that the rigid and cross-linked SA matrix plays an essential role in PA signal amplification.10, 56 It is usually considered that surface modification with materials of finite heat conductance and capacity will only broaden a heat pulse and deteriorate the PA signal. However, a low interfacial thermal resistance on nanoparticle may facilitate the transfer of the generated heat from the solid interface to the ambient environment, leading to a larger temporal temperature gradient and subsequently to the stronger amplification of the PA signal.8 For surface attachment of albumin onto Au NRs, the involvement of Au-S bonding may dominate the interfacial thermal transfer process, as covalent bonding has often been proven to be efficient in enhancing interfacial heat conductance.57-58 Moreover, as compared to CTAB-capped Au NRs, modification with silica coating8 or polymeric shell59-60 has also been reported to speed up heat dissipation in an aqueous solution. It suggests that the penetration of water into the hydrophilic shell can add a significant conduction path, leading to an effective thermal conductivity and heat capacity of the surface layer. The finding indicates that NR@SAs produce greater PA signal than NR@SAs (EM) in Figure 9B, which agrees with the aforementioned literature study, suggesting the signal amplification was attributable to the more 26


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hydrated coating layers constructed by GTA cross-linking as compared to that obtained through desolvation method. Therefore, proteins that may be in intimate contact with water were assumed to be beneficial for heat transfer (interfacial heat conductance between protein and water: 100 to 270 WK-1m-2 61; between silica and water: ~200–330 WK-1m-2 62).

NR@SA nanoplatform (GTA) for in vivo cancer theranostics Next, NR@SAs (GTA) was used to acquire in vitro PA images of Tramp-C1 cells. In contrast to non-treated cancer cells, significantly enhanced PA imaging was observed for exposed cells (Figure 9D), demonstrating that NR@SAs can serve as an efficient theranostic agent in cancer treatments. By contrast, bare Au NRs exerted high toxicity to living cells (Figure S19), suggesting its inadequacy for further PA image acquisition. The maximum non-toxic concentration of NR@SAs and Au NRs is 6 nM and 9 pM, respectively. To demonstrate the capability of NR@SAs for cancer theranostics, the PA tumor imaging in C57BL/6J mice bearing prostate tumor xenograft was acquired during, and after intratumoral injection of NR@SAs using a needle. As displayed in Figure S20, an obvious contrast enhancement was observable at the tumor injection site of mice followed by NR@SAs administration, while tumors treated with PBS showed a negligible change of PA signal (data not shown). Next, the suppression of tumor growth was observed in vivo by the combination of NR@DOX@SAs and NIR laser irradiation. The laser beam (808 nm, 2.65 W/cm2, 3 min) was targeted to the injection site and the rise in the local temperature of tumors subjected to NIR 27



irradiation was recorded using a thermal infrared camera, as displayed in Figure S21A. For mice treated with NR@SAs and NR@DOX:SAs, a rapid increase in the temperature to 90 °C appeared at the tumor site while the remaining parts showed a negligible response upon laser light exposure. In contrast, tumors treated with PBS and free DOX increased to approximately 50 °C after 3 min of irradiation. In addition, the tumor growth rate at subsequent time points was studied for 10 days. As shown in Figure S21B, photo- and chemo-combined treatments of NR@DOX:SA (+) significantly suppressed tumor growth by 80%–90% of the size of control tumors, as compared to a single treatment of NR@SA (+), NR@DOX:SA (−) and free DOX (-). Moreover, PBS (+) compared with PBS (-) showed a negectable change in tumor size, suggesting the induced non-specific photothermal effect was insufficient. The combined effect of NR@DOX:SA (+) in prostate tumor was further assessed using tail vein injections. Here, the output power of the NIR laser was reduced to 1.1 W/cm2, but irradiation occurred for 9 min. Two breaks were involved during light exposure to minimize non-specific thermal ablation (Figure S22). To ensure intratumoral drug content was sufficient for photothermal therapy, the biodistribution of NR@SAs in tumor-bearing mice at 24 h post intravenous administration has been investigated using inductively coupled plasma mass spectrometry (ICP-MS). As shown in Figure S23, NR@SAs (GTA) in comparison with its NR@SAs (EM) counterpart, presented increased gold accumulation at the target tumor site, with less sequestered in RES clearance organs such as the spleen. In agreement with the aforementioned results (Figure 2E–2G), 28


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the bioprosthetic shell of the NR@SAs (GTA) was more efficient for tumor targeted delivery, reaching an accumulation level of approximately 0.4 percentage injected dose (%ID) for Tramp-C1 tumors. This value is quite similar to the median (0.7% ID) of the administered nanoparticle dose accumulated in a solid tumor after surveying the literature from the past 10 years.63 Furthermore, the resulting temperature during irradiation was monitored using thermal images and the average temperature rise over three cycles is illustrated in Figure 10A. As expected, mice injected with NR@SAs and NR@DOX:SAs exhibited an immediate response to NIR laser irradiation, with a rise in the local temperature to 57.2 (±7.8) °C and 52.2 (±5.7) °C, respectively, after 3 min of irradiation. The temperature of all remaining groups under the same irradiation conditions was less than 38 °C. Lastly, the tumor growth curve was recorded every day and is plotted in Figure 10B. In agreement with the aforementioned results, NR@DOX:SA (+) delivering dual photo- and chemo-actions exhibited superior anti-tumor activities compared to the other groups. Free DOX at an equivalent dose of 15 mg/kg demonstrated a limited efficacy in delaying tumor growth. Furthermore, the median survival times for Tramp-C1 tumor-bearing mice receiving NR@DOX:SA (+) was 28 days versus only 12 days for DOX-treated mice (Figure 10C). Both tumor-bearing (Figure S24A) and healthy mice (Figure S24B) receiving free DOX had an average weight loss of 8.6% at 7 days post-dosing (p < 0.01); no statistically significant difference in body weight (p > 0.05) was observed for NR@DOX:SA-treated mice in comparison with the PBS-treated controls. Similar to PBS control mice, it is worthy to note that the NR@DOX:SA-injected mice also survived more than 8 months. 29



These results indicated that the safe and targeted delivery platform, NR@DOX:SA was effective in reducing adverse side effects without jeopardizing the treatment efficacy. The superior therapeutic efficacy mediated by NR@DOX:SAs subjected to NIR irradiation represents a more promising tool for future oncology medicine when compared with conventional treatment.

CONCLUSIONS On the basis of aldehyde-based crosslinking for in vitro biology, a novel biomimetic nanointimation (NR@SA, GTA) was successfully developed using a relatively high concentration of glutaraldehyde. According to our review of the relevant literature, this study was the first to employ the crosslinking and biomimetic concept for nanoparticle formulation. The half-native and half-artificial nanohybrid exhibited a few excellent characteristics for cancer theranostics. The native part of the bioprosthetic shell considerably reduced free SA adsorption, thus reducing further protein adsorption to enable less protein corona formation. Furthermore, the adsorbed protein was also more close to its native form. The native properties successfully reduced macrophage phagocytosis and increased the interaction with tumor cells for higher cellular uptake. The artificial part endowed NR@SAs with higher drug loading capacity, intact colloidal stability for stable photothermal effects and enhanced drug release, and a larger photoacoustic signal than its pristine counterpart. The delivery of anticancer drugs by using NR@SAs can effectively reduce the adverse side effects of conventional medicine. The combined photothermal therapy and enhanced chemotherapy also 30


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exhibited superior therapeutic efficacy for delaying in vivo tumor growth compared with individual treatment. Notably, the optical properties and aforementioned behaviors of the artificial part of the crosslinked NR@SAs (GTA) are comparable with those of silica-coated Au NRs.8, 64-66 Albumin proteins are natural polymers that provide distinct advantages over inorganic silica with respect to biocompatibility and biodegradability. Overall, the nanointimation NR@SAs demonstrated the interaction between biomimetic nanomaterials and biological fluids and the subsequent response to cells. Moreover, artificial crosslinking can improve the drug loading capacity and enlarge photoacoustic signal intensity of these nanoparticles for cancer theranostics. These results emphasized the importance of exploring both preserved fidelity and unavoidable artificial characterizations in biomimetic nanomaterial design.

MATERIALS AND METHODS Materials. Sodium tetrachloroaurate(III) dihydrate (99 %), glutaraldehyde (25 % wt in H2O), bovine serum albumin, transferrin, fibrinogen, hexadecyltrimethylammonium bromide (CTAB), propidium iodide (PI), dithiothreitol, phosphotungstic acid hydrate, cathepsin B, citric acid, dibasic sodium phosphate, L-cysteine, doxorubicin (DOX) and CelLytic M were obtained from Sigma–Aldrich (St. Louis, MO, USA). Methanol, ethanol, Tris⋅Cl and glycine, hydrochloric acid and nitric acid were purchased from J.T.Baker (Center Valley, PA, USA). Fetal bovine serum (catalog number: 16000044), RPMI 1640 (Roswell Park Memorial Institute), DMEM (Dulbecco's modified Eagle's

31



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medium), and phosphate buffered saline (PBS) were obtained from GIBCO (Grand Island, NY, USA).

Penicillin-Streptomycin

and

Trypsin-EDTA

Hoechst®33342 and LysotrackerTM RED DND-99,

(ethylenediaminetetraacetic

acid),

transferrin from human serum, Alexa Fluor®

633 conjugate, 4′,6-diamidino-2-phenylindole dihydrochloride (DAPI), Alexa Fluor™ 488 NHS Ester, tetramethylrhodamine-5-Isothiocyanate (TRITC) and Alexa Fluor™ 633 NHS Ester were bought from Invitrogen (Invitrogen, Carlsbad, CA, USA). Dulbecco's phosphate-buffered saline (DPBS) was brought from Biological Industries (Camarillo, CA, USA). Alamar blue® was brought from AbD Serotec (Oxford, OX5 1GE, UK). Fluorescamine was obtained from Alfa Aesar (Heysham, Lancs, USA). 5-aminofluorescein was bought from AbD Serotec (Oxford, UK). Deionized water (18.2 MΩ cm) was used to prepare all of the aqueous solutions. For the cellular experiments, all of the reagents, buffers and culture medium were sterilized by steam autoclave (134 °C, 30 min) or filtration (0.22 µm pore size, Millipore), and maintained under a sterile condition. Preparation of NR@SAs and NR@DOX:SAs. To synthesize Au NRs, a seed and growth method following the previous study was applied herein.67 The final stock solution in 7.5 nM was preserved for any further experiments. For the synthesis of NR@SAs or NR@DOX:SAs (GTA), the 0.1 mL of 20 mg/mL SA stock solution first mixed with 2 mM or 20 mM DOX solution was moderately swung for 2 hours. The stock solution was then diluted to 1.6 mL with DI water and a portion of 80 µL was added to 40 µL of the condensed Au NRs solution (7.5 nM) and mixed with 20 µL of GTA solution 32


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(25 wt%). The complex was gently swung for 16 hours. The resultant product was diluted 1 mL with DI water and subsequently washed at least 3 times by centrifugation (6000 rpm, 15 min) to remove free GTA, SA and DOX. The precipitate was re-dispersed in 1% SA solution in the first 2 times centrifugation, 0.1% SA for the third time and finally re-disperse in DI water, PBS or culture medium for further use. As for the synthesis of NR@SAs or NR@DOX:SAs (EM), the protocol was followed from the previous reports which use ethanol and methanol (1:1 v/v) to denature SA and fabricate another homogenous core-shell NR@SA (EM).26 The washing protocol was the same as the process for washing NR@SA (GTA). Characterization of NR@SAs and NR@DOX:SAs. UV-Vis spectra were obtained by UV-Vis spectrometer (Cary 100, Varian, Palo Alto, CA, USA) or plate-reader (Tecan Infinite® 200, Tecan Group AG, Basel, Switzerland) while the hydrodynamic size or zeta potential was determined by Zetasizer Nano (Malvern Instruments, England). Samples were loaded in either the disposable sizing cuvette (DTS0012) or zeta potential cuvette (DTS1070). The experiments were carried out at least triplicate. Transition electron microscopy (TEM) images were captured by (H-7100, Hitachi, Tokyo, Japan). Before analysis, the copper grid was immersed in the sample for 30 s followed by 4% phosphotungistic acid staining for 20 s and dried for 1 h. To verify DOX loading of NR@DOX:SA, the supernatant after centrifugation (5500 rpm, 15 min) was collected. The DOX absorbance at 490 nm was measured by UV-Vis spectrometer to determine the DOX encapsulation efficacy by using the following equation: %EE = [(DOX in feed –unentrapped DOX) / DOX in feed] × 100%. 33



Microscale Thermophoresis (MST). The detail of microscale thermophoresis (MST) method has been described in the previous reports.68 Briefly, proteins were followed with the manufacturer's protocol to label with fluorescent dye Cy 5.5. Fluorescence-labeled protein was purified by centrifugal filter (Amicon®, Ultra-4, 10K). The selected concentration of each unlabeled protein in the presence of Cy 5.5-labeled protein was then prepared for further experiments.69 A serial concentrations of unlabeled SA and Tf that was ranged from 23 nM–758 µM and 76 nM–625 µM) respectively both with 30 nM fluorescent labeled protein was incubated with the 2.4 nM samples (1:1 v/v) for 1 h at 37 ℃ in PBS buffer. Notably, due to low efficiency of Cy 5.5 labeling on Fib, we substitute Cy 5.5-labeled SA for Cy 5.5-labeled Fib for further experiments. Briefly, Cy 5.5 labeled-SA was pre-mixed with unlabeled SA, which the volume ratio was approximately 1:90 and 1:150 for NR@SAs (GTA) and NR@SAs (EM) fabrication based on their fluorescent intensity. After Cy 5.5-labeled NR@SAs (GTA) and NR@SAs (EM) were fabricated, a series concentration of Fib (17.85 pM–73.10 nM) was incubated with the 2.4 nM samples (1:1 v/v) for 1 h at 37 ℃ in PBS buffer. Then, the samples were loaded into silica capillaries and delivered to MST setups (NanoTemper Technologies, Germany). Measurements were also carried out by using 20% MST power and 20% IR-laser power. Data analyses were performed using Nanotemper Analysis software, v.1.5.41. SDS-PAGE. 1.2 nM NR@SAs were first incubated with STF solution (PBS buffer) or DFP (DMEM + 10 % FBS + 1% PS) at 37 ℃ for 1 and 6 h. After incubation, samples were diluted with PBS 34


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buffer (1:4 v/v) and centrifuged by 5500 g for 15 minutes and re-suspended in PBS buffer. The remained free protein after dilution and centrifugation was 250-fold reduction to reduce the effect from free SA. Notably, although the amount of remained free protein compared to original protein solution (DFP or STF) has been largely reduced, the free protein was unable to be completely removed because more centrifugation caused irreversible aggregation.37 Samples were then diluted to 6-fold reduction, DFP and free proteins from STF was diluted to 60-fold and 150-fold reduction, respectively. 20 µL of samples were incubated with 4 µL of 6× loading buffer (Ipswich, MA, USA) and heated at 95 ℃ for 5 minutes. 10 µL of samples and 5 µL protein ladder (PageRuler™ Prestained Protein Ladder) were further analyzed by 1D SDS-PAGE in an electric field under a constant voltage of either 200 V for 60 min (DFP) or 200 V for 100 min (STF) and stained with Coomassie® G-250 stain. The digital image was taken after several times cleaning for removing unstained dye from the gel. Raman Spectra. The sample preparation was similar to the protocol of the experiments for SDS-PAGE. 1.2 nM NR@SAs were first diluted with PBS buffer, centrifuged and finally suspended in PBS. After sample processing, the concentration of remained free SA was diluted to 250-fold decrease. The as prepared NR@SAs were condensed to 2.4 nM and 1 µL of the sample was dropped onto the silica wafer. Then those drops were dried by the vacuum pumper. This procedure was repeated for five times. Similarly, 10 wt% free SA served as the control was dropped onto the silica wafer and dried for one time before analysis. The Raman measurement was carried out through a 35



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Micro-Raman spectrometer (UniRam Series, ProTrus. Tech. Co., Ltd) that is equipped with 532 nm laser (ProTrus. Tech. Co., Ltd), and the laser beam aimed at the samples on microscope stage (Olympus, Center Valley, PA, USA) by the 50× Olympus slmpln objective (NA = 0.35). The laser power intensity on samples was 20 mW, and the signal dispersed by 1200 l/mm was detected by CCD (Andose 401). For each measurement, the data acquisition time was 2 seconds for 10 accumulations. As for the Raman analysis of secondary structure of adsorbed protein on nanoparticles, 1.2 nM NR@SAs was first incubated with 0.6 wt% SA at 37 ℃ for 6 h. Samples were followed the aforementioned protocol for sample processing. The dried sample was then followed the same protocol for Raman measurement of NR@SAs as described before.

.

Circular Dichroism (CD) Spectroscopy. As-prepared 1.2 nM of NR@SAs was first diluted to 0.6 nM by PBS buffer. Then nanoparticles along with corresponding free SA or CTAB-capped Au NRs was individually added to 1 mm path length quartz cuvette and performed by J-815 circular dichroism spectrometer (JASCO International Inc., Ltd. Tokyo, Japan) at 25℃. Wavelength from 200−700 nM were measured every 1 nm with scanning speed in 50 nm/min. Visible region was measured to make sure the concentration of each sample during the processing was same.37 For the protein adsorption analysis, the same samples obtained before Raman measurement were used for CD measurement. Each CD spectra was carried out by following the same protocol as described. Quantitative determination of protein adsorption on nanoparticles. Prior to determine the protein loading amount of each nanoparticles, FBS, SA, Tf and Fib followed the protocol from 36


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manufacturer was artificially modified with Cy5.5, TRITC, Alex-633 and Alex-488, respectively. A series concentration of fluorescence-labeled DFP was incubated with 1.2 nM nanoparticles for 6 h. The samples were diluted to 5-fold decrease with PBS, and the supernatant was collected and determined by fluorescence spectrometer after centrifugation (5500 rpm, 15 minutes). As for SA, Tf and Fib protein adsorption, a series of unlabeled protein was first mixed with fluorescence-labeled protein which was fixed in 0.35 µM, 0.44 µM and 17.3 nM. Each protein could be single or cooperative with other proteins, but only one kind of dye was in the protein solution in each experiment. Then, the selected protein solution was incubated with 1.2 nM nanoparticles for 1 h. Unbounded or only weak-bounded protein was obtained after centrifugation (5500 rpm, 15 minutes) and analyzed through fluorescence spectrometer. Cell culture. Tramp-C1 cells (CCL-2730, epithelial of prostate cancer cell from transgenic mouse) were obtained from American Type Culture Collection (ATCC; Manassas, VA, USA). Cells were cultured in DMEM medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin (PS). The cells were cultured at 37℃ with 5% CO2 in a humidified incubator and were passaged when reached 80% confluence. In vitro intracellular drug release and the corresponding cell viability. 1.5×104 Tramp-C1 cells were seeded in the 96-cell plate to culture 12 h. Then, cells were incubated with 2.4 nM NR@SA-TRITC (GTA and EM) and NR@DOX16:SA (GTA and EM) for 6 h. The corresponding DOX concentrations was 9.4 µM and 7.9 µM with respect to 0.6 nM NR@SA (GTA) and NR@SA 37



(EM) while free DOX was 37.6 µM. After incubation, cells were washed with DPBS twice and incubated with fresh culture medium. To visualize the performance of cell nuclei and lysosome, cells followed the protocol from manufacturer were exposed to Hoechst®33342 and LysotrackerTM RED DND-99 for 30 min after each selected recovery time. Finally, cells were either immersed in CelLytic M for protein extraction or SDS lytic buffer (50 mM Tris⋅Cl, 1 mM dithiothreitol, and 0.5 wt% SDS) for DOX and nuclei determination. The measurement was conducted by Tecan Safire platereader (Tecan Infinite 200, Tecan Group AG, Basel, Switzerland). For investigation of intracellular drug release by fluorescence microscopy or confocal microscopy, 1.5×104 Tramp-C1 cells were seeded onto 10 mm glass coverslip and set in the 48-cell plate to culture 12 h. Cells were treated with NR@DOX:SA or free DOX for 6, or 12 h and then washed with DPBS for three times before being fixed with 4 % formaldehyde. Cells were subsequently stained with 1 µM DAPI for 15 min and the fluorescence images were captured by fluorescence microscopy (IX-71, Olympus, Center Valley, PA, USA). For in vitro cell viability analysis, 5×103 Tramp-C1 cells were plated on to the 96-well plate and allowed to adhere for overnight before further use. Various DOX concentration ranged from 0.25–64 µM was picked to fabricate [email protected]:SA, NR@DOX1:SA, NR@DOX4:SA, NR@DOX16:SA and NR@DOX64:SA. The corresponding DOX concentrations were 26.6, 9.4, 4, 1, and 0.25 µM, respectively (0.6 nM Au NRs). The selected concentration of NR@SA, NR@DOX:SA, and free DOX were then incubated with cells for 24 h. Soon afterwards cells were washed with DPBS three times and re-incubated with the fresh culture medium for 48 h. 38


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To study the photo-thermal effect against Tramp-C1 cells, additional NIR light irradiation by 808 nm in a power density of 2.65 W/cm2 for 1 h (VD-ⅢA, DPSS LASER DRIVER, Unice E-O services Inc., Taiwan) was conducted prior to additional 48 h incubation. The cell viability was measured by AlamarBlue assay. In vitro intracellular uptake. 1×105 Tramp-C1 and RAW 264.7 cells were seeded onto 24-cell plate to culture 12 h. For the observation of dark-field microscopy, the same number of Tramp-C1 cells or RAW 264.7 cells were seeded onto 24-cell plate which contained one 10 mm glass coverslip in each well. To verify the mechanism of cellular uptake, 10 µM of phagocytosis inhibitor, cytochalasin D was incubated with the selected groups of cells for 1 h prior to further experiments. Then, cells were incubated with 1.2 nM NR@SA (GTA) and NR@SA (EM) for 6 h and rinsed with DPBS buffer for 3 times. Finally, cells were immersed in mixed HCl (12 M) and HNO3 (70%) solution (1:1 v/v) for 12 h before inductively coupled plasma mass spectrometry (ICP-MS, Agilent 7700 Series ICP-MS, USA) analysis for determining intracellular gold content. Meanwhile, dark-field images were obtained through dark-field microscopy (IX-71, Olympus, Center Valley, PA, USA) NIR-Activation drug release. 2.4 nM NR@DOX32:SA (GTA) and NR@DOX32:SA (EM) was either immersed in PBS or lysosome mimicking buffer (LB buffer, 200 mM citric acid with 65.5 mL of 200 mM dibasic sodium phosphate and 2.2 g L-cysteine) with 1.2 U/ml enzyme cathepsin B. Each samples were irradiated with 808 nm of NIR laser (2.65 W/cm2) for 10 minutes. Meanwhile, the temperature was recorded by a thermal couple. Then, the samples were centrifuged (5500 g, 15 39



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minutes) and re-suspended in the same buffer. The experimental cycle was conducted for two times. For evaluation of NIR-activated drug release, two cycles of laser on/off was conducted through a 808 nm NIR laser in a power density of 2.65 W/cm2 (20 min for laser on, 40 min for laser off). Supernatant was collected from each selected point by centrifugation (5500 g, 15 minutes) and DOX fluorescence was determined by fluorescence spectrometer. NIR-Activation

of

Nanoparticle

Activity

in

Multicellular

Tumor

Spheroids.

Multicellular-constructed tumor spheroids were followed from the protocol of hanging drop methods in pervious study.70 Cell density was optimized to 1×105 Tramp-C1 cells/ml on culture dish for 7 days. The fresh culture medium would be additionally supplied every 2-3 days. The resultant tumor spheroids were washed with PBS buffer twice and then transferred to 24-well plate which contained one coverslip (10 mm diameter) on each wall. Then, tumor spheroids were incubated with 2.4 nM NR@DOX16:SA (GTA), NR@DOX16:SA (EM) and corresponding free DOX (0.6 nM Au NRs versus 9.4 µM DOX) for 6 h. After incubation, tumor spheroids were washed with PBS buffer twice and exposed to 808 nm of NIR laser irradiation (2.65 W/cm2) for 10 minutes. They were then stained with 5 µM calcein-AM for 30 minutes and fixed by 4% paraformaldehyde. Finally, they were also stained with 5 µM DAPI for 15 minutes prior to confocal microscopic observation. Photoacoustic microscopy system. Dark field confocal photoacoustic microscopy system was built up by Prof. Li et al. and the whole setting in detail was illustrated in the previous reports.71 Briefly, a Nd:YAG Q-switched pumped tunable laser (Surlite II-10, Continuum,USA) was utilized to provide 40


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laser pulses at 808 nm with 10-Hz pulse-repetition frequency (PRF) and 6.5-ns pulse width. The laser light from the multi-mode fiber was aligned to be confocal with a 25 MHz ultrasonic transducer (v324, Olympus, USA) at the target site through a convex lens, an axicon, a plexiglass mirror. 25-MHz focused ultrasonic transducer (−6 dB fractional bandwidth: 55%, focal length: 13 mm, v324, Olympus) provided 68 µm and 171 µm of axial and lateral resolution, respectively. The transducer was immersed in the water tank and a hole at the bottom of the water tank served as the available area for detection. The subject (transparent conduit, cell culture dish or tumor bearing mouse) was positioned at the target site at the bottom of the water tank. The set of piezoelectric motors (HR8 Ultrasonic Motor, Nanomotion) controlled the location of the single transducer or was used to perform 2-D raster scanning. Detected signals received from the ultrasonic transducer were pre-amplified by a low-noise amplifier (AU-3A-0110, Miteq, USA). The signals were then cascaded to an ultrasonic pulser/receiver (5073 PR, Olympus, USA) and low-pass filtered. Then, they were further digitized by a 14-bit digital (A/D) card (CompuScope 14200, GaGe, USA), processed at 200 MHz, and finally stored in the PC. 16-time signal average was processed for each A-line signal to gain higher signal-to-noise ratio. All the experiments were independently carried out for four times. For in vivo intratumoral photoacoustic images, 3-D C-scan images at tumor foci before and 24 h-post injection were reconstructed, then projected to x-y and y-z planes and quantitatively analyzed through MATLAB software (R2016a).

41



In vivo photo-and chemo-therapy by intravenous injection. All the 6-8 weeks male mice (C57BL/6J) purchased from National Laboratory Animal Center and were handled according to the guideline from Laboratory Animal Center of National Tsing Hua University, Taiwan. 2×106 Tramp-C1 cells were subcutaneously injected into the left limp of the mice. When the tumor size reached to approximately 150 mm3, all the tumor bearing mice were randomly allocated into several groups and separately received 100 µL of PBS, DOX15, NR@SA, or NR@DOX15:SA (Au: 3.36 mg/kg, DOX: 15 mg/kg) through tail vain injection. Treatment of NIR light irradiation (808 nm, 1.1 W/cm2, 3 min) were given three times at 2, 12 and 24 post-injection. The temperature change was monitored by IR camera (Thermo Shot F30, NFC Avio Infrared Technologies Co., Ltd) during the NIR light irradiation. Tumor size was measured everyday through a caliper and the tumor volume was determined by V = 1/2 × (W)2 × (L) where W and L represented the width and length of tumor. Statistical Analysis. Experimental data was quantitatively presented as the mean ± standard deviation. Statistical significance by a two tailed student ′s test (P < 0.05) was determined unless otherwise stated.

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B

A

C

D

Figure 1. Characterization of the synthesis of NR@SAs. (A) UV-Vis spectra, (B) hydrodynamic size and (C) zeta potential of Au NRs, NR@SAs (GTA) and NR@SAs (EM) in phosphate buffered saline (PBS). Pure Au NRs was immersed in DI water. Before DLS measurements, particles were diluted 20-fold with DI water and then added to the zeta potential cuvette. Hydrodynamic size and zeta potential measurements were carried out together under the same conditions. (D) The TEM images of (a) Au NRs, (b–c) NR@SA (GTA) and (d–e) NR@SA (EM). Scale bar: pictures of (a), (b) and (d) are 50 µm, and pictures of (c) and (e) are 200 µm.

43



A

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C

B

D

E

F

G

Figure 2. Analysis of protein adsorption and its resultant influence on cellular uptake. (A) SDS-PAGE of the DFP, free culture medium (Dulbecco’s Modified Eagle Medium [DMEM] + 10% FBS + 1% PS), and nanoparticles, 1.2 nM NR@SAs (GTA) and NR@SAs (EM) in DFP. The concentration of NR@SAs (DFP) and free culture medium (DFP) were diluted 6- and 60-fold before SDS-PAGE was performed. (B) 1 ml of 1.2 nM samples were prepared and added to the disposable size cuvette for further measurement. DLS spectra of NR@SAs (GTA) and NR@SAs (EM) in PBS

44


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buffer or free culture medium (DFP) were then obtained. Labels 1st and 2nd indicated the two distributions of DLS profile from NR@SAs (GTA) and NR@SAs (EM), respectively. (C) An increment in the particle size of NR@SAs occurred during incubation in DFP. n = 3, ***p < 0.001. (D) Schematic of NR@SAs (GTA) and NR@SAs (EM) after FBS adsorption. FBS was more prone to adsorption on NR@SAs (EM) than on NR@SAs (GTA). The composition of the protein shell and adsorbed protein influences cellular uptake. (E) Dark-field images and ICP-MS results of the uptake of NR@SAs (GTA) and NR@SAs (EM) by (F) macrophages (RAW 264.7) and (G) cancer cells (Tramp-C1) which were either pre-treated with cytochalasin D or not. n = 3–4, *p < 0.05, **p < 0.01, and n.s. > 0.05.

45



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A

B

C

Figure 3. Protein structure analysis. (A) Raman spectra and (B) CD spectra of native SA, NR@SAs (GTA), NR@SAs (EM), and both nanoparticle types after further incubation with 0.6 wt% SA. The nanoparticles prepared after incubation with 0.6 wt% SA for 6 h were centrifuged to remove free proteins and suspended in PBS buffer before further experiments. To analyze the Raman spectra, the spectra within 1200–1700 nm were fitted using Origin 8.0 software, which could effectively reconstruct the original peaks. Amide Ⅰ region (beige area): 1640–1700 cm-1; amide Ⅲ (pale blue area): 1220–1300 cm-1.33, 72 (C) Schematic of the interaction between the differently constructed

46


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shells and the proteins in biological fluids: (a) native and (b) denatured SA of the constructed SA shell; adsorbed (c) native and (d) denatured SA from 0.6 wt% SA nanoparticles.

A

B

C

D

E

Figure 4. Protein binding analysis: single protein or multiple protein interactions with NR@SA (GTA) and NR@SA (EM). MST analysis of single protein adsorption. (A) MST results of 47



single-protein binding of 1.2 nM NR@SA after thermophoretic movement of Cy5.5 labeled protein changes under thermal gradient, which is expressed as the normalized fluorescence (Fnorm) and defined as Fhot/Fcold. (B) Bind affinity KD (Koff/Kon) was determined through microscale thermophoresis (MST). KD was expressed as 10-6 M. The loading amount of protein adsorption of NR@SAs by single or multiple protein mixing simultaneously. NR@SA (GTA, up panel) and NR@SA (EM, down panel) were co-incubated with a series concentration of (C) SA, (D) Tf and (E) Fib that was ranged from 3.0-151.5 µM, 1-75.0 µM and 0.3-13.2 µM, respectively. The incubation of SA, Tf and Fib could be solely (expressed as none) or additionally with fixed concentration of SA (151.5 µM), Tf (37.5 µM) and Fib (13.2 µM). The red arrow indicates in vivo conditions. (SA: 151.5 µM, Tf: 37.5 µM, and Fib: 5 µM or 13.2 µM). n = 3–4, *p < 0.05, **p < 0.01 or n.s. > 0.05. #p < 0.05, ##p < 0.01 versus NR@SA (GTA) control.

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A

B

C

D

Figure 5. Monitoring the mechanism of protein adsorption by fluorescence resonance energy transfer (FRET). (A) Prior to further experiments, 2% (0.26 µM), 10% (15.2 µM) and 1.67 % (0.63 µM) of Fib, SA and Tf in STF solution was individually labeled with Alexa-488, TRITC and Alexa-633

prior

to

the

incubation.

1.2

nM

of

nanoparticles

49


were

incubated

with


fluorescence-contained STF solution for 1 h, and diluted to 0.24 nM. Then, nanoparticles were washed with PBS buffer once by centrifugation and condensed to 0.6 nM. Each fluorescence spectra was obtained by excitation (Ex) at 488 nm, 550 nm, 633 nm, 685 nm through fluorescence spectroscopy. (B) To see whether the energy transfer was occurred in the surface of nanoparticles between bounded proteins, the equivalent loading amount of free protein solution was served as the negative control for comparison. The change of energy transfer was presented as fluorescent ratio of emission to clearly quantify the different energy transfer between the bounded protein and free form. n = 3, *p < 0.05 or **p < 0.01. (C) DLS spectra of 1.2 nM NR@SA after 1 h incubation with STF mixed protein solution (151.5 µM SA+ 37.5 µM Tf and 13.2 µM). Labels 1st and 2nd indicated the two distributions of DLS profile of NR@SAs (GTA) and NR@SAs (EM), respectively. (D) An increment in the particle size of NR@SAs occurred during incubation in STF.

50


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A

B

C

D

Figure 6. Schematic of the interaction between differently constructed SA shells and biofluids. (A) SA and Tf interacted rapidly with the protein-constructed surface. SA was more likely to be adsorbed 51



in its denatured form, and some SA on the GTA shell was preserved in its native form. According to a previous study, Tf is mostly adsorbed on the hydrophobic site in its native form.38 Overall, NR@SA (GTA) exhibited less protein adsorption. (B) The native Tf and portion of SA that provided hydrophilic characterizations can prevent Fib adsorption. However, some of the bound Tf and SA were replaced with Fib.39 (C) After Fib encountered nanoparticles, it denatured and subsequently aggregated, acting as the interprotein bridge for further protein adsorption, particularly SA adsorption. The induced hydrophobic interaction denatured these adsorbed SA proteins and induced more protein adsorption. (D) Until more Tf was bound on the outer surface layer, the protein adsorption achieved equilibrium. NR@SAs (GTA) that are closer to their native form can efficiently reduce their exposure and interaction with free proteins, leading to a smaller protein corona formation.

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B

A

C

D

F

E

G

Figure 7. Drug loading analysis represented by (A) encapsulation efficacy (EE%) and (B) corresponding loading amount of DOX in 0.6 nM NR@SAs (GTA). Crosslinking effect for 53



intracellular drug transport. Cells were incubated with 1.2 nM (C) TRITC-labeled NR@SAs, (D) NR@DOX:SAs, or the corresponding free molecules for 6 h and then recovered at selected time intervals. Each fluorescence spectrum was obtained after the staining of hoechst or (E) lysotracker and cell lysis. The fluorescence signal was normalized by dividing by the intensity of hoechst fluorescence for further comparison. n = 4, *p < 0.05 and **p < 0.01. #p < 0.05 and ##p < 0.01 versus NR@SA (EM). (F) Dose-dependent cytotoxic effect of NR@DOX:SAs (GTA) and DOX on cells. Cells were allowed to recover for an additional 48 h in a fresh culture medium prior to the AlamarBlue assay. (G) The intracellular transport of NR@DOX:SAs (GTA) and its uptake by Tramp-C1 cancer cells after 24 h was visualized using laser-scanning confocal fluorescence microscopy. The nuclei were stained blue with DAPI, and the acidic endolysosomal compartments were stained green with commercial transferrin-Alexa 633. White arrows indicate the overlapping of the DOX and Tf fluorescence signals. Scale bar: 20 µm. The Au NRs of the samples were maintained at 0.6 nM with respect to approximately 9.4 µM DOX.

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A

D

55



B

C

Figure 8. Hard and soft corona for the photothermal effect. (A) Time-dependent temperature elevation proﬁles of 2.4 nM NR@SAs (GTA), NR@SAs (EM), or control (PBS or LB) suspended in PBS or the lysosomal-mimicking buffer (LB) under 808 nm of 2.65 W/cm2 NIR laser irradiation. n = 3. (B) Schematic of NR@SAs (GTA) and NR@SAs (EM) in LB buffer after NIR laser irradiation. 56


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(C) NIR-laser-activated drug release from 2.4 nM NR@DOX:SAs (GTA) or NR@DOX:SAs (EM) in PBS or LB buffer after two bouts of 2.65 W/cm2 NIR laser irradiation (20 min per bout). (D) In

vitro drug penetration of 2.4 nM NR@DOX:SAs (GTA), NR@DOX:SAs (EM), or free DOX in 3D tumor spheroids was observed using confocal microscopy. The observation was conducted along the

x, y, and z directions. DAPI (blue) represents the cell nuclei, and calcein-AM (green) refers to the cytoplasm. Red arrows indicate the location of released DOX. The Au NRs of the samples were maintained at 0.6 nM with respect to approximately 9.4 µM DOX.

A

B 57



D

C

Figure 9. NR@SA nanoplatform as a new photoacoustic amplifier. (A) UV-Vis spectra of fresh samples: 0.6 nM Au NRs, NR@SAs (GTA) and NR@SAs (EM) in background culture meidum (10% FBS). (B) The PA signal generated from the fresh Au NRs, and NR@SAs was detected using a transducer and was collected using an ultrasonic pulser/receiver. 16-time signal average was processed for each A-line signal. n=4–5, **p < 0.01 and ***p < 0.001. (C) Schematic drawing demonstrating the physical adsorption of SA onto the surface of Au NRs and NR@SAs. (D) 2 mm × 2 mm area of projected C-scan PA images of non-treated and NR@SAs treated tumor cells.

A

58


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C

B

Figure 10. In vivo anti-tumor activity of various therapeutic agents towards tumor growth delay. (A) Thermal imaging of different tumor-bearing mice during photothermal treatment (808 nm, 1.1 W/cm2, irradiated 3 times for 3 min each time) at day 1 post intravenous injection. (B) Tumor growth curve (n=3–5) and (C) survival curve of mice after receiving various treatments (Au: 3.36 mg/kg, DOX: 15 mg/kg) through tail vein administration (one dose on day 0). n = 5–8, *p < 0.05, **p < 0.01 or ***p < 0.001 versus PBS (-) control. #p < 0.05 or ###p < 0.001.

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ASSOCIATED CONTENT

Supporting Information The Supporting Information is available free of charge on the ACS Publications website. Figures were provided for a serial comparison between NR@SAs (GTA) and NR@SAs (EM) associated with the physicochemical properties (e.g., hydrodynamic size, UV-Vis absorption, colloidal stability, surface functional groups, FBS or STF adsorption, and payload release), as well as their interactions with macrophages, cancer cells, or tumor spheroids, and in vivo biodistribution. Theranostic effects of NR@SAs have also been evaluated both in vitro and in vivo by cell viability test, photoacoustic images, and tumor growth delay.

AUTHOR INFORMATION

Corresponding Author

Prof. Yu-Fen Huang, Department of Biomedical Engineering and Environmental Sciences, National Tsing Hua University, Hsinchu 30013, Taiwan, ROC. Email: [email protected] Notes The authors declare no competing financial interest.

ACKNOWLEDGMENT We appreciate financial support from the Ministry of Science and Technology (NSC 102-2113-M-007-005-MY3, 105-2113-M-007 -021, 105-2627-M-019 -001 -, 106-2113-M-007 -008 -, 106-2627-M-019 -001 -) of Taiwan, ROC.

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Serum Albumin

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