Serum Albumin

Article pubs.acs.org/cm

Cite This: Chem. Mater. 2018, 30, 729−747

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. S Supporting Information *

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

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Au NRs modified with silica8 or graphene oxide9 as core−shell nanoparticles exhibit an enhancement in photoacoustic signals greater than that of their pristine counterparts, leading to a better imaging capability for in vivo studies. Physical adsorption and desolvation/cross-linking are two common strategies used to successfully combine Au NRs with serum albumin (SA). Through the use of 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 cross-linker, glutaraldehyde (GTA), is required in desolvation methods to stabilize nanoparticles through the cross-linking of resoluble SA.15 The soft corona influences the resultant particle size, stability, and drug release because of the 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

INTRODUCTION Albumin and gold nanorods (Au NRs) have emerged as a versatile carrier for the systematic delivery of therapeutic and diagnostic agents. The former 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 to 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, 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 antitumor 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 a rapid thermal expansion and contraction and the generation of pressure transients. The outgoing thermoacoustic wave can be detected with an ultrasound transducer and then used to reconstruct photoacoustic (PA) images. A few studies have further reported that © 2017 American Chemical Society

Received: September 28, 2017 Revised: December 13, 2017 Published: December 13, 2017 729

DOI: 10.1021/acs.chemmater.7b04127 Chem. Mater. 2018, 30, 729−747

Article

Chemistry of Materials

Figure 1. Characterization of the synthesis of NR@SAs. (A) UV−vis spectra, (B) hydrodynamic size, and (C) z-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 z-potential cuvette. Hydrodynamic size and z-potential measurements were carried out together under the same conditions. (D) TEM images of (a) Au NRs, (b and c) NR@SA (GTA), and (d and e) NR@SA (EM). Scale bar: pictures of a, b, and d are 50 μm, and pictures of c and e are 200 μm.

bioprosthetic shell 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 a simple one-step fabrication of the bioprosthetic SA shell to provide another perspective of biomimetic nanomaterial design. Furthermore, although the artificial cross-linking 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 successfully in vivo tumor growth inhibition through combined photothermal therapy and 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.

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 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 the 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 allow these materials to be worthy of further investigation. This study developed a facile and controllable synthesis method for the successful fabrication of 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 GTA to protein to directly immobilize albumin and form a 730


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Figure 2. Analysis of protein adsorption and its resultant influence on cellular uptake. (A) SDS-PAGE of the free culture medium (DFP, 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) One milliliter of 1.2 nM samples was prepared and added to the disposable size cuvette for further measurement. DLS spectra of NR@SAs (GTA) and NR@SAs (EM) in PBS or 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 pretreated with cytochalasin D or not. n = 3−4, *p < 0.05, **p < 0.01, and ns > 0.05.

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RESULTS AND DISCUSSION Cross-Linking Effect: One-step Synthesis and Characterizations of NR@SA Nanoplatform. In the beginning, to construct a homogeneous 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 = 0−33 300). 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 731


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whereas NR@SAs (GTA) had only a 22.6 ± 7.7 and −1.1 ± 2.1 nm increase (Figure 2C). 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) (Figures 2E and F). 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 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 cross-linking 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 (Figures 2E and G), 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, the 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

concentration of the dispersions was adjusted from 40 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 decreasing CTAB concentration with low GTA concentration leads to a reduced level of protein adsorption. Even a high concentration of GTA was beneficial 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 concentrations of 0.1 M and 40 μM, respectively, 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 the 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 z-potential of CTAB-coated NRs and NR@SAs was +21.5 (±0.8) 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 nanoparticle that was introduced from the previous study using desolvation methods was also fabricated.26 The similar protein loading amount, size, surface charge, and morphology of nanoparticles made it a good candidate to serve as a negative control for the following experiments (Figure 1). 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’s 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, 732


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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 before further experiments. To analyze the Raman spectra, the spectra within 1200−1700 nm were fitted using Origin 8.0 software, which effectively reconstructed 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 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.

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 drying 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 as 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 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

internalization, 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, without wash). Unavoidable protein adsorption on both nanoparticles during purification could balance the loss of functional groups during fabrication (Figure S6, with 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 733


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Figure 4. Protein binding analysis: single protein or multiple protein interactions with NR@SA (GTA) and NR@SA (EM). (A) MST results of 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) Binding affinity KD (Koff/Kon) was determined through MST. The adsorption of protein on NR@SAs was done by single or multiple protein mixing simultaneously. NR@SA (GTA, upper panel) and NR@SA (EM, lower panel) were coincubated with a series concentration of (C) SA, (D) Tf, and (E) Fib that ranged from 3.0−151.5, 1−75.0, and 0.3−13.2 μM, respectively. The incubation of SA, Tf, and Fib was done alone (expressed as none) or in addition to a 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 ns > 0.05. #p < 0.05, ##p < 0.01 versus NR@SA (GTA) control.

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) exhibited only 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. 734


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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. Nanoparticles (1.2 nM) were incubated with fluorescence-contained STF solution for 1 h and diluted to 0.24 nM. Then, nanoparticles were washed with PBS once by centrifugation and condensed to 0.6 nM. Each fluorescence spectrum was obtained by excitation (Ex) at 488, 550, 633, and 685 nm through fluorescence spectroscopy. (B) To see whether the energy transfer occurred in the surface of nanoparticles between bounded proteins, the equivalent loading amount of free protein solution served as the negative control for comparison. The change of energy transfer was presented as a 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 of 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 increase in the particle size of NR@SAs occurred during incubation in STF.

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

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. Not only did the constructed shell (GTA) still retain its amide III structure (1235 and 1270 cm−1) and partial amide I structure (941 cm−1) 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 remain amide III structure at approximately 1674 cm−1, indicating that the adsorbed SA was 735


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

Bioprosthetic Albumin Shell Reduced Protein Corona Formation by Blood Proteins. Three blood proteins (bovine SA, Tf, and 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

(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). 736


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(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 Figures 3A and B, 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 (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 (Figures 5C and D). 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. On the basis of 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 adsorb onto the hydrophobic parts and remains to be nondenatured.38 This assumption was supported by the results presented in Figures 4C and E, 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

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 4A and B). 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 solutions were incubated with NR@SAs (Figures 4C−E), and the loading amount was then quantified. For the single protein adsorption profile (Figures 4C−E, black line), NR@SAs (EM) exhibited a larger amount of SA and Tf adsorption 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 (Figures 4C and D, 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 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 737


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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). Cross-linking effect for 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 versus cell only. #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) 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.

particle synthesis.10,49 The rapid cross-linking 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 DOX fluorescence intensity similar to that of 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 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, a 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 a 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 cross-linking. 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 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 longlasting 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 740


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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 medium (10% FBS). (B) The PA signal generated from the fresh Au NRs, and NR@SAs were detected using a transducer and collected using an ultrasonic pulser/receiver. Sixteen-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 × 2 mm area of projected C-scan PA images of nontreated and NR@SA treated tumor cells.

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 (SATRITC). 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 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 ( 0.05) was observed for NR@DOX:SAtreated 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 eight months. 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.

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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, United States). Methanol, ethanol, Tris·Cl and glycine, hydrochloric acid, and nitric acid were purchased from J.T.Baker (Center Valley, PA, United States). Fetal bovine serum (catalog number: 16000044), RPMI 1640 (Roswell Park Memorial Institute), DMEM (Dulbecco’s modified Eagle’s medium), and phosphate buffered saline (PBS) were obtained from GIBCO (Grand Island, NY, United States). Penicillin−streptomycin and Trypsin-EDTA (ethylenediaminetetraacetic acid), Hoechst33342 and LysotrackerTM RED DND-99, transferrin from human serum, Alexa Fluor 633 conjugate, 4′,6-diamidino-2-phenylindole dihydrochloride (DAPI), Alexa Fluor 488 NHS Ester, tetramethylrhodamine-5isothiocyanate (TRITC), and Alexa Fluor 633 NHS Ester were bought from Invitrogen (Invitrogen, Carlsbad, CA, United States). Dulbecco’s phosphate-buffered saline (DPBS) was bought from Biological Industries (Camarillo, CA, United States). Alamar blue was bought from AbD Serotec (Oxford, OX5 1GE, UK). Fluorescamine was obtained from Alfa Aesar (Heysham, Lancs, United States). 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), 0.1 mL of 20 mg/mL SA stock solution first mixed with 2 or 20 mM DOX solution was moderately swung for 2 h. 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 (25 wt %). The complex was gently swung for 16 h. The resultant product was diluted with 1 mL of 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 redispersed in 1% SA solution in the first 2 round of centrifugation, 0.1% SA for the third round, and finally redispersed 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 used ethanol and methanol (1:1 v/v) to denature SA and fabricate another homogeneous 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 a UV−vis spectrometer (Cary 100, Varian, Palo Alto, CA, United States) or plate-reader (Tecan Infinite 200, Tecan Group AG, Basel, Switzerland) while the hydrodynamic size or z-potential was determined by Zetasizer Nano (Malvern Instruments,

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CONCLUSIONS On the basis of aldehyde-based cross-linking for in vitro biology, a novel biomimetic nanoimitation (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 cross-linking and biomimetic concept for nanoparticle formulation. The halfnative 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 photoacoustic signal larger than that of 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 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 cross743


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Chemistry of Materials England). Samples were loaded in either the disposable sizing cuvette (DTS0012) or z-potential cuvette (DTS1070). The experiments were carried out at least in triplicate. Transition electron microscopy (TEM) images were captured by using an H-7100 instrument (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 drying 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%. Microscale Thermophoresis (MST). The details of the MST method were described in the previous reports.68 Briefly, the manufacturer’s protocol was followed to label proteins with fluorescent dye Cy 5.5. Fluorescence-labeled protein was purified by a 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 Serial concentrations of unlabeled SA and Tf that ranged from 23 nM to 758 μM and 76 nM to 625 μM, respectively, both with 30 nM fluorescent labeled protein were incubated with the 2.4 nM samples (1:1 v/v) for 1 h at 37 °C in PBS. Notably, due to the low efficiency of Cy 5.5 labeling on Fib, we substituted Cy 5.5-labeled SA for Cy 5.5-labeled Fib for further experiments. Briefly, Cy 5.5 labeled-SA was premixed 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 to 73.10 nM) was incubated with the 2.4 nM samples (1:1 v/v) for 1 h at 37 °C in PBS. 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. NR@SAs (1.2 nM) were first incubated with STF solution (PBS) or DFP (DMEM + 10% FBS + 1% PS) at 37 °C for 1 and 6 h. After incubation, samples were diluted with PBS (1:4 v/v) and centrifuged at 5500g for 15 min and resuspended in PBS. The remained free protein after dilution and centrifugation was 250-fold reduction to reduce the effect from free SA. Notably, although the amount of remaining free protein compared to original protein solution (DFP or STF) was largely reduced, the free protein was unable to be completely removed because more centrifugation caused irreversible aggregation.37 Samples were then diluted to a 6-fold reduction, and DFP and free proteins from STF were diluted to 60fold and 150-fold reduction, respectively. Twenty microliters of samples were incubated with 4 μL of 6× loading buffer (Ipswich, MA, United States) and heated at 95 °C for 5 min. Ten microliters of samples and 5 μL of 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 cleaning several times to remove unstained dye from the gel. Raman Spectra. The sample preparation was similar to the protocol of the experiments for SDS-PAGE. NR@SAs (1.2 nM) were first diluted with PBS, 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 five times. Similarly, 10 wt % free SA served as the control was dropped onto the silica wafer and dried before analysis. The Raman measurement was carried out through a micro-Raman spectrometer (UniRam Series, ProTrus. Tech. Co., Ltd.) that was equipped with 532 nm laser (ProTrus. Tech. Co., Ltd.), and the laser beam was aimed at the samples on microscope stage (Olympus, Center Valley) 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 s 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 °C for 6 h. The aforementioned protocol was followed for sample processing. Raman measurement of NR@SAs in the dried samples followed the same procedure as described before. Circular Dichroism (CD) Spectroscopy. As-prepared 1.2 nM NR@SAs were first diluted to 0.6 nM by PBS. Then, nanoparticles along with corresponding free SA or CTAB-capped Au NRs was individually added to 1 mm path length quartz cuvette and analyzed using a J-815 circular dichroism spectrometer (JASCO International Inc., Ltd. Tokyo, Japan) at 25 °C. Wavelength from 200−700 nM was measured every 1 nm with the scanning speed of 50 nm/min. Visible region was measured to make sure the concentration of each sample during the processing was the same.37 For the protein adsorption analysis, the same samples obtained before Raman measurement were used for CD measurement. CD spectra were obtained by following the same protocol as previously described. Quantitative Determination of Protein Adsorption on Nanoparticles. Prior to the determination of the protein loading amount of each nanoparticle, FBS, SA, Tf, and Fib, following the protocol from the manufacturer, were 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 spectrometry after centrifugation (5500 rpm, 15 min). For SA, Tf, and Fib protein adsorption, a series of unlabeled protein was first mixed with fluorescence-labeled protein which was fixed at 0.35, 0.44, 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 min) and analysis using a 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, United States). Cells were cultured in DMEM medium supplemented with 10% fetal bovine serum (FBS) and 1% penicillin−streptomycin (PS). The cells were cultured at 37 °C with 5% CO2 in a humidified incubator and were passaged when 80% confluence was reached. In Vitro Intracellular Drug Release and Corresponding Cell Viability. Tramp-C1 cells (1.5 × 104) were seeded in the 96-cell plate to culture for 12 h. Then, cells were incubated with 2.4 nM NR@SATRITC (GTA and EM) and NR@DOX16:SA (GTA and EM) for 6 h. The corresponding DOX concentrations were 9.4 and 7.9 μM with respect to 0.6 nM NR@SA (GTA) and NR@SA (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, following the protocol from the manufacturer, we exposed cells to Hoechst33342 and Lysotracker 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 using a Tecan Safire plate reader (Tecan Infinite 200, Tecan Group AG, Basel, Switzerland). For investigation of intracellular drug release by fluorescence microscopy or confocal microscopy, 1.5 × 104 TrampC1 cells were seeded onto a 10 mm glass coverslip and set in the 48cell 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 3 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). For in vitro cell viability analysis, 5 × 103 Tramp-C1 cells were plated on to the 96well plate and allowed to adhere overnight before further use. Various DOX concentrations ranging from 0.25−64 μM were picked to 744


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Chemistry of Materials 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 concentrations of NR@SA, NR@DOX:SA, and free DOX were then incubated with cells for 24 h. Soon afterward, cells were washed with DPBS 3 times and reincubated with the fresh culture medium for 48 h. To study the photothermal 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 EO services Inc., Taiwan) was conducted prior to additional 48 h incubation. The cell viability was measured using an AlamarBlue assay. In Vitro Intracellular Uptake. Tramp-C1 and RAW 264.7 cells (1 × 105) were seeded onto 24-cell plates to culture for 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 plates which contained 1 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 3 times. Finally, cells were immersed in mixed HCl (12 M) and HNO3 (70%) solution (1:1 v/v) for 12 h before ICP-MS (Agilent 7700 Series ICP-MS, United States) analysis for determining intracellular gold content. Meanwhile, dark-field images were obtained through darkfield microscopy (IX-71, Olympus, Center Valley). NIR-Activation Drug Release. NR@DOX32:SA (GTA) and NR@DOX32:SA (EM) (2.4 nM) were immersed in either 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 sample was irradiated with 808 nm of NIR laser (2.65 W/cm2) for 10 min. Meanwhile, the temperature was recorded by a thermal couple. Then, the samples were centrifuged (5500g, 15 min) and resuspended in the same buffer. The experimental cycle was conducted twice. For evaluation of NIRactivated drug release, two cycles of laser on/off were 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). The supernatant was collected from each selected point by centrifugation (5500g, 15 min) and DOX fluorescence was determined using a fluorescence spectrometer. NIR-Activation of Nanoparticle Activity in Multicellular Tumor Spheroids. The hanging drop method found in a previous study70 was used for multicellular-constructed tumor spheroids. Cell density was optimized to 1 × 105 Tramp-C1 cells/ml on a culture dish for 7 days. Fresh culture medium was additionally supplied every 2−3 days. The resultant tumor spheroids were washed with PBS 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 twice and exposed to 808 nm of NIR laser irradiation (2.65 W/cm2) for 10 min. They were then stained with 5 μM calcein-AM for 30 min and fixed by 4% paraformaldehyde. Finally, they were also stained with 5 μM DAPI for 15 min prior to confocal microscopic observation. Photoacoustic Microscopy System. Dark field confocal photoacoustic microscopy system was built 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, United States) was utilized to provide laser pulses at 808 nm with 10Hz pulse-repetition frequency (PRF) and 6.5 ns pulse width. The laser light from the multimode fiber was aligned to be confocal with a 25 MHz ultrasonic transducer (v324, Olympus, United States) at the target site through a convex lens, an axicon, a plexiglass mirror. Focused ultrasonic transducer (25-MHz, −6 dB fractional bandwidth: 55%, focal length: 13 mm, v324, Olympus) provided 68 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 2D raster scanning. Detected signals received from the ultrasonic transducer were preamplified by a low-noise amplifier (AU-3A-0110, Miteq, United States). The signals were then cascaded to an ultrasonic pulser/receiver (5073 PR, Olympus, United States) and low-pass filtered. Then, they were further digitized by a 14-bit digital (A/D) card (CompuScope 14200, GaGe, United States), processed at 200 MHz, and finally stored in the PC. Sixteen-time signal average was processed for each A-line signal to gain higher signal-to-noise ratio. All of the experiments were independently carried out four times. For in vivo intratumoral photoacoustic images, 3D 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). In Vivo Photo- and Chemotherapy by Intravenous Injection. Mice, all 6−8 weeks old and male (C57BL/6J), were purchased from National Laboratory Animal Center and handled according to the guideline from Laboratory Animal Center of National Tsing Hua University, Taiwan. Tramp-C1 cells (2 × 106) were subcutaneously injected into the left limb of the mice. When the tumor size reached 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) was done 3 times at 2, 12, and 24 h postinjection. The temperature change was monitored using an IR camera (Thermo Shot F30, NFC Avio Infrared Technologies Co., Ltd.) during the NIR light irradiation. Tumor size was measured every day using 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 the 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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ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.chemmater.7b04127. Figures of 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; evaluation of theranostic effects of NR@SAs both in vitro and in vivo by cell viability test, photoacoustic images, and tumor growth delay (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Yuh-Chang Sun: 0000-0002-5955-1881 Yu-Fen Huang: 0000-0001-7686-8090 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We appreciate financial support from the Ministry of Science and Technology (Grants NSC 102-2113-M-007-005-MY3, 105745


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2113-M-007-021, 105-2627-M-019-001, 106-2113-M-007-008, and 106-2627-M-019-001) of Taiwan, ROC.

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NOTE ADDED AFTER ASAP PUBLICATION This paper was published ASAP on January 2, 2018, with an incorrect Supporting Information file. The corrected version was reposted on January 12, 2018.

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Serum Albumin

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