Functional Silver Nanocomposites as Broad ... - ACS Publications

Research Article www.acsami.org

Functional Silver Nanocomposites as Broad-Spectrum Antimicrobial and Biofilm-Disrupting Agents Qianqian Guo, Yu Zhao, Xiaomei Dai, Tianqi Zhang, Yunjian Yu, Xinge Zhang,* and Chaoxing Li* Key Laboratory of Functional Polymer Materials of Ministry of Education, Institute of Polymer Chemistry, Nankai University, Tianjin 300071, China S Supporting Information *

ABSTRACT: Biofilms’ tolerance has become a serious clinical concern due to their formidable resistance to conventional antibiotics and prevalent virulence. Therefore, there is an urgent need to develop alternative antimicrobial agents to eradicate biofilms but avoid using antibiotics. Herein, we successfully developed polymer functional silver nanocomposites by reduction of silver nitrate in the presence of a biocompatible carbohydrate polymer and a membrane-disrupting cationic polymer. The nanocomposites presented effective antimicrobial activity against Gram-negative bacteria (Pseudomonas aeruginosa, Escherichia coli) and Gram-positive bacteria (Staphylococcus aureus and Bacillus amyloliquefaciens). Confocal laser scanning macroscopy imaging demonstrated that the nanocomposites could efficiently disperse and eradicate the mature biofilms formed by the above four bacterial strains. The introduction of carbohydrate polymers onto nanocomposites effectively improved the biocompatibility, and these nanocomposites induced no significant red blood cell hemolysis and cytotoxicity toward mammalian cells. More importantly, the nanocomposites were able to well eradicate the bacterial biofilms formed on the silicone implants in vivo. In conclusion, the nanocomposites as the broadspectrum biofilm-disrupting agent are significant in the design of new strategies to eradicate biofilms on indwelling medical devices. KEYWORDS: silver nanocomposites, cationic polymer, carbohydrate polymer, antimicrobial activity, biofilm-disrupting

■

INTRODUCTION Biofilms are bacterial communities encapsulated within selfsecreted extracellular polymeric substances, such as exopolysaccharides (EPS), deoxyribonucleic acid (DNA), and lipid, which are highly resilient and difficult to be eradicated.1−4 They occur frequently on synthetic implants and indwelling medical devices including urinary catheters, arthro-prostheses, and dental implants. Bacterial growth in biofilms induces novel behaviors when compared to planktonic bacteria, such as characteristic increased tolerance to antibiotics, stress, and host immunological defense.5 To date, biofilms formed by pathogentic bacteria have been an important cause of chronic infections that are notoriously difficult to treat. On the basis of an understanding of signals and features during biofilm development, the innovative treatment strategies against biofilm infections have been developed in recent years. One strategy is to exploit biofilm self-produced components to only disrupt biofilms by targeting exopolysaccharide matrix but not kill biofilm bacteria.6−10 The other strategy is usage of nanocarriers to deliver antibiotics into biofilms resulting in bacterial death and biofilm removal, which offers an opportunity to reduce the dosage of antibiotics against obliterating biofilms.11−17 For example, Benoit et al. reported a pH-activated nanoparticle for controlled topical delivery of farnesol to treat oral biofilm virulence.16 The farnesol-loaded nanoparticles © XXXX American Chemical Society

improved the efficiency of antibiofilm through multitargeted binding and pH-responsive drug release. Yet the dependence on antibiotics in these approaches cannot prevent the onset of acquired resistance. Another more fashionable strategy is to exploit the antimicrobial materials that can kill bacteria and eradicate biofilms by themselves due to their antimicrobial activity.18−26 Among these antibacterial agents, silver nanoparticles (AgNPs) are probably the most powerful antimicrobial agent that exhibits a strong toxicity toward a broad range of microorganisms, and possibly far lower inclination to induce bacterial resistance as compared to current antibiotic therapies,27−31 which have been used increasingly in catheters, orthopedic, and prosthetic cardiac devices.32,33 Nevertheless, practical applications of AgNPs are often frustrated due to their easily oxidized weakness, which may cause the loss of antimicrobial activity.34−38 To settle this problem, different ligands have been employed to modify AgNPs.23,39−42 For instance, Fei et al. reported a silk fibroin/Ag nanocomposite that was stable in a usual environment (room temperature, exposure to light, and so forth) for at least one month, and meanwhile owned good antibacterial and antibiofilm activity Received: February 24, 2017 Accepted: May 8, 2017 Published: May 8, 2017 A

DOI: 10.1021/acsami.7b02775 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX

Research Article

ACS Applied Materials & Interfaces Scheme 1. Synthetic Routes of (A) PAGA, (B) PDMAEMA-C4, and (C) AgNPs@PAGA/PDMAEMA-C4

■

against the methicillin-resistant Staphylcoccus aureus.39 As a consequence, such a strategy may be a fascinating way to surmount the resistance and tolerance of biofilms as well as put an end to antibiotics in clinical therapeutics. The functional ligands on nanoparticle surface can afford multivalent interactions with biological molecules, allowing nanoparticles to be self-therapeutic agents.43−45 This strategy can avoid the potential limitation of existing antibiotics in nanocarrier systems.46 The resistance of Gram-negative bacteria to most antibiotics arises from the low permeability of the outer membrane. Fortunately, cationic polymers can disrupt bacterial cytoplasmic membranes displaying impressive selectivity for bacterial over mammalian,47,48 and can inhibit growth of planktonic bacteria through strong electrostatic interactions with the negatively charged microbial cell surface.49,50 Therefore, we chose positively charged molecules as ligands of AgNPs to increase the permeability of the outer membrane of bacteria. Of course, it is critical to ensure that the antibacterial agents are nontoxic to human cells. On the basis of the aforementioned rationale and our previous study on antibacterial activity,41,42 alkylated poly(2-(dimethylamino)ethy methacrylate) and poly(2-(acrylamido) glucopyranose) (PAGA) are suitable ligands to functionalize AgNPs as the antibiofilm agents. We investigated the antibacterial activity of polymer-functionalized silver nanocomposites against Gram-positive bacteria (Pseudomonas aeruginosa and Escherichia coli) and Gram-negative bacteria (Staphylococcus aureus and Bacillus amyloliquefaciens). Furthermore, the dispersion and eradication of mature biofilms were explored by in vitro and in vivo study.

MATERIALS AND METHODS

Materials. Silver nitrate (AgNO3, 99.95%, metals basis, Ag 63% min) was purchased from Alfa Aesar (Ward Hill, MA). D-Glucosamine hydrochloride was bought from Beijing Ouhe Technology Co. Ltd. (Beijing, China). 2-(Dimethylamino)ethyl methacrylate (DMAEMA) was bought from Sigma-Aldrich (Saint Louis, MO) and filtered through an alumina column before use. 2,2′-Azobis(isobutyronitrile) (AIBN) was obtained from Heowns Biochem Technologies LLC (Tianjin, China) and recrystallized twice. 4-Cyanopentanoic acid dithiobenzoate (CPADB) was prepared according to the previously reported method.51,52 Sodium borohydride (NaBH4, 98% min) and crystal violet were bought from Aladdin (Shanghai, China). Acridine oange (AO, Fluk) and ethidium (EB, Fluk) were bought from Beijing Solarbio Science & Technology Co., Ltd. (Beijing, China). Methylthiazolyldiphenyl-tetrazolium bromide (MTT) was obtained from J&K China Chemical Co., Ltd. (Beijing, China). Alamar blue was purchased from Yisheng Institute of Biotechnology (Shanghai, China). Bacillus amyloliquefaciens (B. amyloliquefaciens) ATCC 23842, Escherichia coli (E. coli) ATCC 8739, Pseudomonas aeruginosa (P. aeruginosa) ATCC 9027, and Staphylococcus aureus (S. aureus) ATCC 6538 strains were provided by the Department of Microbiology of Nankai University (Tianjin, China). Synthesis of Quaternizated PDMAEMA. First, PDMAEMA was synthesized by reversible additional-fragmentation chain transfer (RAFT) polymerization using DMAEMA as a monomer, CPADB as RAFT agent, and AIBN as an initiator. Briefly, DMAEMA (1.69 mL, 10 mmol), CPADB (27.9 mg, 0.1 mmol), and AIBN (3.28 mg, 0.02 mmol) were dissolved in absolute 1,4-dioxane (4.0 mL) and charged into a polymerization tube. The blending was purged with N2 for 30 min and reacted for 24 h at 70 °C. After the reaction was quenched in an ice bath, PDMAEMA was obtained after the mixture was diluted with tetrahydrofuran (THF) and precipitated into hexane. B


Research Article

ACS Applied Materials & Interfaces

hydrochloride and acryloyl chloride using the modified method.53,54 PAGA was synthesized by RAFT polymerization using AGA as a monomer, CPADB as RAFT agent, and AIBN as an initiator. Briefly, AGA (2.33 g, 10 mmol), CPADB (27.9 mg, 0.1 mmol), and AIBN (3.28 mg, 0.02 mmol) were dissolved in dimethylformamide (DMF)/ H2O (v/v, 9:1) and charged into a polymerization tube. The blending then was purged with N2 for 30 min and reacted at 70 °C for 24 h. After quenching the reaction in an ice bath, PAGA was harvested by precipitating the reaction mixture into ethyl acetate. Preparation and Characterization of Nanocomposites. Typically, AgNO3 (100 μL, 10 mg/mL), PDMAEMA-C4 (21 mg, 2.0 mmol/L), and PAGA (18 mg, 2.0 mmol/L) were added into 4.75 mL of ultrapure water under vigorous stirring at room temperature. Fresh NaBH4 (150 μL, 10 mg/mL) then was rapidly added into the blending, and the reaction was performed for 30 min. The nanocomposites then were centrifuged at 12 580g (Cence, H1850) and washed to remove any unreacted polymer. The obtained silver nanocomposites were denoted as AgNPs@PAGA/PDMAEMA-C4. To better understand the properties of AgNPs@PAGA/PDMAEMA-C4, another two nanocomposites of AgNPs@PAGA and AgNPs@ PDMAEMA-C4 were also prepared with the corresponding polymers using the same method. The surface morphologies of AgNPs and three nanocomposites were analyzed by transmission electron microscopy (TEM, FEI, TecnaiG2F20), for which a droplet of sample was placed on a copper grid and dried at room temperature. Hydrodynamic size (DH) and zeta potential measurements were carried out at 25 °C by dynamic light scattering (DLS) using a Malvern Zetasizer Nano S apparatus equipped with a 4.0 mV laser λ = 636 nm. Hemolysis Assay in Vitro. The hemolysis assay was performed using human blood. Briefly, human blood was centrifuged at 955g for 3 min, and serum was discarded. The red blood cells were washed three times using phosphate buffer solution (PBS, 0.01 M, pH 7.4) and diluted using PBS with the volume fraction of 4%. We used Triton-100 as positive control (+) and PBS as negative control (−). Red blood cells (0.5 mL) were added into different concentrations of nanocomposites (0.5 mL). After being gently shaken, the samples were cocultured at 37 °C for 45 min. After centrifugation, the supernatant was recorded with an absorbance value at 576 nm using UV−vis spectrometer (Shimadzu, UV-2550). The percentage of hemolysis (hemolysis %) was calculated from the following formula:

Figure 1. (A) UV spectra measured by a UV-2550 spectrometer and (B) zeta potential values measured by DLS of AgNPs and three nanocomposites in aqueous solution. Each sample of zeta potential values was analyzed in triplicate, and results were reported as mean ± standard deviation (n = 3). Quaternized PDMAEMA was prepared as follows. PDMAEMA and a 10-fold molar ratio amount of 1-bromobutane were added into 10 mL of absolute ethanol. The reaction system was stirred under reflux at 70 °C for 48 h. The obtained mixture then was dialyzed (3.5 kDa MW cutoff) against ethanol for 48 h, precipitated into cold hexane, and dried. The obtained quaternized polymer was named as PDMAEMA-C4. The quaternization degree was determined by elemental analysis. Synthesis of Poly(2-acrylamido glucopyranose) (PAGA). 2-Acrylamido glucopyranose (AGA) was synthesized with D-glucosamine

hemolysis % =

Abs − Abs(− ) × 100% Abs(+ ) − Abs(− )

(1)

where Abs, Abs(−), and Abs(+) are the absorbances of blending sample, the negative control of PBS, and the positive control of Triton-100, respectively.

Figure 2. Characterization of three nanocomposites: TEM images (A−C) and size distribution measured by DLS (D−F). C


Research Article


Figure 3. Biotoxicity of three nanocomposites. (A) Hemolysis assay, (B) MTT assay, and (C) Alamar blue assay. Triton-100 and PBS were used as positive control (+) and negative control (−) in hemolytic assay, respectively. Each sample of hemolytic assay was analyzed in triplicate (n = 3); both MTT assay and Alamar blue assay were analyzed in quintuplicate (n = 5). All results were reported as mean ± standard deviation.

Figure 4. Growth inhibition of (A) P. aeruginosa, (B) E. coli, (C) S. aureus, and (D) B. amyloliquefaciens in the presence of AgNPs@PAGA, AgNPs@ PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4, respectively. Bacteria treated with PBS were set as negative control (−). Each sample was analyzed in triplicate, and results were reported as mean ± standard deviation (n = 3). Alamar Blue Assay. For Alamar blue assay, 10 μL of Alamar blue solution was added to each well after the coincubation of cells and nanocomposites for 16 h, and the samples were incubated for another 8 h. The fluorescence intensity was read at 530−560 nm (excitation wavelength) and 630 nm (emission wavelength) with a fluorescence spectrometer. The number of viable cells related to the magnitude of dye reduction and was indicated as percentage of Alamar blue reduction.55 The percentage of Alamar blue reduction (Alamar blue reduction (%)) was calculated from the manufacturer’s protocol:

Cytotoxicity Assay. The cytotoxicity of nanocomposites was evaluated using NIH3T3 cells by MTT assay and microplate Alamar blue assay, respectively. Cells were cultured in Dulbecco’s modified Eagles medium (DMEM, Gibco) supplemented with 1% nonessential amino acid, 10% fetal bovine serum, and 1% streptomycin/penicillin in 5% CO2 at 37 °C. Cells were seeded into 96-well plates at a density of 104 cells per well and incubated overnight. The culture medium then was removed, and nanocomposites were diluted to the predetermined concentrations with culture medium and then added into the wells. MTT Assay. For MTT assay, MTT solution (10 μL, 5 mg/mL) was added to each well after the coincubation of cells and nanocomposites for 24 h, and the samples were cultured for another 4 h. The supernatant was discarded, and dimethyl sulfoxide (150 μL) was added to dissolve the formazane crystals. The optical density was read at 492 nm on a microplate reader.

Alamar blue reduction (%) =

(εox λ 2)(Aλ1) − (εox λ1)(Aλ 2) × 100% (εredλ1)(A′λ 2) − (εredλ 2)(A′λ1)

(2) where ελ1 and ελ2 represented the molar extinction coefficients of Alamar blue at 540 and 630 nm, respectively, in the oxidized (εox) and D


Research Article


Figure 5. Antibacterial activity observed by CLSM of (A) P. aeruginosa, (B) E. coli, (C) S. aureus, and (D) B. amyloliquefaciens after treatment with PBS, fluorescence-labeling AgNPs@PAGA, AgNPs@PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4 for 30 min at 37 °C, respectively. The concentrations of AgNPs@PAGA, AgNPs@PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4 were 1.25, 0.16, and 0.16 μmol/L, respectively. Green fluorescence: live bacteria stained by AO in PBS group, or the BODIPY molecule in fluorescence-labeling nanocomposite groups. Red fluorescence: dead bacteria stained by EB. 15 min in darkness. After staining, bacteria were rinsed five times with PBS and imaged using a confocal scanning laser microscope (CLSM, Leica TCS SP5, Germany). Bacteria treated with PBS were set as the control group. Especially, the control group was stained with both EB (10 μL, 1.0 mg/mL) and AO (10 μL, 1.0 mg/mL) at 4 °C for 15 min in darkness. For the observation by scanning electron microscopy (SEM, JSM7500F, Japan), bacterial samples were prepared as per the following method. Glass slides were treated with saturated NaOH/ethanol solution and then rinsed thoroughly with water. The treated bacterial samples were tiled on the glass slides, fixed overnight with 2.5% glutaraldehyde, and washed with water. Bacterial samples then were dehydrated using a series of ethanol solutions (30%, 50%, 70%, 90%, 95%, and 100% v/v in water) and lyophilized. After the treatment, bacterial samples were observed and imaged using SEM. Biofilms’ Formation and Antibiofilm Activity. Biofilms were cultured using the following method. Bacteria were cultured overnight at 37 °C in LB medium and then diluted with the medium to be OD600 = 0.025 as the seeding solution. 100 μL of seeding solution was added into each well of 96-well microplates and incubated for 24 h at 37 °C under static conditions. The formed biofilms were rinsed with

reduced (εred) forms. Aλ1 and Aλ2 were the absorbances of the sample wells at 540 and 630 nm, respectively. A′λ1 and A′λ2 were the absorbances of the negative control wells at 540 and 630 nm, respectively. The values of % Alamar blue reduction were related to the background value of negative control containing medium without cells. Bacterial Growth Inhibitory Assay. Bacteria were cultured overnight at 37 °C in Luria−Bertani (LB) medium and then diluted with medium to be an absorbance of 0.2 at 600 nm (OD600 = 0.2) measured by UV−vis spectrometer. 0.5 mL of bacterial suspension was blended with 0.5 mL of nanocomposites, and then incubated for 8 h at 37 °C. The growth inhibition of bacteria was evaluated by testing OD600. Bacteria suspension treated with an equal volume of PBS was set as the control group. Following inhibitory assay, 10 μL of bacterial suspension was spread onto agar plates, and photos were taken after incubation for 20 h at 37 °C. Microscope Observation. Bacterial suspension (1.5 mL, OD6 00 = 1.0) was centrifuged for 5 min at 1698g and washed three times with PBS. The bacteria were suspended in PBS, and treated for 30 min with fluorescence-labeling AgNPs@PAGA, AgNPs@PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4 (Scheme S1), respectively. The treated bacteria then were stained with EB (10 μL, 1.0 mg/mL) at 4 °C for E


Research Article


(FITC-ConA, Sigma) and EB, respectively, in darkness for 15 min at 4 °C. Finally, the biofilm samples were observed using the CLSM 3D imaging. Following the antibiofilm assay, the residual biofilms were scraped off the glass slides and resuspended in PBS. The samples then were grown on LB agar for 20 h at 37 °C. CFU/mL = volume used to plate on LB agars × dilution factor × average number of colonies. To fully understand the behaviors of bacteria in biofilms, another assay was performed by staining the biofilm samples with AO and EB for 15 min in darkness. After being washed, the samples were observed using CLSM. Mouse Implant Infection Model. Male Sprague−Dawley (SD) mice were bought from Beijing HFK Bioscience Co., Ltd. (Beijing, China). All of the protocols for animal experiments were performed in accordance with the guidelines of the Animal Care and Use Committee of University of Science and Technology of China. The mouse implant infection model was built according to a previously described method with minor modification.9 Biofilms were initially cultured on cylindrical silicone implants (3 mm × 10 mm) in LB medium for 24 h at 37 °C under static conditions. Five SD mice were used per group and anesthetized with ketamine and xylene. The biofilm-coated implants were transferred into the peritoneum of mice after being washed five times with sterile PBS. After 2 h, sterile PBS (3 mL) and AgNPs@PAGA/PDMAEMA-C4 (3 mL, 5 μmol/L) were injected into the peritoneum, respectively. After the infection for 24 h, mice were sacrificed to harvest the implants. The implants were sonicated in 1 mL of PBS for 10 min in an ice−water bath to disrupt and homogenize the biofilm cells from the implants. The samples then were diluted and grown on LB agar for 20 h at 37 °C. CFU/mL = volume used to plate on LB agars × dilution factor × average number of colonies. Statistical Analysis. All data were expressed as mean ± standard deviation of parallel experiments, and the results were analyzed by oneway analysis of Kruskal−Wallis ANOVA. The level of statistical significance was set at *p < 0.05.

Figure 6. SEM images of P. aeruginosa, E. coli, S. aureus, and B. amyloliquefaciens after treatment with PBS, AgNPs@PAGA, AgNPs@ PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4 for 30 min at 37 °C, respectively. The concentrations of AgNPs@PAGA, AgNPs@ PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4 were 1.25, 0.16, and 0.16 μmol/L, respectively. PBS three times to remove planktonic bacteria. Biofilms then were treated with PBS and nanocomposites, respectively. The treated biofilms were rinsed with PBS three times, thoroughly dried, and 150 μL of absolute methanol was added into each well for 15 min. For quantification, 150 μL of crystal violet solution (w/w, 1%) was added to each well for 20 min to stain the biofilms after discarding methanol and drying the plates. After crystal violet was discarded, the biofilm samples were washed with PBS and dried. The stained biofilms then were thoroughly dissolved by adding 200 μL of acetic acid (v/v, 33%) and measured at 595 nm on a microplate reader. Antibiofilm Activity Observed by CLSM Imaging. Bacteria were cultured overnight at 37 °C in LB medium and then diluted with medium to be OD600 = 0.025 as the seeding solution. Sterile coverslips were vertically placed into 24-well microplates, and 400 μL of seeding solution was added into each well. After being incubated for 24 h at 37 °C under static conditions, the coverslips were rinsed with PBS three times to remove planktonic bacteria. Biofilms then were treated with PBS and nanocomposites, respectively. After that, the biofilms were rinsed with PBS three times, fixed with 4% glutaraldehyde for 4 h, and stained with fluorescein isothiocyanate conjugated concanavalin A

■

RESULTS AND DISCUSSION Preparation and Characterization of Nanocomposites. Herein, we prepared the antimicrobial nanocomposites AgNPs@PAGA/PDMAEMA-C4 by NaBH4 reduction of AgNO3 in the presence of a carbohydrate (PAGA) and a cationic polymer (PDMAEMA-C4) (Scheme 1C). Notably, the concentrations of nanocomposites denoted as the theoretical polymer concentrations and the primary concentration of all of the nanocomposites were 400 μmol/L. Two polymers were synthesized by RAFT polymerization, and CPADB acted as chain transfer agent to afford dithioester end group (Scheme 1A,B). PDMAEMA were further quaternized with 1-butyl bromide on the tertiary amino groups to obtain PDMAEMA-C4 (Scheme 1B), and the degree of quaternization was 83.4% estimated by elemental analysis. The information on 1H NMR spectra and

Scheme 2. Eradication of Bacterial Biofilms with AgNPs@PAGA/PDMAEMA-C4

F


Research Article


Figure 7. Disperse percentage of biofilms with different concentrations (from 0.15 to 10 μmol/L) of AgNPs@PAGA, AgNPs@PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4 against (A) P. aeruginosa, (B) E. coli, (C) S. aureus, and (D) B. amyloliquefaciens, respectively. The disperse percentage (%) = the absorbance of nanocomposite group/the absorbance of PBS group × 100% measured at 590 nm. Each sample was analyzed in quintuplicate, and results were reported as mean ± standard deviation (n = 5).

PAGA and PDMAEMA-C4 (Scheme S1A,B). PAGA/fluorescence-labeling PDMAEMA-C4 and fluorescence-labeling PAGA/ PDMAEMA-C4 were used to prepare AgNPs@PAGA/PDMAEMA-C4, respectively (Scheme S1C). The molar ratios of two polymers on the nanocomposites surface were calculated from the fluorescence intensity of BODIPY measured by a fluorescence spectrophotometer (RF-5301PC, Shimadzu). We found that the determined molar ratio of PAGA to PDMAEMA-C4 on AgNPs@PAGA/PDMAEMA-C4 (1:1) was about 1.17 (Table S1). Simultaneously, we used the elemental analysis and sulfuric acid−phenol method to calculate the polymeric content on AgNPs@PAGA/PDMAEMA-C4 (1:1), and found that the total number of two polymers was approximately 20 200 on the surface of each AgNP. It is well-known that AgNPs are easy to aggregate and precipitate in the absence of stabilizer. Herein, two hydrophilic polymers of PAGA and PDMAEMA-C4 were used to functionalize and stabilize AgNPs. We found that three nanocomposites were stable in ultrapure water (Figure S9) and PBS (Figure S10) for 2 months at 37 °C without any aggregation and precipitation, while AgNPs completely precipitated after 2 days in both solutions. These results indicated PAGA and PDMAEMA-C4 were able to effectively stabilize the silver nanoparticles in ultrapure water and PBS. Figure 1 showed the UV spectra and zeta potential values of AgNPs and three nanocomposites. As compared to AgNPs, the maximum absorption showed a red shift from 397 to 439, 427, and 454 nm for AgNPs@PAGA, AgNPs@PDMAEMA-C4, and AgNPs@PAGA/ PDMAEMA-C4, respectively (Figure 1A). Besides, lower peak

molecular weights for three polymers is given in Figures S1−5. To better understand the property of AgNPs@PAGA/ PDMAEMA-C4, another two nanocomposites AgNPs@PAGA and AgNPs@PDMAEMA-C4 were prepared with the corresponding polymers using the same method. To optimize chain length and content of PAGA on AgNPs@ PAGA/PDMAEMA-C4 nanocomposites, zeta potential test, hemolysis assay, and cell viability were performed, and the results are shown in Figures S6−8. Three different molecular weights of PAGA were used to prepare AgNPs@PAGA (AgNPs@PAGA 5 0 , AgNPs@PAGA 1 0 0 , and AgNPs@ PAGA150), and the results indicated that there was no significant difference in the zeta potential and cytotoxicity of AgNPs@PAGA for different molecular weights of PAGA. To ensure the homogeneity of theoretical chain length and quantity, PAGA100 was chosen as the model to prepare AgNPs@PAGA/PDMAEMA-C4 with three different molar ratios of PAGA100 to PDMAEMA-C4 (0.5:1, 1:1, and 2:1). As shown in Figures S6−8, with the increasing content of PAGA on AgNPs@PAGA/PDMAEMA-C4, the zeta potential slightly decreased and the biocompatibility obviously improved. Therefore, AgNPs@PAGA/PDMAEMA-C4 (1:1) was chosen as the model to be discussed. On the basis of the above results, only AgNPs@PAGA100 and AgNPs@PAGA100/PDMAEMA-C4 (1:1) were mentioned below, and they were simplified to be AgNPs@PAGA and AgNPs@PAGA/PDMAEMA-C4 in the present work. To determine the molar ratios of two polymers on AgNPs@ PAGA/PDMAEMA-C4, we synthesized fluorescence-labeling G


Research Article


Figure 8. Disperse percentage of biofilms by AgNPs@PAGA, AgNPs@PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4 at different times (from 0 to 24 h) to (A) P. aeruginosa, (B) E. coli, (C) S. aureus, and (D) B. amyloliquefaciens, respectively. The concentration of three nanocomposites was all 5 μmol/L. The dispersion of percent (%) = the absorbance of nanocomposite group/the absorbance of PBS group × 100% measured at 590 nm. Each sample was analyzed in quintuplicate, and results were reported as mean ± standard deviation (n = 5).

intensity and broader peak width appeared for polymer-functionalized AgNPs when compared to the uncapped AgNPs. The results were possibly caused by a loss of localized surface plasmon resonance (LSPR) absorbance of AgNPs,35,36,56,57 whose position, intensity, and bandwidth were correlated with the size and shape of the nanoparticles.58 The surface modification of AgNPs led to the confinement of free electrons on Ag metal, which strongly influenced the absorbance, leading to the red shift with a higher absorbance or reflection from its original intensity. After conjugating polymers onto AgNPs, zeta potential values were −21.6, 46.5, and 20 mV for AgNPs@PAGA, AgNPs@PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4, respectively, as compared to AgNPs of −33.9 mV (Figure 1B). The morphology observed by TEM (FEI, TecnaiG2F20) and size distribution obtained from DLS of AgNPs and three nanocomposites are shown in Figures 2 and S11. AgNPs and three nanocomposites were well dispersed in size and spherical in shape. The crystalline structures of three nanocomposites were of the cubic phase (Figure S12 and Table S2). The average size of AgNPs@PAGA, AgNPs@PDMAEMA-C4, and AgNPs@ PAGA/PDMAEMA-C4 obtained by TEM was about 6.85 ± 0.86, 11.2 ± 0.24, and 14.1 ± 2.92 nm, respectively, while those determined by DLS were about 19.7 ± 0.25, 27.8 ± 1.04, and 34.1 ± 1.26 nm, respectively. The differences in size obtained by TEM and DLS may be ascribed to the following reasons. When measured by TEM, polymer chains were shrinking in dry state, and only AgNPs were clearly observed, so the size obtained from TEM was mainly the size of AgNPs in the nanocomposites. Yet when measured by DLS, polymer chains were

outstretched in aqueous solution, and the size obtained was related to the whole nanocomposite. Hemolysis and Cytotoxicity Assay. Hemolysis assay was used to evaluate the toxicity of nanocomposites to red blood cells. It was observed (Figure 3A) that the hemolysis percentage of AgNPs@PAGA and AgNPs@PAGA/PDMAEMA-C4 was under the acceptable value of 10% across all nanocomposite concentrations, while AgNPs@PDMAEMA-C4 presented the obvious toxicity to red blood cells above the concentration of 0.63 μmol/L, indicating the introduction of PAGA effectively improved the biocompatibility to red blood cells. The cytotoxicity of nanocomposites against NIH3T3 cells was assessed using MTT assay and Alamar blue assay. The result of MTT assay was shown in Figure 3B. It was observed that the cell viability was over 80% for all concentrations of AgNPs@PAGA, below 0.31 μmol/L of AgNPs@PDMAEMA-C4, and below 2.5 μmol/L of AgNPs@PAGA/PDMAEMA-C4, respectively. That is, AgNPs@PAGA and AgNPs@PAGA/ PDMAEMA-C4 displayed better cell viability than AgNPs@ PDMAEMA-C4 across all concentrations tested. As compared to the MTT assay, Alamar blue assay did not cause toxicity and destruction to cells but was more sensitive on the assessment of cell viability.59,60 The results of the Alamar blue assay were shown in Figure 3C. There was a decreased trend of fluorescence intensity with the increased concentrations for three nanocomposites, indicating an increased toxicity at high concentrations. Also, the introduction of PAGA obviously enhanced the fluorescence intensity of AgNPs@PAGA/PDMAEMA-C4 across all concentrations tested as compared to AgNPs@PDMAEMA-C4. H


Research Article


Figure 9. CLSM 3D images of P. aeruginosa, E. coli, S. aureus, and B. amyloliquefaciens biofilms after the treatment with PBS, AgNPs@PAGA, AgNPs@PDMAEMA-C4, and AgNPs@PDMAEMA-C4/PAGA, respectively. The concentration of three nanocomposites was all 5 μmol/L. Green fluorescence: the EPS stained by FITC-ConA. Red fluorescence: dead bacteria stained by EB.

bacterial cells.65 Therefore, there was a synergistic effect on antibacterial activity between AgNPs and PAGA. The stronger antibacterial activity of AgNPs@PDMAEMA-C4 than PDMAEMA-C4 (Figure S13) was ascribed to the high density of positive charges leading to a powerful electrostatic interaction with the negative bacterial cell membranes and the synergistic interaction with AgNPs.41,42 Therefore, there was a synergistic effect on antibacterial activity among AgNPs, PAGA, and PDMAEMA-C4, leading to more superior antibacterial activity of AgNPs@PAGA/PDMAEMA-C4 than both AgNPs@PAGA and AgNPs@PDMAEMA-C4. Simultaneously, we used agar plates method to evaluate the antibacterial activity of AgNPs@ PAGA, PDMAEMA-C4, and AgNPs@PDMAEMA-C4. As shown in Figure S14, there were much fewer colonies after the treatment with three nanocomposites for four bacteria as compared to PBS groups, indicating all of the three nanocomposites exhibited excellent antibacterial activity. To investigate the interaction of nanocomposites with bacteria, we prepared the fluorescence-labeling nanocomposites with BODIPY-CTA (Scheme S1) to appear as the green fluorescence signals with the excitation/emission at 494/525 nm. The dead cells stained with EB appeared as the red fluorescence signals.

Antibacterial Activity. The antibacterial activity of nanocomposites was explored with P. aeruginosa, E. coli, S. aureus, and B. amyloliquefaciens, and the results were shown in Figures 4 and S13. As shown in Figure 4, the growth of bacteria strains was inhibited at concentrations higher than 0.62, 0.16, and 0.16 μmol/L for AgNPs@PAGA (AgNPs: 5 μg/mL), AgNPs@ PDMAEMA-C4 (AgNPs: 1.25 μg/mL), and AgNPs@PAGA/ PDMAEMA-C4 (AgNPs: 1.25 μg/mL), respectively. In Figure S13, AgNPs@PAGA showed much better antibacterial activity than both AgNPs and PAGA at the same concentration, indicating the introduction of PAGA efficiently enhanced the antibacterial activity of AgNPs. It has been proved that AgNPs can destroy bacteria by several mechanisms,36,61,62 such as releasing Ag+ and inducing to produce reactive oxygen species (ROS), and AgNPs can increase the permeability of bacterial membrane to help biocides enter into the bacterial cells more easily.62 However, the individual AgNPs are easily aggregated to minimize their surface energy, which may lead to limited antimicrobial activity of AgNPs due to a loss of surface area.63,64 Herein, the introduction of PAGA not only stabilized AgNPs (Figures S10 and 11) but made a contribution to the antibacterial activity of AgNPs through glucosamine-mediated penetration into I


Research Article


aforementioned antibacterial activity, it was envisaged that these nanocomposites could be exploited to eradicate the preformed biofilms as shown in Scheme 2. To evaluate the contribution of AgNPs@polymers to biofilm disassembly, biofilms were grown by seeding bacteria (OD600 = 0.025) into a 96-well microplate and incubating for 24 h at 37 °C under static conditions. After the treatment with PBS, AgNPs@PAGA, AgNPs@PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4, respectively, biofilms were washed with PBS three times, fixed with methanol for 15 min, and stained with crystal violet for 20 min. The stained biofilms then were dissolved into 33% acetic acid, and the biomass was measured by a microplate reader at 590 nm. Figure 7 showed the disperse percentage of biofilms with three nanocomposites as a function of concentration against P. aeruginosa, E. coli, S. aureus, and B. amyloliquefaciens biofilms, respectively. It was observed that the disperse percentages of biofilms by AgNPs@PAGA, AgNPs@ PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4 were 40−60%, ∼80%, and ∼80%, respectively. We further evaluated the disperse percentage of biofilms by AgNPs, PAGA, and PDMAEMA-C4 as a function of concentration. As shown in Figure S15, the disperse percentages of biofilms by AgNPs, PAGA, and PDMAEMA-C4 were 10−20%, ∼0%, and 30−50%, respectively. AgNPs@PAGA showed a better disperse effect than both AgNPs and PAGA at the same concentration, presenting a synergistic effect on antibiofilm due to increased stability, facilitating the release of high concentration Ag+ of AgNPs, and glucosamine-mediated penetration of nanocomposites into bacterial cells. AgNPs@PDMAEMA-C4 showed an excellent disperse capability of biofilms that was ascribed to a synergistic effect between AgNPs and PDMAEMA-C4. Moreover, EPS, known as the major component of biofilms, had many roles including enhancing bacterial adhesion, promoting structural development of biofilms, providing a protective barrier, as well as adsorbing and storing nutrients for biofilms growth.66 Also, it was proved that EPS was a polyanion with negative charges under neutral conditions,67,68 which facilitated the positive charges to combine with biofilms and improve antibiofilm capability of AgNPs@PDMAEMA-C4. Therefore, there was a synergistic effect among AgNPs, PAGA, and PDMAEMA-C4, leading to AgNPs@PAGA/PDMAEMA-C4 with the powerful efficacy in antibioflims as compared to AgNPs@PAGA and AgNPs@PDMAEMA-C4 at the same concentration. Figures 8 and S16 showed the disperse percentage of biofilms as a function of time after the treatment with AgNPs, PAGA, PDMAEMA-C4, and three nanocomposites. It was observed that AgNPs and AgNPs@PAGA displayed an enhanced disperse capability over time and achieved the plateau after 12 h, while PDMAEMA-C4, AgNPs@PDMAEMA-C4, and AgNPs@ PAGA/PDMAEMA-C4 rapidly achieved the best disperse effect over 4 h. The rapid and efficient antibiofilm behaviors of AgNPs@PDMAEMA-C4 and AgNPs@PAGA/PDMAEMA-C4 probably derived from the multiply positive charges, which facilitated nanocomposites to combine with the negatively charged EPS of biofilms. For CLSM 3D imaging, biofilm samples were fixed with 4% glutaraldehyde and stained with FITC-ConA and EB in darkness. FITC-ConA was used as a probe to determine the polysaccharides matrix,69 and EB was used to probe dead cells in biofilms. Figures 9 and S17 showed the antibiofilm effect of PBS and three nanocomposites against P. aeruginosa, E. coli, S. aureus, and B. amyloliquefaciens measured by CLSM 3D

Figure 10. Bacterial colonies of (A) the dispersed bacteria in suspension and (B) the remained bacteria in biofilms after biofilms were treated with PBS, AgNPs@PAGA, AgNPs@PDMAEMA-C4, and AgNPs@PAGA/PDMAEMA-C4 for 24 h, respectively. The concentration of three nanocomposites was all 5 μmol/L. Each sample was analyzed in quintuplicate, and results were reported as mean ± standard deviation (n = 5).

Especially, bacteria treated with PBS were stained by EB and AO. CLSM imaging of P. aeruginosa, E. coli, S. aureus, and B. amyloliquefaciens after treatment with PBS and nanocomposites for 30 min was shown in Figure 5. After the treatment with three nanocomposites, red fluorescence was observed, suggesting the bacteria were efficiently killed. The aggregation with green and red fluorescence signals was observed in AgNPs@ PDMAEMA-C4 and AgNPs@PAGA/PDMAEMA-C4 groups, which was ascribed to a strong interaction between the positively charged nanocomposites and negatively charged bacteria via the electrostatic interaction. In the AgNPs@PAGA group, bacteria aggregated and formed smaller clusters due to a weak interaction between AgNPs@PAGA and bacteria. SEM imaging of four bacterial strains was pursued to investigate the morphological changes after the treatment with lethal doses of three nanocomposites (Figure 6). In PBS groups, bacteria exhibited a smooth surface with an unbroken membrane structure. In contrast, the surface of bacteria was seriously broken after treatment with three nanocomposites for 30 min. Collectively, the aforementioned results suggested the introduction of functional ligands onto AgNPs greatly increased the antibacterial effect against both Gram-negative bacteria (P. aeruginosa and E. coli) and Gram-positive bacteria (S. aureus and B. amyloliquefaciens). Antibiofilm Activity of Nanocomposites. Biofilms were inherently resistant to antibiotics and played important roles in many chronic bacterial infections. On the basis of the J


Research Article


Figure 11. Bacterial colonies of biofilms in vivo after treatment with PBS and AgNPs@PDMAEMA-C4/PAGA. (A) P. aeruginosa (p = 0.000), (B) E. coli (p = 0.001), (C) S. aureus (p = 0.009), and (D) B. amyloliquefaciens (p = 0.018) (*p < 0.05). The untreated biofilm group presented the biofilms without further treatment after their formation. The concentration of AgNPs@PAGA/PDMAEMA-C4 was 2.5 μmol/L. CFU/mL = volume used to plate on LB agars × dilution factor × average number of colonies.

Mouse Implant Infection Assay. Synthetic implants such as arthro-prostheses and dental implants played a vital role in modern healthcare fields. However, they were highly vulnerable to colonization by bacterial biofilm leading to the development of persistent implants-associated infections.70 To reduce the complexity, male mice were chosen in the research, because male mice are large, as well as they do not have estrous cycles that can complicate pharmacology.71 Therefore, we used male murine models for implant-associated infections to determine whether AgNPs@PAGA/PDMAEM-C4 could work effectively in the disruption of biofilms in vivo (Figure S19). Implants coated with biofilms were put into the mouse peritoneum and then treated locally with sterile PBS and AgNPs@PAGA/ PDMAEMA-C4, respectively. After 24 h of incubation, all mice were alive. It was observed in Figure 11 that AgNPs@PAGA/ PDMAEMA-C4 reduced the bacterial colonization on the implant as compared to the PBS group, indicating the superior antibiofilm capability in vivo. The reduction of biofilm in PBS group may be ascribed to the host clearance of implant biofilms. These results indicated that AgNPs@PAGA/PDMAEMA-C4 nanocomposites have a potential to eradicate biofilms formed on medical devices in clinic.

imaging. In the control group, the structure of biofilms was intact, and bacteria were dense in biofilms. After the treatment with three nanocomposities, the structure of biofilms was destroyed, and bacteria dramatically reduced. Especially, AgNPs@PDMAEMA-C4 and AgNPs@PAGA/PDMAEMA-C4 displayed better antibiofilm capability than AgNPs@PAGA in accordance with the results in Figures 7 and 8. To fully understand the behaviors of these biofilm communities, another assay was performed by staining the biofilm samples with AO and EB. As shown in Figure S18, there were compact and quite a number of bacteria in PBS groups, and most of the bacteria were live showing green fluorescence. After the biofilm communities were treated with three nanocomposites, especially in AgNPs@PDMAEMA-C4 and AgNPs@PAGA/PDMAEMA-C4 groups, bacteria dramatically reduced and most of them were dead, indicating three nanocomposites could effectively kill the bacteria in biofilms. However, there was a large population of viable cells that remained in biofilms in CLSM images (Figures 9 and S18). To determine the bacteria viability in biofilms and disperse states, bacterial counts were performed, and the results were shown in Figure 10. It was found that three nanocomposites were able to kill the dispersed bacteria in suspension as compared to PBS groups, as well as the biofilm bacteria. However, there were still residual live bacteria for both dispersed bacteria and biofilm bacteria. This phenomenon may be ascribed to the stubborn resistance of bacteria to antibiocides. In a word, our nanocomposite antibiocides could disrupt and eradicate biofilms of Gram-negative bacteria (P. aeruginosa and E. coli) and Gram-positive bacteria (S. aureus and B. amyloliquefaciens).

■

CONCLUSIONS We successfully prepared functionalized silver nanocomposites with a biocompatible carbohydrate polymer (PAGA) and a membrane-disrupting cationic polymer (PDMAEMA-C4) as potent antibacterial and antibiofilm agents. These nanocomposites K


Research Article


(8) Petrova, O. E.; Cherny, K. E.; Sauer, K. The Diguanylate Cyclase GcbA Facilitates Pseudomonas Aeruginosa Biofilm Dispersion by Activating BdlA. J. Bacteriol. 2015, 197, 174−187. (9) Yu, S.; Su, T. T.; Wu, H. J.; Liu, S. H.; Wang, D.; Zhao, T. H.; Jin, Z. J.; Du, W. B.; Zhu, M. J.; Chua, S. L.; Yang, L.; Zhu, D. Y.; Gu, L. C.; Ma, L. Z. PslG, A Self-produced Glycosyl Hydrolase, Triggers Biofilm Disassembly by Disrupting Exopolysaccharide Matrix. Cell Res. 2015, 25, 1352−1367. (10) Sanchez, C. J., Jr; Prieto, E. M.; Krueger, C. A.; Zienkiewicz, K. J.; Romano, D. R.; Ward, C. L.; Akers, K. S.; Guelcher, S. A.; Wenke, J. C. Effects of Local Delivery of D-Amino Acids from Biofilm-Dispersive Scaffolds on Infection in Contaminated Rat Segmental Defects. Biomaterials 2013, 34, 7533−7543. (11) Liu, Y.; Busscher, H. J.; Zhao, B. R.; Li, Y. F.; Zhang, Z. K.; Van der Mei, H. C.; Ren, Y. J.; Shi, L. Q. Surface-Adaptive, Antimicrobially Loaded, Micellar Nanocarriers with Enhanced Penetration and Killing Efficiency in Staphylococcal Biofilms. ACS Nano 2016, 10, 4779− 4789. (12) Duncan, B.; Li, X. N.; Landis, R. F.; Kim, S. T.; Gupta, A.; Wang, L. S.; Ramanathan, R.; Tang, R.; Boerth, J. A.; Rotello, V. M. Nanoparticle-Stabilized Capsules for the Treatment of Bacterial Biofilms. ACS Nano 2015, 9, 7775−7782. (13) Du, J.; Bandara, H. M. N.; Du, P.; Huang, H.; Hoang, K.; Nguyen, D.; Mogarala, S. V.; Smyth, H. D. C. Improved Biofilm Antimicrobial Activity of Polyethylene Glycol Conjugated Tobramycin Compared to Tobramycin in Pseudomonas Aeruginosa Biofilms. Mol. Pharmaceutics 2015, 12, 1544−1553. (14) Cheow, W. S.; Chang, M. W.; Hadinoto, K. Antibacterial Efficacy of Inhalable Levofloxacin-Loaded Polymeric Nanoparticles against E. Coil Biofilm Cells: The effect of Antibiotic Release Profile. Pharm. Res. 2010, 27, 1597−1609. (15) Cheow, W. S.; Chang, M. W.; Hadinoto, K. Antibacterial Efficacy of Inhalable Antibiotic-Encapsulated Biodegradable Polymeric Nanoparticles against E. Coil Biofilm Cells. J. Biomed. Nanotechnol. 2010, 6, 391−403. (16) Horev, B.; Klein, M. I.; Hwang, G.; Li, Y.; Kim, D.; Koo, H.; Benoit, D. S. W. pH-Activated Nanoparticles for Controlled Topical Delivery of Farnesol to Disrupt Oral Biofilm Virulence. ACS Nano 2015, 9, 2390−2404. (17) Zhou, J. Y.; Horev, B.; Hwang, G.; Klein, M. I.; Koo, H.; Benoit, D. S. W. Characterization and Optimization of pH-Responsive Polymer Nanoparticles for Drug Delivery to Oral Biofilms. J. Mater. Chem. B 2016, 4, 3075−3085. (18) Contreras-García, A.; Bucio, E.; Brackman, G.; Coenye, T.; Concheiro, A.; Alvarez-Lorenzo, C. Biofilm Inhibition and DrugEluting Properties of Novel DMAEMA-Modified Polyethylene and Silicone Rubber Surfaces. Biofouling 2011, 27, 123−135. (19) Li, Y.; Fukushima, K.; Coady, D. J.; Engler, A. C.; Liu, S. Q.; Huang, Y.; Cho, J. S.; Guo, Y.; Miller, L. S.; Tan, J. P. K.; Ee, P. L. R.; Fan, W. M.; Yang, Y. Y.; Hedrick, J. L. Broad-Spectrum Antimicrobial and Biofilm-Disrupting Hydrogels: Stereocomplex-Driven Supramolecular Assemblies. Angew. Chem., Int. Ed. 2013, 52, 674−678. (20) Frei, R.; Breitbach, A. S.; Blackwell, H. E. 2-Aminobenzimidazole Derivatives Strongly Inhibit and Disperse Pseudomonas Aeruginosa Biofilm. Angew. Chem., Int. Ed. 2012, 124, 5226−5229. (21) Faure, E.; Falentin-Daudré, C.; Lanero, T. S.; Vreuls, C.; Zocchi, G.; Van De Weerdt, C.; Martial, J.; Jérôme, C.; Duwez, A. S.; Detrembleur, C. Functional Nanogels as Platforms for Imparting Antibacterial, Antibiofilm, and Antiadhesion Activities to Stainless Steel. Adv. Funct. Mater. 2012, 22, 5271−5282. (22) Durmus, N. G.; Taylor, E. N.; Kummer, K. M.; Webster, T. J. Enhanced Efficacy of Superparamagnetic Iron Oxide Nanoparticles Against Antibiotic-Resistant Biofilms in the Presence of Metabolites. Adv. Mater. 2013, 25, 5706−5713. (23) Qin, H.; Cao, H. L.; Zhao, Y. C.; Zhu, C.; Cheng, T.; Wang, Q. J.; Peng, X. C.; Cheng, M. Q.; Wang, J. X.; Jin, G. D.; Jiang, Y.; Zhang, X. L.; Liu, X. Y.; Chu, P. K. In Vitro and in Vivo Anti-Biofilm Effects of Silver Nanoparticles Immobilized on Titanium. Biomaterials 2014, 35, 9114−9125.

presented broad-spectrum antimicrobial and antibiofilm activity against Gram-negative bacteria (P. aeruginosa and E. coli) and Gram-positive bacteria (S. aureus and B. amyloliquefaciens). The introduction of carbohydrate polymers into nanocomposites effectively improved the biocompatibility. Moreover, the obtained nanocomposites efficiently destroyed and eradicated the biofilms in vitro. The mouse implant infection assay indicated nanocomposites efficiently eradicated the bacterial colonization on implants in vivo, which may facilitate their clinical application in biofilm-related infectious diseases.

■

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.7b02775. 1 H NMR spectra and GPC characterization for monomers and polymers, zeta potential, hemolytic assay, cell viability assay, antibacterial activities, photographs of bacterial colonies on agar plates, CLSM images of bacteria for AgNPs, PAGA, PDMAEMA-C4, and three nanocomposites, and photographs of the performed process of mouse implant infection model (PDF)

■

AUTHOR INFORMATION

Corresponding Authors

*Tel.: +86-22-23501645. Fax: +86-22-23505598. E-mail: [email protected]. *E-mail: [email protected]. ORCID

Xinge Zhang: 0000-0003-3399-1659 Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS This work was supported by the National Natural Science Foundation of China (Grant nos. 21174071, 21474055, 51673102, and 81170773), the Natural Science Foundation of Tianjin, China (Grant no. 14JYBJC29400), and the Specialized Research Fund for the Doctoral Program of Higher Education (Grant no. 20130031110014).

■

REFERENCES

(1) Costerton, J. W.; Lewandowski, Z.; Caldwell, D. E.; Korber, D. R.; Lappin-Scott, H. M. Microbial Biofilms. Annu. Rev. Microbiol. 1995, 49, 711−745. (2) Stoodley, P.; Sauer, K.; Davies, D. G.; Costerton, J. W. Biofilms as Complex Differentiated Communities. Annu. Rev. Microbiol. 2002, 56, 187−209. (3) Flemming, H. C.; Wingender, J. The Biofilm Matrix. Nat. Rev. Microbiol. 2010, 8, 623−633. (4) Costerton, J. W.; Stewart, P. S.; Greenburg, E. P. Bacterial Biofilms: A Common Cause of Persistent Infections. Science 1999, 284, 1318−1322. (5) Hall-Stoodley, L.; Costerton, J. W.; Stoodley, P. Bacterial Biofilms: From the Natural Environment to Infectious Diseases. Nat. Rev. Microbiol. 2004, 2, 95−108. (6) Kolodkin-Gal, I.; Cao, S. G.; Chai, L.; Böttcher, T.; Kolter, R.; Clardy, J.; Losick, R. A Self-Produced Trigger for Biofilm Disassembly That Targets Exopolysaccharide. Cell 2012, 149, 684−692. (7) Kolodkin-Gal, I.; Romero, D.; Cao, S. J.; Clardy, J.; Kolter, R.; Losick, R. D-Amino Acid Trigger Biofilm Disassembly. Science 2010, 328, 627−629. L


Research Article


Metastasis by a Self-Therapeutic Nanoparticle. Proc. Natl. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 6700−6705. (44) Zhao, Y. Y.; Tian, Y.; Cui, Y.; Liu, W. W.; Ma, W. S.; Jiang, X. Y. Small Molecule-Capped Gold Nanoparticles as Potent Antibacterial Agents That Target Gram-Negative Bacteria. J. Am. Chem. Soc. 2010, 132, 12349−12356. (45) Li, X. N.; Robinson, S. M.; Gupta, A.; Saha, K.; Jiang, Z. W.; Moyano, D. F.; Sahar, A.; Riley, M. A.; Rotello, V. M. Functional Gold Nanoparticles as Potent Antimicrobial Agents against Multi-DrugResistant Bacteria. ACS Nano 2014, 8, 10682−10686. (46) Capeletti, L. B.; de Oliveira, L. F.; Goncalves, K. D. A.; de Oliveira, J. F. A.; Saito, A.; Kobarg, J.; Santos, J. H. Z. D.; Cardoso, M. B. Tailored Silica-Antibiotic Nanoparticles: Overcoming Bacterial Resistance with Low Cytotoxicity. Langmuir 2014, 30, 7456−7464. (47) Ilker, M. F.; Nüsslein, K.; Tew, G. N.; Coughlin, E. B. Coughlin, Tuning the Hemolytic and Antibacterial Activities of Amphiphilic Polynorbornene Derivatives. J. Am. Chem. Soc. 2004, 126, 15870− 15875. (48) Kenawy, E. R.; Worley, S. D.; Broughton, R. The Chemistry and Applications of Antimicrobial Polymers: A State-of-the-Art Review. Biomacromolecules 2007, 8, 1359−1384. (49) Nederberg, F.; Zhang, Y.; Tan, J. P. K.; Xu, K. J.; Wang, H. Y.; Yang, C.; Gao, S. J.; Guo, X. D.; Fukushima, K.; Li, L. J. Biodegradable Nanostructures with Selective Lysis of Microbial Membranes. Nat. Chem. 2011, 3, 409−414. (50) Ng, V. W. L.; Ke, X. Y.; Lee, A. L. Z.; Hedrick, J. L.; Yang, Y. Y. Synergistic Co-Delivery of Membrane-Disrupting Polymers with Commercial Antibiotics Against Highly Opportunistic Bacteria. Adv. Mater. 2013, 25, 6730−6736. (51) Mitsukami, Y.; Donovan, M. S.; Lowe, A. B.; McCormick, C. L. Water-Soluble Polymers. 81. Direct Synthesis of Hydrophilic StyrenicBased Homopolymers and Block Copolymers in Aqueous Solution via RAFT. Macromolecules 2001, 34, 2248−2256. (52) Thang, S. H.; Chong, Y. K.; Mayadunne, R. T. A.; Moad, G.; Rizzardo, E. A Novel Synthesis of Functional Dithioesters, Dithiocarbamates, Xanthates and Trithiocarbonates. Tetrahedron Lett. 1999, 40, 2435−2438. (53) Ting, S. R. S.; Min, E. H.; Zetterlund, P. B.; Stenzel, M. H. Controlled/Living Ab Initio Emulsion Polymerization via a Glucose RAFT stab: Degradable Cross-Linked Glyco-Particles for Concanavalin A/Fim H Conjugations to Cluster E. Coli Bacteria. Macromolecules 2010, 43, 5211−5221. (54) Guo, Q. Q.; Zhang, T. Q.; An, J. X.; Wu, Z. M.; Zhao, Y.; Dai, X. M.; Zhang, X. G.; Li, C. X. Block Versus Random Amphiphilic Glycopolymer Nanoparticles as Glucose-Responsive Vehicles. Biomacromolecules 2015, 16, 3345−3356. (55) Al-Nasiry, S.; Geusens, N.; Hanssens, M.; Luyten, C.; Pijnenborg, R. The Use of Alamar Blue Assay for Quantitative Analysis of Viability, Migration and Invasion of Choriocarcinoma Cells. Hum. Reprod. 2007, 22, 1304−1309. (56) Gorham, J. M.; Mac Cuspie, R. I.; Klein, K. L.; Fairbrother, D. H.; Holbrook, R. D. UV-Induced Photochemical Transformations of Citrate Capped Silver Nanoparticle Suspensions. J. Nanopart. Res. 2012, 14, 1139−1155. (57) Veerapandian, M.; Lim, S. K.; Nam, H. M.; Kuppannan, G.; Yun, K. S. Glucosamine-Functionalized Silver Glyconanoparticles: Characterization and Antibacterial Activity. Anal. Bioanal. Chem. 2010, 398, 867−876. (58) Wiley, B.; Sun, Y.; Xia, Y. Synthesis of Silver Nanostructures with Controlled Shapes and Properties. Acc. Chem. Res. 2007, 40, 1067−1076. (59) O’Brien, J.; Wilson, I.; Orton, T.; Pognan, F. Investigation of the Alamar Blue (Resazurin) Fluorescent Dye for the Assessment of Mammalian Cell Cytotoxicity. Eur. J. Biochem. 2000, 267, 5421−5426. (60) Munshi, S.; Twining, R. C.; Dahl, R. Alamar Blue Reagent Interacts with Cell-Culture Media Giving Different Fluorescence over Time: Potential for False Positives. J. Pharmacol. Toxicol. Methods 2014, 70, 195−198.

(24) Goswami, S.; Thiyagarajan, D.; Das, G.; Ramesh, A. Biocompatible Nanocarrier Fortified with a Dipyridinium-Based Amphiphile for Eradication of Biofilm. ACS Appl. Mater. Interfaces 2014, 6, 16384−16394. (25) Zhang, A. M.; Mu, H. B.; Zhang, W. X.; Cui, G. T.; Zhu, J.; Duan, J. Y. Chitosan Coupling Makes Microbial Biofilms Susceptible to Antibiotics. Sci. Rep. 2013, 3, 3364. (26) Lee, A. L. Z.; Ng, V. W. L.; Wang, W. X.; Hedrick, J. L.; Yang, Y. Y. Block Copolymer Mixtures as Antimicrobial Hydrogels for Biofilm Eradication. Biomaterials 2013, 34, 10278−10286. (27) Lara, H. H.; Garza-Treviño, E. N.; Ixtepan-Turrent, L.; Singh, D. K. Silver Nanoparticles Are Broad-Spectrum Bactericidal and Virucidal Compounds. J. Nanobiotechnol. 2011, 9, 1−8. (28) Russell, A. D.; Hugo, W. B. Antimicrobial Activity and Action of Silver. Prog. Med. Chem. 1994, 31, 351−370. (29) Dallas, P.; Sharma, V. K.; Zboril, R. Silver Polymeric Nanocomposites as Advanced Antimicrobial Agents: Classification, Synthetic Paths, Applications, and Perspectives. Adv. Colloid Interface Sci. 2011, 166, 119−135. (30) Kumar, A.; Vemula, P. K.; Ajayan, P. M.; Hohn, G. SilverNanoparticle-Embedded Antimicrobial Paints Based on Vegetable Oil. Nat. Mater. 2008, 7, 236−241. (31) Knetsch, M. L. W.; Koole, L. H. New Strategies in the Development of Antimicrobial Coatings: The Example of Increasing Usage of Silver and Silver Nanoparticles. Polymers 2011, 3, 340−366. (32) Catauro, M.; Raucci, M. G.; de Gaetano, F. D.; Marotta, A. Antibacterial and Bioactive Silver-Containing Na2Ox·CaOx·2SiO2 Glass Prepared by Sol-Gel Method. J. Mater. Sci.: Mater. Med. 2004, 15, 831−837. (33) Crabtree, J. H.; Burchette, R. J.; Siddiqi, R. A.; Huen, I. T.; Handott, L. L.; Fishman, A. The Efficacy of Silver-Ion Implanted Catheters in Reducing Peritoneal Dialysis-Related Infections. Peritoneal Dial. Int. 2003, 23, 368−374. (34) Kumar, R.; Münstedt, H. Silver Ion Release from Antimicrobial Polyamide/Silver Composites. Biomaterials 2005, 26, 2081−2088. (35) Taglietti, A.; Arciola, C. R.; D’Agostino, A.; Dacrro, G.; Montanaro, L.; Campoccia, D.; Cucca, L.; Vercellino, M.; Poggi, A.; Pallavicini, P. Antibiofilm Activity of a Monolayer of Silver Nanoparticles Anchored to an Amino-Silanized Glass Surface. Biomaterials 2014, 35, 1779−1788. (36) Chernousova, S.; Epple, M. Silver as Antibacterial Agent: Ion, Nanoparticle, and Metal. Angew. Chem., Int. Ed. 2013, 52, 1636−1653. (37) Yliniemi, K.; Vahvaselka, M.; Van Ingelgem, Y.; Baert, K.; Wilson, B. P.; Terryn, H.; Kontturi, K. The Formation and Characterization of Ultra-Thin Films Containing Ag Nanoparticles. J. Mater. Chem. 2008, 18, 199−206. (38) De Jong, W. H.; Van Der Ven, L. T.; Sleijffers, A.; Park, M. V.; Jansen, E. H.; Van Loveren, H.; Vandebriel, R. J. Systemic and Immunotoxicity of Silver Nanoparticles in an Intravenous 28 Days Repeated Dose Toxicity Study in Rats. Biomaterials 2013, 34, 8333− 8343. (39) Fei, X.; Jia, M. H.; Du, X.; Yang, Y. L.; Zhang, R.; Shao, Z. Z.; Zhao, X.; Chen, X. Green Synthesis of Silk Fibroin-Silver Nanoparticle Composites with Effective Antibacterial and Biofilm-Disrupting Properties. Biomacromolecules 2013, 14, 4483−4488. (40) Loo, C. Y.; Rohanizadeh, R.; Young, P. M.; Traini, D.; Cavaliere, R.; Whitchurch, C. B.; Lee, W. H. Combination of Silver Nanoparticles and Curcumin Nanoparticles for Enhanced Anti-Biofilm Activities. J. Agric. Food Chem. 2016, 64, 2513−2522. (41) Mei, L.; Lu, Z. T.; Zhang, X. G.; Li, C. X.; Jia, Y. X. Polymer-Ag Nanocomposites with Enhanced Antimicrobial Activity against Bacterial Infection. ACS Appl. Mater. Interfaces 2014, 6, 15813−15821. (42) Mei, L.; Lu, Z. T.; Zhang, W.; Wu, Z. M.; Zhang, X. G.; Wang, Y. N.; Luo, Y. T.; Li, C. X.; Jia, Y. X. Bioconjugated Nanoparticles for Attachment and Penetration into Pathogenic Bacteria. Biomaterials 2013, 34, 10328−10337. (43) Arvizo, R. R.; Saha, S.; Wang, E. F.; Robertson, J. D.; Bhattacharya, R.; Mukherjee, P. Inhibition of Tumor Growth and M


Research Article

ACS Applied Materials & Interfaces (61) Sharma, V. K.; Yngard, R. A.; Lin, Y. Silver Nanoparticles: Green Synthesis and Their Antimicrobial Activities. Adv. Colloid Interface Sci. 2009, 145, 83−96. (62) Panácě k, A.; Smékalová, M.; Kilianová, M.; Prucek, R.; Bogdanová, K.; Večeřová, R.; Kolár,̌ M.; Havrdová, M.; Płaza, G. A.; Chojniak, J. Strong and Nonspecific Synergistic Antibacterial Efficiency of Antibiotics Combined with Silver Nanoparticles at Very Low Concentrations Showing No Cytotoxic Effect. Molecules 2016, 21, 26−42. (63) Lok, C. N.; Ho, C. M.; Chen, R.; He, Q. Y.; Yu, W. Y.; Sun, H. Z.; Tam, P. H.; Chiu, J. F.; Che, C. M. Silver Nanoparticles: Partial Oxidation and Antibacterial Activities. JBIC, J. Biol. Inorg. Chem. 2007, 12, 527−534. (64) Liu, Z.; Yan, J.; Miao, Y. E.; Huang, Y.; Liu, T. Catalytic and Antibacterial Activities of Green-Synthesized Silver Nanoparticles on Electrospun Polystyrene Nanofiber Membranes Using Tea Polyphenols. Composites, Part B 2015, 79, 217−223. (65) Panácě k, A.; Kvítek, L.; Prucek, R.; Kolár,̌ M.; Večeřová, R.; Pizúrová, N.; Sharma, V. K.; Nevěcň á, T.; Zbořil, R. Silver Colloid Nanoparticles: Synthesis, Characterization, and Their Antibacterial Activity. J. Phys. Chem. B 2006, 110, 16248−16253. (66) Xue, Z.; Sendamangalam, V. R.; Gruden, C. L.; Seo, Y. Multiple Roles of Extracellular Polymeric Substances on Resistance of Biofilm and Detached Clusters. Environ. Sci. Technol. 2012, 46, 13212−13219. (67) Liu, X. M.; Sheng, G. P.; Luo, H. W.; Zhang, F.; Yuan, S. J.; Xu, J.; Zeng, R. J.; Wu, J. G.; Yu, H. Q. Contribution of Extracellular Polymeric Substances (EPS) to the Sludge Aggregation. Environ. Sci. Technol. 2010, 44, 4355−4360. (68) Mayer, C.; Moritz, R.; Kirschner, C.; Borchard, W.; Maibaum, R.; Wingender, J.; Flemming, H. C. The Role of Intermolecular Interactions: Studies on Model Systems for Bacterial Biofilms. Int. J. Biol. Macromol. 1999, 26, 3−16. (69) Bar-Zeev, E.; Berman-Frank, I.; Girshevitz, O.; Berman, T. Revised Paradigm of Aquatic Biofilm Formation Facilitated by Microgel Transparent Exopolymer Particles. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 9119−9124. (70) Noimark, S.; Dunnill, C. W.; Wilson, M.; Parkin, I. P. The Role of Surfaces in Catheter-Associated Infections. Chem. Soc. Rev. 2009, 38, 3435−3448. (71) Mogil, J. S. Why Sex Matters in Mouse Models. Scientist 2007, 7, 49.

N


Functional Silver Nanocomposites as Broad ... - ACS Publications

Recommend Documents