Highly Stabilized Nanoparticles on Poly-l-Lysine-Coated Oxidized

Subscriber access provided by - Access paid by the | UCSB Libraries

Biological and Medical Applications of Materials and Interfaces

HIGHLY-STABILIZED NANOPARTICLES ON POLY-LLYSINE-COATED OXIDIZED METALS: A VERSATILE PLATFORM WITH ENHANCED ANTIMICROBIAL ACTIVITY. Fiorela Ghilini, Miriam Candelaria Rodriguez Gonzalez, Alejandro Guillermo Miñán, Diego Ezequiel Pissinis, Alberto Hernández Creus, Roberto Carlos Salvarezza, and Patricia Laura Schilardi ACS Appl. Mater. Interfaces, Just Accepted Manuscript • DOI: 10.1021/acsami.8b07529 • Publication Date (Web): 21 Jun 2018 Downloaded from http://pubs.acs.org on June 23, 2018

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Applied Materials & Interfaces

HIGHLY-STABILIZED NANOPARTICLES ON POLY-L-LYSINE-COATED OXIDIZED METALS: A VERSATILE PLATFORM WITH ENHANCED ANTIMICROBIAL ACTIVITY. Fiorela Ghilini1, Miriam C. Rodríguez González2, Alejandro G. Miñán1, Diego Pissinis1, Alberto Hernández Creus2, Roberto C. Salvarezza1, Patricia L. Schilardi1*

1

Instituto de Investigaciones Fisicoquímicas Teóricas y Aplicadas (INIFTA), Facultad de Ciencias

Exactas, UNLP – CONICET, CC16 Suc4 (1900) La Plata, Buenos Aires, Argentina. 2

Área de Química Física, Departamento de Química, Facultad de Ciencias, Universidad de La

Laguna, Instituto de Materiales y Nanotecnología (IMN), 38200, La Laguna, Tenerife, Spain.

*Corresponding author:

P. L. Schilardi Tel. +54 221 4257430 Fax: +54 221 4254642 e-mail: [email protected]; [email protected]

Keywords: titanium functionalization; poly-L-lysine; silver nanoparticles; biofilms; bactericidal

Abstract The increasing incidence of infections in implantable devices has encouraged the search for biocompatible antimicrobial surfaces. In order to inhibit the bacterial adhesion and proliferation on biomaterials, several surface functionalization strategies have been 1 ACS Paragon Plus Environment

ACS Applied Materials & Interfaces 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 2 of 45

developed. However, most of these strategies lead to bacteriostatic effect and only few of these are able to reach the bactericidal condition. In this work, bactericidal surfaces were designed through the functionalization of titanium surfaces with poly-L-lysine (PLL) as mediator for the incorporation of antimicrobial silver nanoparticles (AgNPs). This functionalization influences the adsorption of the particles on the substrate impeding the agglomeration observed when bare titanium surfaces are used, leading to a homogeneous distribution of AgNPs on the surfaces. The antimicrobial activity of this surface has been tested against two different strains, namely Staphylococcus aureus and Pseudomonas aeruginosa. For both strains and different AgNPs sizes, the surface modified with PLL and AgNPs shows a much-enhanced antimicrobial activity in comparison with AgNPs deposited on bare titanium. This enhanced antibacterial activity is high enough to reach bactericidal effect, a condition hard to achieve in antimicrobial surfaces. Importantly, the designed surfaces are able to decrease the bacterial viability more than 5 orders with respect to the initial bacterial inoculum. That means that a relative low load of AgNPs on the PLL-modified titanium surfaces reaches 99.999 % bacterial death after 24 h. The results of the present study are important to avoid infections in indwelling materials, by reinforcing the preventive antibiotic therapy usually dosed throughout the surgical procedure and during the postoperative period. 1. Introduction A generalized surgical practice to improve the patients’ quality life is the incorporation of indwelling materials into the body. These highly biocompatible materials comprise screws, wires, nails, tweezers and/or prostheses. The main function of all these materials is to keep damaged bones in proper alignment as well as replace damaged bone or connective tissue. However, these surgical interventions imply several adverse complications, being the most important the infection of the prosthesis.1 For example, its incidence in primary knee arthroplasties is 2-3%2, being recurrent infections the most common reason for revision

2 ACS Paragon Plus Environment

Page 3 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


surgery.3 It has been estimated that by 2050 bacterial infections due to multidrug-resistant microorganisms will overcome cancer as cause of death.4 This complication is consequence of the formation of biofilms on the biomaterial surface, due to improper sterilization and/or mishandling during the surgical procedure. Bacteria living in biofilms are immersed in an extracellular polymeric substances (EPS) matrix, which constitutes the main protective mechanism against external aggressive agents, increasing the resistance of cells.5 Thus, the search for indwelling material surfaces able to inhibit the bacterial adhesion and the biofilm growth is crucial in order to avoid harmful consequences in implanted patients. Surface modification of implants has emerged as a new alternative to impede these processes. The surface modifying-agent should be such as to avoid the development of antimicrobial resistance. This desirable property arises in response to the great concern worldwide about the increasing resistance to antibiotics6, leading to the need for the development of alternatives to conventional antimicrobial treatments. Importantly, antimicrobial compounds are classified as bacteriostatic (i.e. inhibit the microorganisms growth) or bactericidal (i.e. kill the microorganisms). Bacteriostatic agents need the help of the host defense in order to eliminate microorganisms. Thus, immunocompromised patients are easy target for infections development since the pathogen will grow again once the bacteriostatic agent runs out with the consequent risk of relapse into infection. Under these circumstances, bacteriostatic agents are not enough and the use of bactericidal agents is necessary. In the search for new approaches, silver nanoparticles (AgNPs) have become one of the most studied elements to create antimicrobial surfaces. The antimicrobial activity of silver is very well-known since a long time7 and with the occurrence of AgNPs, a new approach against pathogen microorganisms has emerged to avoid bacterial antibiotic-resistance. In the last years, multiple reports have reviewed and analyzed different theories and methodologies involved in the antibacterial action of AgNPs.8–10 The mechanisms involved in the bacteria killing by AgNPs are not yet clearly understood and they are still controversial. The small size 3 ACS Paragon Plus Environment


Page 4 of 45

and high surfaces to volume ratio of AgNPs allow them to establish a strong interaction with the cell membrane producing a specific bactericidal effect11, as well as increasing the silver ions release when compared with planar silver surfaces.12 Beyond the different proposed mechanisms, it is generally accepted that the Ag(I) ions released from nanoparticles are certainly involved in the AgNPs antibacterial effect.13,14 In addition, it has been recently demonstrated that anisotropic silver nanoparticles are able to efficiently absorb NIR laser radiation, to dissipate it as heat, and then kill bacteria cells by hyperthermia treatment, enhancing the antibacterial effect due to Ag(I) ions release. 13,15 Among the desired properties of antibacterial functionalized surfaces, biocompatibility is crucial for the successful incorporation of the biomaterial into the body. Several coatings has been developed in order to enhance the interaction between the implantable device and surrounding cells

16

. In particular, poly-L-Lysine (PLL) (Figure 1) is one of the most common

cationic polymers to attach and immobilize eukaryotic cells to different surfaces. In fact, the amine groups of the lysine molecules are protonated under physiological conditions (the PLL isoelectric point is 9.6 17) , which make them attractive for the coating of negatively charged surfaces such as glass18, mica19, some polymers20 and oxidized metals, for instance, titanium and aluminum, by electrostatic interactions.21 On the other hand, the positive charges of PLL exposed to the environment make them appropriate for the immobilization of negatively charged species, such as DNA, and eukaryotic cells since, as it is well known, the cell surface carries negative charges. Specially, citrate-capped AgNPs carry negative charge, and thus, are appropriate to be immobilized on PLL layers, making this PLL-AgNPs combination suitable for biocompatible coating of different biomaterials. This combination would enhance the interaction of implantable devices with the biological environment and also would avoid the microbial colonization.


Page 5 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 1. PLL chemical structure at physiological pH. In this work, we explore the surface modification of Ti/TiO2 by PLL and the post functionalization with AgNPs as an efficient platform for avoiding surface bacterial adhesion and proliferation. We have chosen titanium as substrate due to its high strength-to-weight ratio, corrosion resistance and mechanical wear, inert nature and biocompatibility, etc.,

22

which make it widely used in dental and orthopedic implants. Our results show that PLL impedes the agglomeration of AgNPs on the Ti/TiO2 surface, as it has been observed for PLLcoated mica substrates23, being the adsorbed particles stable in liquid media for long times. More important, they exhibit enhanced bactericidal activity (defined as at least three orders decrease in viability) when compared to the initial inoculum of two different strains, namely S. aureus and P. aeruginosa. Thus, the Ti/TiO2/PPL-AgNPs system assayed in this work is an effective and stable platform for avoiding bacterial growth and proliferation on titanium based implantable devices. Fundamental aspects of PLL deposition on mica and glass, as well as the formation of AgNPs layers on these PLL-modified surfaces, have been previously extensively studied 17,23–25 and are out of the aims of this work.



Page 6 of 45

2. Materials and methods 2.1. Silver nanoparticles synthesis and characterization The AgNPs synthesis was carried out according to Frank et al.

26

Briefly, the following

solutions are added in order: 2.0 mL of 1.25 x 10-2 M sodium citrate, 5.0mL of 3.75 x 10-4 M silver nitrate, 5.0 mL of 5.0 x 10-2 M hydrogen peroxide and 0 µL (from now, blue AgNPs) and 20 µL (from now, yellow AgNPs), respectively, of 1.0 x 10-3 M KBr. The silver reduction was achieved by adding 2.5 mL of freshly prepared 5.0 x 10-3M sodium borohydride under vigorously magnetic stirring. This synthesis leads to citrate-capped silver nanoparticles with controlled size, according to the amount of KBr added. The colloidal dispersion was then dialyzed for 2h to eliminate the excess of reactive. The pH of the resulting colloidal suspensions is 7.02 for yellow and 7.34 for blue AgNPs. The final total Ag concentration in the nanoparticle dispersion was determined by inductively coupled plasma optical emission spectrometry (ICP-OES). To this aim, 1 mL of H2O2 5.0 x 10-2 M was added to the AgNPs dispersion. The resulting solution was acidified with HNO3 solution (65 % w/w) to reach 2 % final concentration and finally, ultrapure water was added up to 12 mL. Then, the obtained solution was analyzed. The resulting concentrations are 15.4 ± 0.5 µg/mL and 14 ± 1 µg/mL for yellow and blue AgNPs respectively. TEM imaging was carried out in a Jeol JEM 2100 TEM by dropping 20 µL of the AgNPs dispersion on a TEM grid and dried in air. The nanoparticles size distribution was carried out by using ImageJ software. The size is presented as diameter for nanospheres and nanodiscs and triangle height for nanoprisms. UV-visible spectra were carried out with a Perking Elmer Lambda 35 UV-visible spectrophotometer. The z potential was measured with a Malvern Zetasizer Nano series equipment at 25 oC (detection angle: 173o). Before each run, the original dispersion were


Page 7 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


diluted 1/10 with ultrapure water. The z potential values were calculated by using the Smoluchowski’s approximation for the Henry’s equation27.

2.2 Substrate preparation, functionalization and characterization Titanium discs (1 cm in diameter, 99.9 %, Advent Research Materials) polished to mirror grade with diamond paste were used as substrates. Before each assay, the substrates were washed in ethanol, rinsed with ultrapure water (Milli Q®) and dried under N2 stream. Since Ti surfaces exposed to air or aqueous media are covered by a layer of native oxide (see section 3.1), we will denote these surfaces as Ti/TiO2 from now on. PLL functionalization was carried out by immersing the samples in a 0.1 % w/v α-PLL aqueous solution (Aldrich, used as received) and sonicated for 10 min at 37°C. After that, the PLL-functionalized Ti/TiO2 discs (Ti/TiO2 /PLL) were gently rinsed with ultrapure water and dried in N2 stream. For AgNPs adsorption, a 24 multiwell plate was used. Each sample was located at the bottom of a well and covered by 1 mL of the AgNPs dispersions. After 3 h, the AgNPs-coated Ti/TiO2/PLL (Ti/TiO2/PLL-AgNPs) and Ti/TiO2 (Ti/TiO2/AgNPs) substrates were removed from the wells, rinsed twice with ultrapure water and dried in N2 stream. The total amount of silver on the substrates was determined by ICP-OES: the AgNPs-modified substrates, Ti/TiO2/AgNPs and Ti/TiO2/PLL-AgNPs, were immersed in a 5.0 x 10-2 M hydrogen peroxide solution in order to dissolve the adsorbed AgNPs, sonicated for 10 min and acidified with 2 % HNO3 solution to reach 2 % final concentration (final volume: 12 mL). Then, the resulting solutions were analyzed. Topographic AFM images were acquired in Tapping and PeakForce Tapping modes using a multimode microscope with a Nanoscope V control unit from Bruker. Scan rates of 0.51.2 Hz were used. Measurements were done with RTESP (251-314 kHz, and 40 N/m) tips (from



Page 8 of 45

Bruker). Roughness analysis was carried out from at least 3 different images (2.0 μm× 2.0 μm in size) and calculated as N

∑ (z − z)

2

i

w=

i =1

N

(1)

where N is the number of points considered on the surface, zi the height of the point i on the surface, and z the average height of the N points. Fourier transform infrared (FTIR) spectra were performed in a Varian 660 spectrometer equipped with an attenuated total reflection (ATR) accessory (MIRacle ATR, Pike Technologies) with a ZnSe prism. Contact angle measurements were carried out with a Ramé-Hart 2900 goniometer, by dropping 2 µL of ultrapure water (MilliQ®) on each substrate. Triplicate assays were performed.

2.3 Bacterial Culture Staphylococcus aureus (S. aureus, ATCC 25923) and Pseudomonas aeruginosa (P. aeruginosa, clinical isolate) were grown overnight in nutrient broth (NB; Merck, Darmstadt, Germany) at 37 °C in a rotary shaker (∼170 rpm). Each bacterial inoculum was diluted in NB in order to get ∼108 CFU mL-1 colony-forming units (CFU) mL-1 of bacteria for viability assays. Enumeration of bacterial suspensions was carried out by plate count method.

2.4 Antimicrobial assays. Evaluation of antimicrobial properties of Ti/PLL-AgNPs substrates on preformed biofilms was carried out as described in literature28 with minor modifications. Briefly, bacterial 8 ACS Paragon Plus Environment

Page 9 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


suspensions (∼108 CFU mL-1) were used immediately for the inoculation of the substrates, which were vertically placed in the bacterial suspension and incubated at 35 °C for 2 h. After that, the substrates were removed and gently washed by immersing them in double-distilled sterile water in order to discard cells that were not tightly attached to the surface. Then, the biofilmed-substrates were incubated in a rich phosphate- buffered medium containing 5 g/L glucose, 5 g/L mannitol, and 10 g/ L glycine in phosphate buffer, pH 7, 0.01 M (from now on, GMP) for 24 h at 35 °C in 24-well culture plates. Finally, substrates with biofilms were washed, individually placed in glass tubes for sonication and then quantified by plate count method. The results were expressed per cm2 of each substrate. As controls, bare Ti foils and Ti/PLL substrates were used. The experiments were made thrice and statistical analysis was carried out using one-way analysis of variance (ANOVA) to evaluate differences between groups of bacteria. A p value < 0.05 was considered statistically significant.

2.5 Fluorescence Microscopy. Epifluorescence imaging of S. aureus and P. aeruginosa grown on AgNPs modified substrates for 24 h was determined by using Film Tracer® LIVE/DEAD viability kit (Invitrogen). For both bacterial species, the staining dissolution was prepared by mixing 3μL of component A (SYTO 9) and 3 μL of component B (propidium iodide) in 1 ml of double-distilled sterile water. A total of 40 μL of the mixture was poured on each substrate, and then they were kept in the dark for 15 min at room temperature. After that, the biofilmed-substrates were rinsed with sterile water. Fluorescent bacteria were visualized by epifluorescence with an Olympus BX-51 microscope. The microscope filters used were UMWG2 (excitation 510–550 nm and emission 590 nm) and U-MWB2 (excitation 460–490 and emission 520). Bacteria were kept hydrated throughout the entire procedure.



Page 10 of 45

In order to analyze the EPS produced on the surfaces, either 2 h colonization or 24 h growth, FilmTracer™ SYPRO® Ruby Biofilm Matrix Stain (Invitrogen) was used as received. 150 μL of the staining solution were dropped on each biofilmed surfaces, kept in dark for 30 min and gently washed with sterile water. After that, the samples were imaged by using a UMWG2 (excitation 510–550 nm and emission 590 nm) filter. Bacteria were kept hydrated throughout the entire procedure. 2.6 Immobilized AgNPs stability assays The Ti/PLL-AgNPs were immersed in culture media for 2 h, or in water for 24 h and 5 days, gently washed with ultrapure water and dried in N2 stream. After that, AFM images of the resulting surfaces were taken. The supernatant obtained after 24 h or 5 days in water was used for measuring the Ag(I) ions released from the surfaces. To this aim, each supernatant was acidified with HNO3 to reach 2% final concentration and analyzed by ICP-OES.

3. Results and discussion 3.1. AgNPs and modified- Ti/TiO2 surface characterization The AgNPs were characterized by TEM (Figure 2). The size measurements lead to a maximum in about 12 nm for yellow AgNPs and a bimodal distribution for blue AgNPs, having maxima at 12-15 nm and 40 nm. The height of these nanoparticles was determined by cross sectional analysis of the adsorbed nanoparticles on PLL-modified substrates imaged by AFM. This analysis shows that blue AgNPs are 7 ± 1 nm and 8.3 ± 0.8 nm in height for small and large nanoparticles, respectively, whereas yellow AgNPs are 11 ± 2 in height. Thus, yellow AgNPs consist mainly in nanospheres, whereas blue AgNPs contain nanodiscs (small blue AgNPs) and nanoprisms (large blue AgNPs).


Page 11 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 2. TEM image of blue AgNPs (a) and yellow AgNPs (c). Size distribution of blue AgNPs (b) and yellow AgNPs (d).

The UV-visible spectra of both, yellow and blue AgNPs are presented in Figure 3. For yellow AgNPs, a single sharp band at 407 nm is observed, which is consistent with rather monodisperse nanoparticles 29. In contrast, the spectrum corresponding to blue AgNPs shows three bands located at 331 nm, 418 nm, and 804 nm. These bands are typical for prismatic AgNPs

30,31

and have been assigned to the out-of-plane quadrupole, the in-plane quadrupole,

and the in-plane dipole resonance, respectively. 32,33 On the other hand, the broad band at 804 nm reflects the contribution of different sized and shaped nanoparticles to the surface plasmon resonance (SPR) (cf. Figure 2a). As a matter of fact, the full width at half maximum (FWHM) of the SPR bands is determined by the nanoparticles dispersity, where a large FWHM



Page 12 of 45

is attributed to polydispersity.29 Concerning this, it has been recently demonstrated that broad SPR bands can be deconvoluted by employing lasers tuned to the UV-Vis peaks of gold nanoparticles dispersions, being possible to determine which size and/or shape were responsible for the observed resonances.34

Figure 3. UV-vis spectra for yellow and blue AgNPs.

It is well known that Ti surfaces are covered by a native TiO2 layer. This hydroxylated 35 native oxide layer is mainly formed by amorphous TiO2, about 20 nm thick

36,37

. Because the

isoelectric point of native TiO2 is 4.5, the TiO2 surface charge at physiological pH, i.e. at pH≈7, is negative 38 and then, the surface is suitable for anchoring the positively charged PLL. Figure 4 shows the FTIR-ATR spectra of both Ti/TiO2 and Ti/ TiO2/PLL substrates. As expected, on the Ti surface, the asymmetric stretching mode of CO2 at 2345 cm-1 and small bands corresponding to water (around 1200 and 3400 cm-1) are observed. On the other hand, after PLL adsorption the typical signals related to -NH2, -NH3+, amide I and amide II groups,39,40 are clearly visible on Ti/TiO2/PLL substrates, i.e. the Ti/TiO2 surface has been modified.


Page 13 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 4. FTIR-ATR spectra for Ti/TiO2 and Ti/TiO2/PLL substrates Figure 5a shows an AFM image of the bare polished titanium substrate surface. As it can be seen, it consists in a heterogeneous surface where scratches produced during the polishing process are clearly visible. When the surface is modified with PLL (Figure 5b), a homogeneous organic layer covers the surface. The thickness of the PLL layer is low enough to allow the deeper scratches of the substrate be still visible. Moreover, the surface roughness, measured from 2.0 µm x 2.0 µm AFM images, decreases from w = 4.3 nm to w = 2.8 nm after the incorporation of PLL to the surface. In addition, the contact angle for Ti/TiO2 and Ti/TiO2/PLL surfaces decreases from 71° ± 3° to 61° ± 5° after PLL modification. This change in the wetting properties is consistent with the presence of the PLL layer, as it has been reported by other authors.41 In order to proceed to the nanoparticle adsorption on the PLL-modified surface, the substrates were immersed in the AgNPs dispersions. On this regard, citratecapped AgNPs are negatively charged, since the z potential measured for yellow and blue nanoparticles are -24.4 ± 0.7 mV and -23.8 ± 0.8 mV, respectively. Thus, it is straightforward to expect the adsorption of these nanoparticles on the positively charged Ti/TiO2/PLL surface. It is 13 ACS Paragon Plus Environment


Page 14 of 45

worth to mention that zeta potential values about ±30mV has been used for classifying stable nanoparticles dispersions. However, the zeta potential provides information about the electrostatic repulsive forces, whereas the attractive van der Waals forces are not considered. These van der Waals forces depend, in turn, on the Hamaker constant: if the Hamaker constant is low, the van der Waals attractive forces also become weak and then slight electrostatic repulsion reflected by low zeta potential may be enough for ensure the colloidal stability, 42 as it might be the case for yellow and blue AgNPs used in this work. Figures 5c and 5d show the appearance of the surface after the immersion in blue AgNPs and yellow AgNPs dispersions, respectively. In both cases, a massive adsorption of nanoparticles on the surface takes place. This result is consistent with the electrostatic interactions expected between the positively charged Ti/TiO2/PLL surface and the negatively charged AgNPs. The differently shaped particles can be clearly appreciated in the AFM image shown in Figure 5c. The nanoprisms show crystallographic defined edges as expected by the synthesis method used.


Page 15 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 5. AFM topographic images (2 μm x 2 μm) of (a) bare Ti/TiO2 surface and (b) Ti/TiO2/PLL. AFM images (0.6 μm x 0.6 μm) of Ti/TiO2/PLL-AgNPs for (c) blue and (d) yellow AgNPs, including Cross section analysis (e) and (f) . AFM images (2 μm x 2 μm) for Ti/TiO2/PLL-AgNPs with (g) blue and (h) yellow AgNPs.

Importantly, the distribution of the particles on the surface is homogeneous regardless their size (Figures 5g and 5h), indicating that the PLL modification hinders the agglomeration of the nanoparticles, which leads to the aggregation and a non-homogeneous distribution, a problem that has been observed in the case of bare Ti/TiO2 substrates (without PLL) (Figure 6). 15 ACS Paragon Plus Environment


Page 16 of 45

The deposition of citrate-capped AgNPs on bare Ti/TiO2 substrates was previously studied by some of us. The carboxylic functional groups present in citrate molecules are able to form surface complexes on oxidized titanium surfaces. This strong interaction could be responsible for the high stability of the citrate-capped AgNPs on Ti/TiO2 surfaces.

43,44

Also, it has been

proposed that van deer Waals and hydrogen bonds are responsible for nanoparticlenanoparticle interactions stabilizing the aggregates. 43 Furthermore, the coverage by AgNPs on Ti/TiO2 substrates is lower than Ti/TiO2/PLL surfaces and cannot be increased for a longer immersion time of the substrate in the AgNPs dispersion (data not shown). Quantitative analysis of the total amount of silver on both surfaces leads to the values included in Table 1. Results show that PLL is able to retain larger amounts of silver than the bare substrates. The average amount of silver on Ti/TiO2/PLL-AgNPs is around 4 and 8 times higher than on bare Ti/TiO2 for yellow and blue AgNPs, respectively. However, the average values for yellow and blue AgNPs on both Ti/TiO2/AgNPs and Ti/TiO2/PLL-AgNPs surfaces are not significantly different (p< 0.05). Thus, it can be concluded that, irrespective of the nanoparticles dispersion (yellow AgNPs or blue AgNPs) used for AgNPs surface adsorption, the silver amount is similar on both surfaces. It is worth to mention that under diffusion transport control, it would be expected a higher amount of small yellow AgNPs on the surfaces than blue ones. However, it should be taken into account that two physical-chemical processes are involved in the nanoparticles deposition: diffusion and sedimentation. For small nanoparticles, diffusion is the main driving force, while sedimentation becomes more important as the nanoparticles size increases. Indeed, Cho et al. analyzed the relative contribution of these processes on the cellular uptake of gold nanoparticles by two substrate configurations, upright and inverted 45. These authors found that starting from aliquots of the same nanoparticles dispersion, a similar amount of gold are detected for both configurations when the nanoparticle size is 15 nm, indicating that diffusion prevails over sedimentation. On the contrary, larger nanoparticles (more than 40 nm in size, having low diffusion velocity and high sedimentation velocity) tend 16 ACS Paragon Plus Environment

Page 17 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


to sediment quickly, leading, consequently, to a higher amount of gold on cells. Thus, the similar amount of Ag found on Ti/TiO2/PLL-AgNPs for both, yellow and blue nanoparticles might be related to the quick sedimentation expected of larger nanoprisms.

Figure 6. AFM topographic image (2μm x 2μm) of a bare Ti/TiO2 surface modified with blue AgNPs. Inset: cross section taken along the blue line in the AFM image.

Table 1. Total amount of Ag on the substrates (3 h adsorption time)

Blue AgNPs

Yellow AgNPs

Ti/TiO2/AgNPs Ti/TiO2/PLL-AgNPs Ti/TiO2/AgNPs Ti/TiO2/PLL-AgNPs

Ag (μg.cm-2)

0.34 ± 0.06

2.8± 0.6

0.5 ± 0.2

2.1 ± 0.4

3.2. Antibacterial performance of the modified surfaces. The Ti/TiO2/PLL-AgNPs substrates were tested against sessile Gram(+) and Gram(-) bacteria. In these assays, the substrates were first incubated for 2 h in the bacterial culture, left 24 h in sterile GMP media and then, both sessile and planktonic bacteria were counted. Thus, our experimental model comprises the following stages: (1) surface colonization by bacteria in



Page 18 of 45

broth; (2) bacteria detachment from the colonized surface to the GMP sterile medium (phenotype change from sessile to planktonic cell); (3) both, planktonic and sessile bacterial growth. Accordingly, we have quantified the viable cells, either on the surface or in liquid medium, after stages (1) and (3). 3.2.1. Bacterial adhesion and colonization. We will focus first on the bacterial adhesion and colonization on the different substrates (stage 1) which corresponds to t=0 in Figure 7. In the case of S. aureus, there is a slight decrease in the number of sessile viable bacteria on Ti/TiO2/AgNPs with respect to bare Ti/TiO2 substrates. Indeed, on Ti/TiO2/AgNPs substrates, the number of viable bacteria is reduced by 76.34 % and 62.61 % for blue and yellow nanoparticles respectively, which evidences the bacteriostatic effect of AgNPs adsorbed on the substrates. On the other hand, no significant differences were found in the number of attached P. aeruginosa. This apparent discrepancy has been previously observed by our group and attributed to the EPS secreted during the adhesion step in the formation of P. aeruginosa biofilms.

28

Notice that P. aeruginosa is widely used as an EPS

production model strain.46 In fact, we have stained the proteins in the EPS secreted by sessile bacteria (Figure 8). SYPRO® dyes several biofilm proteins such as glycoproteins, phosphoproteins, lipoproteins, calcium binding proteins, fibrillar proteins, among others. Some of these are found both in the extracellular matrix and in the bacteria, so cells could be also stained and observed in the microscope. From these images, it is clearly noticeable a similar P. aeruginosa EPS production on Ti/TiO2 and Ti/TiO2/AgNPs (Figure 8a), which is consistent with the similar bacterial viability. On this regard, it should be taken into account that, among the polysaccharides identified in P. aeruginosa EPS, negatively charged alginate and Pel were recognized as responsible for the biofilm resistance to cationic antimicrobial compounds, such as aminoglycosides. This fact can be explained in terms of electrostatic interactions between them, which deplete the concentration gradient and interfere with the antimicrobial action.


Page 19 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Consequently, it is reasonable to assume that the similar bacterial viability observed for P. aeruginosa on Ti/TiO2 and Ti/TiO2/AgNPs substrates is due to the interference of EPS on silver release. On Ti/TiO2/PLL-AgNPs the EPS secreted by P. aeruginosa seems to be lower than the above mentioned and, thus, the decrease in the bacterial viability can be assigned to the higher amount of silver on the surface (Table 1). On the other hand, S. aureus excreted a noticeable lower amount of EPS when adhered to all surfaces (Ti/TiO2, Ti/TiO2/AgNPs and Ti/TiO2/PLL-AgNPs) (Figure 8b). Therefore, the EPS would not interfere in the bacteriostatic effect of Ti/TiO2/AgNPs and Ti/TiO2/PLL-AgNPs. On regards Ti/TiO2/PLL, the CFU counted at t=0 does not significantly differ from Ti/TiO2 substrates for both S. aureus and P. aeruginosa, which indicates that PLL does not influence the bacterial adhesion or viability for the strains used in this work. Conte et al. reported the antibacterial activity of PLL aqueous solution on planktonic cells of different bacterial strains, including P. aeruginosa and S. aureus.47 This antibacterial effect by PLL has not been evidenced for the bacterial strains and the experimental conditions used in this work.



Page 20 of 45


Page 21 of 45 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 7. Viable bacteria quantification measured after 24 h in both, surface and liquid media. (a) P. aeruginosa, yellow AgNPs; (b) P. aeruginosa, blue AgNPs; (c) S. aureus, yellow AgNPs; (d) S. aureus, blue AgNPs. For each substrate: (s) t=0: initial viable sessile cells; (s) t=24 h sessile bacteria after 24 h; (p) t=24 h, planktonic bacteria after 24 h.* and ** indicate significant differences (p

Highly Stabilized Nanoparticles on Poly-l-Lysine-Coated Oxidized

Recommend Documents