Article pubs.acs.org/Langmuir
Controlling E. coli Adhesion on High-k Dielectric Bioceramics Films using Poly(amino acid) Multilayers Neil J. Lawrence,† Jamie M. Wells-Kingsbury,† Marcella M. Ihrig,† Teresa E. Fangman,‡ Fereydoon Namavar,§ and Chin Li Cheung*,† †
Department of Chemistry and Nebraska Center for Materials and Nanosciences, ‡Morrison Microscopy Core Research Facility, University of Nebraska - Lincoln, Lincoln, Nebraska 68588, United States § Department of Orthopaedic Surgery and Rehabilitation, University of Nebraska Medical Center, Omaha, Nebraska 68198, United States S Supporting Information *
ABSTRACT: The influence of high-k dielectric bioceramics with poly(amino acid) multilayer coatings on the adhesion behavior of Escherichia coli (E. coli) was studied by evaluating the density of bacteria coverage on the surfaces of these materials. A biofilm forming K-12 strain (PHL628), a wild-type strain (JM109), and an engineered strain (XL1-Blue) of E. coli were examined for their adherence to zirconium oxide (ZrO2) and tantalum oxide (Ta2O5) surfaces functionalized with single and multiple layers of poly(amino acid) polyelectrolytes made by the layer-by-layer (LBL) deposition. Two poly(amino acids), poly(L-arginine) (PARG) and poly(Laspartic acid) (PASP), were chosen for the functionalization schemes. All three strains were found to grow and preferentially adhere to bare bioceramic film surfaces over bare glass slides. The bioceramic and glass surfaces functionalized with positively charged poly(amino acid) top layers were observed to enhance the adhesion of these bacteria by up to 4-fold in terms of bacteria surface coverage. Minimal bacteria coverage was detected on surfaces functionalized with negatively charged poly(amino acid) top layers. The effect of different poly(amino acid) coatings to promote or minimize bacterial adhesion was observed to be drastically enhanced with the bioceramic substrates than with glass. Such observed enhancements were postulated to be attributed to the formation of higher density of poly(amino acids) coatings enabled by the high dielectric strength (k) of these bioceramics. The multilayer poly(amino acid) functionalization scheme was successfully applied to utilize this finding for micropatterning E. coli on bioceramic thin films.
■
probability for bacterial colonization.13 Other surface modification studies focus on the chemical functionalization of the surface utilizing antibiotics14 or other organic coatings such as hydrogels,15−17 polyelectrolyte films,18,19 poly(lactic-co-glycolic acid),20 and pegylated polypeptides.21 However, detailed understanding of the relationship between the physical properties of these inorganic implant surfaces and the effectiveness of these organic coatings are seldom discussed in the literature. Structural medical implants are often made of or coated with high-k dielectric bioceramic materials such as tantalum oxide (tantala, Ta2O5), and zirconium oxide (zirconia, ZrO2) due to their durability, biocompatibility, and flexibility, which are similar to those of the tissue being replaced. Tantala is widely applied in dental implants because it permits osseointegration with long-term stability.4,13,22 Zirconia was introduced as a bioceramic material more than 25 years ago to replace alumina, with the expectation that it would ameliorate the problems of alumina’s brittleness and therefore related implant failure.23,24
INTRODUCTION The adhesion of bacteria, especially in the form of biofilms, on inorganic surfaces has large economic impacts in both the medical1 and aquatic reclamation2,3 industries. Biofilms formed on the surface of durable inorganic implants, such as dental, joint implants, catheters, and other medical devices can be a source of recurring infections. Biofilms often exhibit a strong resistance to any form of therapy and are difficult to eliminate.2,4,5 The adherence of bacteria to many solid surfaces and their proposed mechanisms have been well-documented in the literature.6−10 The initial stages of adhesion are widely reported as being reversible by subtle changes in the ambient conditions such as food supply or pH. The electrostatic interactions between the lipopolysaccharides on the surface of the bacteria with the surface of the bioceramic are reported as among the major factors responsible for the initial stages of adhesion of the bacteria to an inorganic surface.11 One of the better understood models is the adherence of negatively charged bacteria such as Escherichia coli (E. coli) to positively charged surfaces.9,12 Thus, both morphological and chemical modifications of implant surfaces have been explored to prevent biofilm formation. Surface texturing and materials composition modification strategies are commonly used to decrease the © 2012 American Chemical Society
Received: August 27, 2011 Revised: January 31, 2012 Published: February 17, 2012 4301
dx.doi.org/10.1021/la2033725 | Langmuir 2012, 28, 4301−4308
Langmuir
Article
adjusted with concentrated HCl to 7.4. This buffer was used to make 1000 ppm aqueous solutions of poly(amino acids). Poly(L-arginine) hydrocholoride (PARG, CAS# 26982−20−7; mol. wt. 7500 Da) was purchased from Sigma-Aldrich (Milwaukee, WI)), whereas poly(Laspartic acid) sodium salt (PASP, CAS#: 31871−95−1; mol. wt. 10 000 Da), was obtained from CarboMer, Inc. (San Diego, CA). IBAD Preparation and Characterization of Zirconia and Tantala Bioceramic Films. Optically transparent nanocrystalline zirconia and tantala films used in this study were prepared by ion beam assisted deposition (IBAD) in the Nanotechnology Laboratory at the University of Nebraska Medical Center similarly as described by Namavar et al.33 Briefly, zirconia films with 1−2 μm thickness were deposited onto the glass slides at room temperature by e-beam evaporation of zirconia (Alfa Aesar, Ward Hill, MA; lot # C01P41) at a deposition rate of 1.5−2.5 Å/s while the films were concurrently bombarded with a mixed ion beam of argon and nitrogen. Similar deposition conditions using tantala as the evaporation source and a mixed ion beam of argon and oxygen were applied to produce the tantala films with a thickness of 3 μm. The identities of these bioceramic films were verified by X-ray diffractometry (XRD, Bruker AXS D8 Discover with GADDS, Madison, WI) using a Cu Kα X-ray source with weighted average wavelength of 1.5417 Å. The surface morphology of the oxide films and the glass slides was imaged by atomic force microscopy (AFM) (Multimode Atomic Force Microscope Nanoscope IIIa, Bruker Nano Surfaces Business, Santa Barbara, CA) in tapping mode with force modulation mode silicon probes (NanoWorld AG, Switzerland). These bioceramic film coated glass slides were cut into 0.5 × 0.5 cm2 squares, cleaned with a modified RCA process26,34−36 using the SC-1 solution (1:1:5 v/v = 18 M NH4OH/30% H2O2/UPW) and SC2 solution (1:1:6 v/v = 12 M HCl/30% H2O2/UPW),35 rinsed well and then stored in a 25 mM TRIS buffer (pH = 7.4, 100 mM NaCl) solution before their chemical surface functionalization. Between each cleaning step in the RCA cleaning, the substrates were rinsed in running UPW for 60 s and kept wet in UPW for at least 5 min. Layer-by-Layer Functionalization. The surfaces of the substrates for the bacterial adhesion evaluation were chemically functionalized using the layer-by-layer deposition technique with alternating layers of oppositely charged poly(amino acid) polyelectrolytes.37 To coat a layer of a particular poly(amino acid), the chosen substrate was first immersed and nutated in the appropriate 1000 ppm poly(amino acid) TRIS buffer solution (0.25 mM, pH = 7.4, 100 mM NaCl) of the particular electrolyte for 5 min, followed by rinsing with the TRIS buffer for 30 s. The substrate was then either returned to the TRIS buffer for storage or immersed and nutated in an alternating polyelectrolyte solution for another 5 min. The surface functionalization sequence was repeated until the required number of layers was deposited onto the substrates according to the design of the experiment. Substrates coated with PARG, PARG/PASP, or PARG/ PASP/PARG were fabricated to evaluate the effect of these polyelectrolyte multilayers coated substrates on the adhesion of different strains of E. coli. Adhesion tests were performed on substrates with up to five alternating layers of poly(amino acids). To verify our successful constructions of the LBL multilayer film structures, ellipsometry (M-2000D, J. A. Woollam Co., Lincoln, NE) was applied to confirm the incremental thickness increase in the multilayer films after the deposition of each additional electrolyte layer (see Supporting Information). Microcontact printing of a square array pattern of multilayer polyelectrolyte squares on the bioceramic films was accomplished using a polydimethylsiloxane (PDMS) stamp following wellestablished procedures.38 The substrates were first coated with a PARG/PASP coating using the procedures described above. The stamp was cleaned and functionalized using low pressure air plasma.39 The “inking” of the stamp was achieved by dipping it into a 1000 ppm solution of PARG in TRIS buffer and then drying with nitrogen. The “inked” stamp pattern was then brought into contact with the substrate for 30 s, after which the stamp was removed and the substrate was returned to storage in the TRIS buffer.
The opaqueness of both tantala and zirconia to X-ray fosters their uses in biomedical applications where high contrast X-ray imaging is desirable. Chemical functionalization of bioceramics with polyelectrolytes has been studied for more than 30 years for their roles in affecting the behavior and functions of eukaryotic cells such as cell adhesion on surfaces and cellular development.25 The functions of the polyelectrolytes in these studies depend on their chemical and physical properties. One major advantage of applying polyelectrolytes in these studies is that they become charged under certain pH conditions with reasonable predictability.26,27 Typically, long chain polyelectrolytes used in these studies have a high charge density at physiological pH and are composed of repeating monomers chemically similar to metabolic products found naturally in human bodies.27 Poly(amino acids) are a class of polyelectrolytes consisting of repeating amino acid monomers. Layer-by-layer (LBL) surface functionalization utilizing layers of oppositely charged poly(amino acids) has been well-studied. This technique is very effective at forming uniform layers of polyelectrolytes.27−30 Recently, we and others reported that nanostructured inorganic films with high dielectric strength may allow the accumulation of high charge density to promote preferential adhesion of proteins.31,32 Nonetheless, to the best of our knowledge, detailed understanding of the possible roles of the dielectric strength (k) of the bioceramics on the density of polyelectrolyte coatings adsorbed on these materials and how these possible changes in the charge density of the polyelectrolyte coatings may affect the bacterial adhesion on these functionalized surfaces are currently lacking in the literature. Herein, we present our study of Gram-negative bacterial adhesion on high-k dielectric zirconia and tantala bioceramic films functionalized with polyelectrolyte multilayers made of poly(amino acids). Escherichia coli (E. coli) were chosen as the study model target because they are the most commonly found bacterium in nosocomial biofilms on urinary tract catheters.1,11 Additionally, they are well-studied organisms with many nonvirulent strains which can be exploited to gain further understanding of other similar bacterial species. Wild-type E. coli strain (JM109) which does not spontaneously form biofilms, a biofilm forming E. coli strain (PHL628),5 and an E. coli strain with longer cell length and larger cell volume than that of the wild-type (XL1-Blue) were selected for comparison in this study. Poly(L-arginine) (PARG) and poly(L-aspartic acid) (PASP) were applied as the positively and negatively charged polyelectrolytes respectively, to modulate the charge density of the surfaces of these bioceramic films.7 The extent of bacterial adhesion was evaluated by determining the density bacterial coverage of the surface of each substrate.
■
MATERIALS AND METHODS
Materials. All water used was Millipore UltraPure Water (UPW) of >18 MΩ resistivity and filtered with 0.22 μm membrane. All purchased chemicals were used as received without further purification. Ammonium hydroxide (18 M NH4OH) and hydrogen peroxide (H2O2, 30% in water) were purchased from Fisher Scientific (Pittsburgh, PA). Concentrated hydrochloric acid (12 M HCl) was obtained from EMD Chemicals (Gibbston, NJ). Plain glass slides (Fisher Finest Premium microscope slides, Fisher Scientific (Pittsburgh, PA)) were used as substrates for the bioceramic films. A 25 mM TRIS buffer solution containing 100 mM NaCl was prepared by mixing the appropriate amount of TRIS(hydroxymethyl) aminomethane (TRIS base (Sigma-Aldrich, Milwaukee, WI), sodium chloride, and UltraPure water. The pH of the resulting solution was 4302
dx.doi.org/10.1021/la2033725 | Langmuir 2012, 28, 4301−4308
Langmuir
Article
E. coli Strains Culture. E. coli of three different K-12 strains, JM109 (Invitrogen, Carlsbad, CA), XL1-Blue (Agilent Technologies, Santa Clara, CA), and PHL628 (A gift from Dr. Anthony Hay of Cornell University), were grown in the Luria Broth (LB) (SigmaAldrich, Milwaukee, WI) and tested for their ability to adhere to glass and bioceramic substrates functionalized with poly(amino acids). These strains have all been reported in the literature to have varying degrees of ability to adhere to inert surfaces. E. coli PHL628 was originally isolated for its ability to form biofilms on glass.5 The ability of E. coli to adhere to glass and bioceramics is usually attributed to an outer membrane lipopolysaccharide, which has been reported as one of the major factors responsible for the initial surface adhesion of bacteria on an inert surface.11 The bacteria were cultured in liquid medium inoculated from a freshly grown overnight culture diluted to an OD600 of 0.01. Bioceramic film substrates (bare or coated with polyelectrolyte multilayers) were placed with the side to be studied facing up, in wells of sterile VDX 24-well plates (Hampton Research, Aliso Viejo, CA). Each well contained 1.5 mL of the selected inoculated medium. The plates were incubated stationary at 37 °C in an elevated humidity environment for approximately 12 h until an OD600 of 0.9 was reached in the inoculated medium. Viability Assay. Live/Dead BacLight assay kits (Invitrogen, Carlsbad, CA) were used, according to manufacturer instructions, to evaluate the coverage of bacteria on the test substrates after the incubation. Briefly, each test substrate was first gently or rigorously rinsing depending on the experimental conditions with TRIS buffer. At this point in the procedure, one control sample of each treatment condition was flame-fixed as a dead control. These dead controls were then processed identically to the remaining staining and fixing procedures as the rest of the samples. The samples were then wrapped in aluminum foil to maintain darkness and nutated in 1 mL of 1× Live/Dead BacLight fluorescent dye solution for 20 min. After staining, the test substrates were rinsed again with the TRIS buffer and then immersed in freshly prepared 4% w/v paraformaldehyde (SigmaAldrich, Milwaukee, WI) for 30 min. The substrates were transferred to TRIS buffer for storage. During the staining procedure, and prior to microscopic observation, each sample container was covered with an aluminum foil to minimize exposure to ambient light. According to the Live/Dead BacLight assay, after the staining procedure, all bacteria were stained to fluoresce green, but those which were dead or had damaged membranes were stained to fluoresce red. Live bacteria which had minimally damaged cell membranes often appear orange in this assay. Confocal fluorescence microscopy for all stained samples was performed on an Olympus confocal laser scanning microscope FluoView 500 system consisting of an Olympus IX81 inverted microscope using 488 and 543 nm excitation lasers with a PlanApo/1.45 100× oil lens (Olympus America Inc., Center Valley, PA). Composite fluorescent images of the test substrates made with these two excitation lasers were constructed to illustrate the spatial distribution of live bacteria (green) and dead bacteria (red) on each test substrate. A large set of random imaged areas on each individual substrate were chosen by picking points at set distances from the center of the microscope stage. The bacterial adhesion on different test substrates was evaluated by calculating the percentage of the corresponding surface area coverage with both red and green simultaneous fluorescence of the test substrates using the NIH software package Image-J.40
Figure 1. Photographs of optically transparent (a) zirconia thin film on glass, (b) tantala thin film on glass, and (c) uncoated glass slide.
permits the transmission of the exciting wavelengths when performing the fluorescence microscopy studies. The topography of the three inorganic substrates was measured using atomic force microscopy to determine if surface topography and surface roughness influenced the adhesion of bacteria and cells to these substrates. The obtained AFM images of these surfaces yield a very smooth surface morphology for the tantala films. Both zirconia films and the microscopy glass slide substrates have comparable surface roughness (see Supporting Information). The surface roughness values (Ra) of the zirconia, tantala, and glass substrates were measured to be 2.2, 0.2, and 3.4 nm, respectively, from estimations using 1 μm2 scanned images. Since the roughness values of all substrates were under 5 nm, from the findings in previous works,40,41 the surface morphology of these substrates was unlikely to play a significant role in the affecting adhesion of the E. coli bacteria to the tested surfaces.41,42 Adhesion Behavior of E. coli. The adhesion behavior of the three different strains of E. coli (JM109, XL1-Blue, and PHL628) on glass, tantala, and zirconia surfaces coated with poly(amino acid) polyelectrolytes was evaluated indirectly by assessing the density of “live” E. coli bacteria covering these surfaces after a 12 h incubation and a gentle (10 mL/min) rinse with TRIS buffer. Control experiments with a more rigorous rinsing (60 mL/min) were found to produce similar trend bacterial adhesion results. We postulate that the higher the coverage of “live” bacteria on these surfaces is, the more likely it is that the bacteria tend to find the surface biocompatible and suitable to adhere onto these surfaces for further colonization. The effect of the poly(amino acid) coatings on the surfaces was postulated to enhance or prevent the adhesion of the bacteria based on the polarization of the surface charge. A control comparison of the adhesion of the different strains of E. coli bacteria after 12 h cultures on clean glass, tantala, and zirconia indicated that JM109 was the least adherent of the three strains tested (Figures 2 and 3, left-most columns). The “live” JM109 strain bacteria coverage on these three substrates varied from 3% to 7%. PHL628 strain bacteria exhibited the highest surface coverage (16−35%) on all nonfunctionalized substrates, indicating their ability to significantly adhere to these surfaces. XL1-Blue strain bacteria displayed similar bacterial density coverage (15−28%) as to that of PHL628 on all these substrates (micrographs not shown). PHL628 was found to form biofilms consisting of multiple layers of bacteria in an extracellular matrix up to 20 μm thick. This drastically contrasts the less-than-10-μm thick biofilms of XL1-Blue which are reported to secrete little extracellular matrix. JM109 was not found to form multiple layers of bacteria on these substrates. The coverage density of the bacteria adhered to the surface of the substrates appears to be positively correlated to the dielectric constants of glass, zirconia, and tantala which are reported as 2, 25, and 30, respectively.43−45 Increases in surface coverage density of all three strains of E. coli bacteria were found for all substrates dip-coated with one layer of PARG (second columns of Figures 2 and 3). The
■
RESULTS Physical Properties of Bioceramic Films Coated Substrates. All as-deposited zirconia and tantala films and the glass slide substrates are optically transparent. The zirconia film coatings appear to have a red tint and the tantala coatings have a green tint in color. (Figure 1a,b) Inverted confocal scanning fluorescence microscopy with 488 and 543 nm excitation lasers was used to evaluate the surface coverage and viability of the E. coli. Optical transparence of these substrates 4303
dx.doi.org/10.1021/la2033725 | Langmuir 2012, 28, 4301−4308
Langmuir
Article
Figure 2. Comparison of representative surface area coverage with JM109 E. coli on poly(amino acids) functionalized and nonfunctionalized glass and bioceramic film surfaces. Confocal laser scanning microscopy fluorescent images of live (green) and dead (red) bacteria demonstrate the bacterial viability on these surfaces. Parts a,e,i are images from samples without functionalization on glass, tantala (Ta2O5), and zirconia (ZrO2), respectively. Parts b,f,j are images from samples functionalized with one layer of poly(L-arginine) (PARG) on glass, tantala, and zirconia, respectively. Parts c,g,k are images from samples functionalized with one layer of PARG followed by one layer of poly(L-aspartic acid) (PASP) on glass, tantala, and zirconia, respectively. Parts d,h,l are images from samples functionalized with PARG followed by PASP and a second layer of PARG on glass, tantala, and zirconia, respectively. Scale bars are 20 μm for all figures.
Figure 3. Comparison of representative surface area coverage with PHL628 E. coli on poly(amino acids) functionalized and nonfunctionalized glass and bioceramic film surfaces. Confocal laser scanning microscopy fluorescent images of live (green) and dead (red) bacteria demonstrate the bacterial viability on these surfaces. Parts a,e,i are images from samples without functionalization on glass, tantala (Ta2O5), and zirconia (ZrO2) respectively. Parts b,f,j are images from samples functionalized with one layer of poly(L-arginine) (PARG) on glass, tantala, and zirconia, respectively. Parts c,g,k are images from samples functionalized with one layer of PARG followed by one layer of poly(L-aspartic acid) (PASP) on glass, tantala, and zirconia, respectively. Parts d,h,l are images from samples functionalized with PARG followed by PASP and a second layer of PARG on glass, tantala, and zirconia, respectively. Scale bars are 20 μm for all figures.
4304
dx.doi.org/10.1021/la2033725 | Langmuir 2012, 28, 4301−4308
Langmuir
Article
relative percentage change, in bacterial coverage of the substrates was calculated using eq 1: {[(bacterial coverage on functionalized substrates) − (bacterial coverage on bare substrates)] /bacterial coverage on bare substrates} × 100%
(1)
The change in the coverage for wild-type JM109 was much more significant than the change in coverage by the biofilm forming PHL628. For an example, for the tantala substrates, the relative percentage change in surface coverage for JM109 was 1050% whereas that for PHL 628 was 200%. The surfaces of substrates dip-coated with one layer of PASP were found to be covered by only a statistically insignificant number of bacteria. Similarly few, if any, bacteria were found on the surface of substrates which were first dip-coated in PARG followed by PASP (third columns of Figures 2 and 3). In contrast, substrates with alternating PARG/PASP/PARG dip-coatings (right-most columns of Figures 2 and 3) had high bacterial surface coverage density similar to those of substrates with only one layer of PARG coating. The final coverage percentages of PARG functionalized substrates were very similar for the three strains of bacteria: (a) JM109: glass (46%), tantala (59%), and zirconia (59%); (b) PHL628: glass (48%), tantala (69%), and zirconia (71%); and (c) XL1-Blue: glass (61%), tantala (78%), and zirconia (83%) (Figure 4). All substrates on which the topmost coating layer was PASP showed very few or no adhered bacteria. Effect of Fixation Agents. The use of paraformaldehyde as the fixation agent was not found to bias our observed bacterial adhesion results by over-cross-linking the bacteria to either PARG or PASP coatings. To evaluate the effect of paraformaldehyde fixation treatments, we performed a series of control samples for each bacteria strain on each type of substrate with no polyelectrolyte functionalization, or with PARG or with PASP functionalization without any fixative treatments. Additionally, flame fixation was utilized on each type of substrates with each type of functionalization and each bacterial strain. Under these conditions, as well as after the flame fixation process, it was observed that the initial bacterial surface coverage densities on these substrates were within the standard deviation as those of the samples fixed with paraformaldehyde. For the samples with no fixation treatments, after a time interval ranging between 30 min and 1 h, the surface coverage density of the JM109 strain was observed to begin decreasing and many mobile bacteria were observed swimming in the TRIS buffer solution in microscopy studies. For the flame-fixed samples, the percentages of surface bacteria coverage on the three different substrates were all about the same densities as those of the paraformaldehyde-fixed samples. All of the flame-fixed bacteria fluoresced red, indicating that they were all dead as expected. Effect of Poly(Amino Acid) Multilayers. Layer-by-layer dip-coating was applied with up to seven alternating layers of PARG and PASP to produce the poly(amino acids) multilayer coating on the bioceramic films that were tested. High density bacteria surface coverage on these multilayer coated films was noticed when the topmost layer was PARG. Near-zero or zero surface coverage was found when the topmost layer was PASP instead (third columns of Figures 2 and 3). The density of bacteria surface coverage was unchanged with a greater number of layers and was observed to be solely determined by the
Figure 4. Histograms of surface coverage by E. coli JM109, PHL628, and XL1-Blue on glass, tantala (Ta2O5), and zirconia (ZrO2) coated substrates functionalized using (blue checkers) no poly(amino acids) as controls, (red slash down to the left) one layer of polyelectrolyte (poly(L-arginine) (PARG)), (red slash down to the right) two layers of polyelectrolytes (PARG/poly(L-aspartic acid) (PASP)), and (green hash) three layers of polyelectrolytes (PARG/PASP/PARG) after 12 h of bacterial growth. Error bars represent standard deviations of the bacteria surface coverage.
exposed topmost layer. The bacteria surface coverage on substrates with PASP as the topmost layer indicated little to no adherent bacteria, even when PASP was the sixth layer of the coating. Similar surface coverage for each strain of bacteria on either zirconia or tantala identically treated with different multilayer coatings was also observed. Effect of Poly(Amino Acids) Patterning by PDMS Stamping. When a pattern of square array of PARG coating was stamped on a substrate, there was a higher density of bacteria on the parts of the pattern where the topmost layer was PARG than that of the background where PASP was exposed (Figure 5). There was some lack of clarity and definition near the edges of the bacteria pattern, likely due to non-optimized stamping conditions such as uneven application of the stamp or diffusion of the poly(amino acids) at the edge of the stamp during the microcontact process. The stamped patterns of PARG squares on a more complex multilayer (PARG/PASP/ PARG/PASP) coated substrates can also be applied to produce similar square patterns of bacteria with no discernible decrease in the clarity and definition of the pattern (data not shown). 4305
dx.doi.org/10.1021/la2033725 | Langmuir 2012, 28, 4301−4308
Langmuir
Article
by-layer dip-coating technique on substrates can produce uniform layers of polyelectrolytes with either overall positively charged or negatively charged top surfaces on the multilayer coatings.28,29,50 The increase in the bacterial adhesion on positively charged PARG treated surfaces is possibly due to the electrostatic attraction of the negatively charged bacteria to the positively charged PARG. Similar arguments for same charge repulsion may also be applied to rationalize the drastic decrease in bacterial adhesion on surfaces functionalized with PASP coating, which is negatively charged under physiological pH conditions. The increase in the initial adhesion of PHL628 strain to the surfaces compared to JM109 strain is expected because PHL628 likely has larger negative surface charge density due to the presence of increased lipopolysaccharides expressed on their cell envelopes than ones on wild-type strains.5,7 The treatment of any of our biomaterial surfaces with negatively charged PASP led to the inability of any of the three examined strains of E. coli to adhere to the surface strongly enough to withstand a gentle rinse in TRIS buffer applied to all samples. There was no evidence revealed in our Live/Dead BacLight assays that the application of any of the polyelectrolytes was antiseptic. Fluorescence microscopy of the bacteria observed on any polyelectrolyte treated surfaces showed the same ratio of live (green) to dead (red) as any other treated or control sample. From these observations, we conclude that these PASP coated surfaces do not allow the bacteria to adhere strongly. The ability of the PASP to minimize the initial adhesion of E. coli even after the buildup of several layers indicates that simple treatments of PASP coatings can be utilized to coat implants and other surfaces for antibacterial applications. The micropatterning of bacteria clearly demonstrates that the observed bacteria−surface interaction is a localized effect caused by the topmost layer of polyelectrolyte. Furthermore, the fact that there are far fewer bacteria adhered to a PASP treated surface suggests that the formation of a classical conditioning layer in a nutrient medium can be drastically reduced by treatment with polyelectrolyte multilayers terminated with a negatively charged surface. Significantly, our observed results indicate that bioceramic films have higher bacterial coverage than that of glass substrates. Similar comparison findings were obtained for substrates coated with multilayer poly(amino acids) terminated with PARG as the topmost layer. Since our data strongly suggest the involvement of electrostatic cell−surface interactions in the adhesion process, we postulate that higher density of polyelectrolytes could have been adsorbed on bioceramic films than on glass substrates. From previous theoretical studies of the interactions between a charged object and a flat surface of a medium, an increase in the dielectric constant of the medium would lead to a decrease in the “charge image potential” experienced by the charged object and hence be more energetically favorable for the attraction of the charged object to the surface.31,32 They suggest that inorganic films with high dielectric strength may allow the accumulation of high charge density to promote preferential adhesion of proteins.31,32 Hence, consequently, since our examined bioceramic films have much higher dielectric constants (k) than that of glass (25 (zirconia), 30 (tantala) vs 2 (glass)),43 it is likely that the charged polyelectrolytes may adsorb onto these bioceramic films to form higher density coating than on glass. As a result, higher and positive charge density polyelectrolyte coatings on high-k bioceramic films are expected to cause an increase in
Figure 5. (a) Schematic illustrating the microcontact printing of a poly(L-arginine) (PARG) film pattern onto a poly(amino acid) double-layer (poly(L-arginine/poly(L-aspartic acid) or PARG/PASP) deposited on a zirconia film coated substrate. (Inset) Cross-section legend of poly(amino acids) multilayers. (b) Laser scanning confocal fluorescent image of XL1-Blue E. coli adhered to the poly(amino acids) multilayer pattern made according to part a. (c) Enlarged image of (b). Scale bars for parts b and c are each 20 μm.
■
DISCUSSION The adhesion of an E. coli bacteria to a substrate is a complicated multistage process that involves the transport of the bacteria to the surface, pilli-mediated cell−surface interactions, interaction-induced overexpression of lipoproteins and lipopolysaccharides on the cell envelope, secretion of extracellular matrix, and other cell structure and physiology changes.5,46,47 Though much is known about such a complex adhesion process, few effective biocompatible surface functionalization approaches have been reported to control bacterial colonization on surfaces.8,15,48,49 Here we show that, by appropriately controlling the surface charge and density of multilayers of poly(amino acid) electrolyte coatings, the surface coverage density of adhered E. coli on substrates was manipulated from near 0% to over 50%. Significantly, such polyelectrolyte coatings were found to be more effective in bacterial coverage control on high-k dielectric bioceramic surfaces. Bacterial patterns with spatial resolution down to ten micrometers were illustrated with positively and negatively charged patterned functionalized surfaces. This further suggests the important role of surface charge density in regulatory adhesion control for E. coli on functionalized substrates. Our observed phenomena can be better understood by considering the initial cell−surface interactions that are mainly composed of electrostatic interactions and van der Waals forces.32 Typically, the electrostatic forces dominate the interactions between two objects in aqueous solutions because van der Waals forces are effective only in a much shorter range. The lipopolysaccharides in the Gram-negative bacteria cell envelopes are negatively charged due to the deprotonation of their hydroxyl groups of the saccharides at physiological pH. Thus, E. coli generally have an overall negatively charged cell surface under our experimental conditions.7,11 Our use of layer4306
dx.doi.org/10.1021/la2033725 | Langmuir 2012, 28, 4301−4308
Langmuir
Article
(3) Bishop, P. L. The role of biofilms in water reclamation and reuse. Water Sci. Technol. 2007, 55 (1−2), 19. (4) Di Bonaventura, G.; Spedicato, I.; D’Antonio, D.; Robuffo, I.; Piccolomini, R. Biofilm Formation by Stenotrophomonas maltophilia: Modulation by Quinolones, Trimethoprim-Sulfamethoxazole, and Ceftazidime. Antimicrob. Agents Ch. 2004, 48 (1), 151−160. (5) Vidal, O.; Longin, R.; Prigent-Combaret, C.; Dorel, C.; Hooreman, M.; Lejeune, P. Isolation of an Escherichia coli K-12 Mutant Strain Able To Form Biofilms on Inert Surfaces: Involvement of a New ompR Allele That Increases Curli Expression. J. Bacteriol. 1998, 180 (9), 2442−2449. (6) Das, T.; Sharma, P. K.; Krom, B. P.; van der Mei, H. C.; Busscher, H. J. Role of eDNA on the Adhesion Forces between Streptococcus mutants and Substratum Surfaces: Influence of Ionic Strength and Substratum Hydrophobicity. Langmuir 2011, 27 (16), 10113−10118. (7) Abu-Lail, N. I.; Camesano, T. A. Role of Lipopolysaccharides in the Adhesion, Retention, and Transport of Escherichia coli JM109. Environ. Sci. Technol. 2003, 37 (10), 2173−2183. (8) Gottenbos, B.; Grijpma, D. W.; van der Mei, H. C.; Feijen, J.; Busscher, H. J. Antimicrobial effects of positively charged surfaces on adhering Gram-positive and Gram-negative bacteria. J. Antimicrob. Chemother. 2001, 48 (1), 7−13. (9) Jucker, B.; Harms, H.; Zehnder, A. Adhesion of the positively charged bacterium Stenotrophomonas (Xanthomonas) maltophilia 70401 to glass and Teflon. J. Bacteriol. 1996, 178 (18), 5472−5479. (10) Li, B.; Logan, B. E. Bacterial adhesion to glass and metal-oxide surfaces. Colloids Surf., B 2004, 36 (2), 81−90. (11) Orme, R.; Douglas, C. W. I.; Rimmer, S.; Webb, M. Proteomic analysis of Escherichia coli biofilms reveals the overexpression of the outer membrane protein OmpA. Proteomics 2006, 6 (15), 4269−4277. (12) Vadillo-Rodríguez, V.; Logan, B. E. Localized Attraction Correlates with Bacterial Adhesion to Glass and Metal Oxide Substrata. Environ. Sci. Technol. 2006, 40 (9), 2983−2988. (13) Mitragotri, S.; Lahann, J. Physical approaches to biomaterial design. Nat. Mater. 2009, 8 (1), 15−23. (14) Li, J.; Zhao, Y. Biocompatibility and antibacterial performance of titanium by surface treatment. J. Coat. Technol. Res. 2009, 11 (6), DOI 10.1007/s11998-009-9221-1. (15) Krsko, P.; Kaplan, J. B.; Libera, M. Spatially controlled bacterial adhesion using surface-patterned poly(ethylene glycol) hydrogels. Acta Biomater. 2009, 5 (2), 589−596. (16) Li, P.; Poon, Y. F.; Li, W.; Zhu, H.-Y.; Yeap, S. H.; Cao, Y.; Qi, X.; Zhou, C.; Lamrani, M.; Beuerman, R. W.; Kang, E.-T.; Mu, Y.; Li, C. M.; Chang, M. W.; Jan Leong, S. S.; Chan-Park, M. B. A polycationic antimicrobial and biocompatible hydrogel with microbe membrane suctioning ability. Nat. Mater. 2011, 10 (2), 149−156. (17) Weibel, D. B.; Lee, A.; Mayer, M.; Brady, S. F.; Bruzewicz, D.; Yang, J.; DiLuzio, W. R.; Clardy, J.; Whitesides, G. M. Bacterial Printing Press that Regenerates Its Ink: Contact-Printing Bacteria Using Hydrogel Stamps. Langmuir 2005, 21 (14), 6436−6442. (18) Etienne, O.; Picart, C.; Taddei, C.; Haikel, Y.; Dimarcq, J. L.; Schaaf, P.; Voegel, J. C.; Ogier, J. A.; Egles, C. Multilayer Polyelectrolyte Films Functionalized by Insertion of Defensin: a New Approach to Protection of Implants from Bacterial Colonization. Antimicrob. Agents Chemother. 2004, 48 (10), 3662−3669. (19) Chua, P.-H.; Neoh, K.-G.; Kang, E.-T.; Wang, W. Surface functionalization of titanium with hyaluronic acid/chitosan polyelectrolyte multilayers and RGD for promoting osteoblast functions and inhibiting bacterial adhesion. Biomaterials 2008, 29 (10), 1412−1421. (20) Mouriño, V.; Boccaccini, A. R. Bone tissue engineering therapeutics: controlled drug delivery in three-dimensional scaffolds. J. R. Soc. Interface 2010, 7 (43), 209−227. (21) Boulmedais, F.; Frisch, B.; Etienne, O.; Lavalle, P.; Picart, C.; Ogier, J.; Voegel, J. C.; Schaaf, P.; Egles, C. Polyelectrolyte multilayer films with pegylated polypeptides as a new type of anti-microbial protection for biomaterials. Biomaterials 2004, 25 (11), 2003−2011. (22) Rosentritt, M.; Behr, M.; Burgers, R.; Feilzer, A. J.; Hahnel, S. In Vitro Adherence of Oral Streptococci to Zirconia Core and Veneering Glass-Ceramics. J. Biomed. Mater. Res., B 2009, 91B (1), 257−263.
bacterial coverage on their surfaces. Similarly, the charged lipopolysaccharides on the bacterial cell envelopes may also have higher tendency to attach onto bare bioceramic film surfaces.
■
CONCLUSIONS The use of a poly(amino acids) multilayer surface functionalization strategy was demonstrated to successfully control the density of E. coli bacterial colonization on surfaces of two distinct bioceramics, zirconia, and tantala. The effectiveness of such polyelectrolyte coatings to either increase or decrease the bacterial adhesion density was enhanced when the polyelectrolyte coating applied to these high-k bioceramic surfaces. Coatings of poly(L-arginine) (PARG) layer were observed to promote bacterial adhesion, whereas coatings of poly(L-aspartic acid) (PASP) layers were examined to minimize bacterial adhesion. Coatings with multilayers of alternating oppositely charged poly(amino acids) were illustrated as a robust functionalization scheme to pattern and promote the adhesion of the examined E. coli strains down to micrometer-scale. A simple model was proposed to explain the initial attraction or repulsion of the bacteria to the substrates based on electrostatic interactions between the functionalized coatings on the substrate and the charged surface of the bacterium when in close proximity. A “charge image potential” model suggests that polyelectrolytes may adsorb and form higher density films on inorganic substrates with high dielectric strength (k). Such postulated phenomena may be applied to explain the higher bacterial coverage on PARG coated high-k bioceramics than similarly functionalized glass substrates.
■
ASSOCIATED CONTENT
S Supporting Information *
Atomic force microscopy images of substrates used in this study and ellipsometic spectra of poly(amino acids) multilayer coatings. This material is available free of charge via the Internet at http://pubs.acs.org.
■
AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Notes
The authors declare no competing financial interest.
■
ACKNOWLEDGMENTS We thank Nebraska Research Initiative and University of Nebraska for financial support and the Nebraska Center for Biotechnology for the use of their Microscopy Core facilities. M.M.I. was partially supported by a NASA Nebraska Space Grant fellowship. We are grateful to Dr. Anthony Hay of Cornell University for the gift of PHL628, Dr. Renat Sabirianov, and Dr. Alex Rubinstein for helpful discussions and James Hilfiker of J. A. Woollam Co. Inc. for assistance with obtaining, and modeling ellipsometry data as well as helpful discussions.
■
REFERENCES
(1) Tambyah, P. A.; Knasinski, V.; Maki, D. G. The Direct Costs of Nosocomial Catheters Associated Urinary Tract Infection in the Era of Managed Care. Infect. Control Hosp. Epidemiol. 2002, 23 (1), 27−31. (2) Eberl, H.; Morgenroth, E.; Noguera, D. Mathematical modeling of biofilms; IWA Pub.: London, 2006. 4307
dx.doi.org/10.1021/la2033725 | Langmuir 2012, 28, 4301−4308
Langmuir
Article
(23) Hisbergues, M.; Vendeville, S.; Vendeville, P. Zirconia: Established facts and perspectives for a biomaterial in dental implantology. J. Biomed. Mater. Res., B 2009, 88B (2), 519−529. (24) Chevalier, J. What future for zirconia as a biomaterial? Biomaterials 2006, 27 (4), 535−543. (25) Chanana, M.; Gliozzi, A.; Diaspro, A.; Chodnevskaja, I.; Huewel, S.; Moskalenko, V.; Ulrichs, K.; Galla, H.-J.; Krol, S. Interaction of Polyelectrolytes and Their Composites with Living Cells. Nano Lett. 2005, 5 (12), 2605−2612. (26) Pang, X.; Zhitomirsky, I. Fabrication of Composite Films Containing Zirconia and Cationic Polyelectrolytes. Langmuir 2004, 20 (7), 2921−2927. (27) Ozin, G. A.; Arsenault, A. C. Nanochemistry: A Chemical Approach to Nanomaterials, 1st ed.; The Royal Society of Chemistry: 2005. (28) Caruso, F.; Donath, E.; Möhwald, H. Influence of Polyelectrolyte Multilayer Coatings on Förster Resonance Energy Transfer between 6-Carboxyfluorescein and Rhodamine B-Labeled Particles in Aqueous Solution. J. Phys. Chem. B 1998, 102 (11), 2011−2016. (29) Decher, G. Fuzzy Nanoassemblies: Toward Layered Polymeric Multicomposites. Science 1997, 277 (5330), 1232−1237. (30) Krogman, K. C.; Lowery, J. L.; Zacharia, N. S.; Rutledge, G. C.; Hammond, P. T. Spraying asymmetry into functional membranes layer-by-layer. Nat. Mater. 2009, 8 (6), 512−518. (31) Rubinstein, A.; Sabirianov, R.; Mei, W.; Namavar, F.; Khoynezhad, A. Effect of the ordered interfacial water layer in protein complex formation: A nonlocal electrostatic approach. Phys. Rev. E 2010, 82 (2), 0219151−0219157. (32) Sabirianov, R. F.; Rubinstein, A.; Namavar, F. Enhanced initial protein adsorption on engineered nanostructured cubic zirconia. Phys. Chem. Chem. Phys. 2011, 13 (14), 6597−6609. (33) Namavar, F.; Wang, G.; Cheung, C. L.; Sabirianov, R. F.; Zeng, X. C.; Mei, W. N.; Bai, J.; Brewer, J. R.; Haider, H.; Garvin, K. L. Thermal stability of nanostructurally stabilized zirconium oxide. Nanotechnology 2007, 18 (41), 4157021−4157021−6. (34) Cras, J. J.; Rowe-Taitt, C. A.; Nivens, D. A.; Ligler, F. S. Comparison of chemical cleaning methods of glass in preparation for silanization. Biosens. Bioelectron. 1999, 14 (8−9), 683−688. (35) Kern, W.; Puotinen, D. A. Cleaning solutions based on hydrogen peroxide for use in silicon semiconductor technology. RCA Rev. 1970, 31, 187−206. (36) Onyiriuka, E. C.; Moore, C. B.; Fehlner, F. P.; Binkowski, N. J.; Salamida, D.; King, T. J.; Couillard, J. G. Effect of RCA cleaning on the surface chemistry of glass and polysilicon films as studied by ToFSIMS and XPS. Surf. Interface Anal. 1998, 26 (4), 270−277. (37) Halthur, T. J.; Claesson, P. M.; Elofsson, U. M. Stability of Polypeptide Multilayers As Studied by in Situ Ellipsometry Effects of Drying and Post-Buildup Changes in Temperature and pH. J. Am. Chem. Soc. 2004, 126 (51), 17009−17015. (38) Xu, C.; Taylor, P.; Ersoz, M.; Fletcher, P. D. I.; Paunov, V. N. Microcontact printing of DNA-surfactant arrays on solid substrates. J. Mater. Chem. 2003, 13 (12), 3044−3048. (39) Hong, S. M.; Kim, S. H.; Kim, J. H.; Hwang, H. I. Hydrophilic Surface Modification of PDMS Using Atmospheric RF Plasma. JPCS 2006, 656−661. (40) Piconi, C.; Maccauro, G. Zirconia as a ceramic biomaterial. Biomaterials 1999, 20 (1), 1−25. (41) Stevens, M. M.; George, J. H. Exploring and Engineering the Cell Surface Interface. Science 2005, 310 (5751), 1135−1138. (42) Truong, V. K.; Lapovok, R.; Estrin, Y. S.; Rundell, S.; Wang, J. Y.; Fluke, C. J.; Crawford, R. J.; Ivanova, E. P. The influence of nanoscale surface roughness on bacterial adhesion to ultrafine-grained titanium. Biomaterials 2010, 31 (13), 3674−3683. (43) Chemical Rubber Company; Handbook of chemistry and physics; CRC Press: Cleveland, OH, 2004. (44) Lin, J.; Masaaki, N.; Tsukune, A.; Yamada, M. Ta2O5 thin films with exceptionally high dielectric constant. Appl. Phys. Lett. 1999, 74 (16), 2370−2372.
(45) Harrop, A. P. J; Wanklyn, J. N. The dielectric constant of zirconia. Brit. J. Appl. Phys. 1967, 18, 739−745. (46) Noimark, S.; Dunnill, C. W.; Wilson, M.; Parkin, I. P. The role of surfaces in catheter-associated infections. Chem. Soc. Rev. 2009, 38 (12), 3435−3448. (47) Nel, A. E.; Madler, L.; Velegol, D.; Xia, T.; Hoek, E. M. V.; Somasundaran, P.; Klaessig, F.; Castranova, V.; Thompson, M. Understanding biophysicochemical interactions at the nano-bio interface. Nat. Mater. 2009, 8 (7), 543−557. (48) Yoshida, T.; Nagasawa, T. ε-Poly-L-lysine: microbial production, biodegradation and application potential. Appl. Microbiol. Biotechnol. 2003, 62 (1), 21−26. (49) Etienne, O.; Gasnier, C.; Taddei, C.; Voegel, J.-C.; Aunis, D.; Schaaf, P.; Metz-Boutigue, M.-H.; Bolcato-Bellemin, A.-L.; Egles, C. Antifungal coating by biofunctionalized polyelectrolyte multilayered films. Biomaterials 2005, 26 (33), 6704−6712. (50) Joanny, J. F. Polyelectrolyte adsorption and charge inversion. Eur. Phys. J. B 1999, 9 (1), 117−122.
4308
dx.doi.org/10.1021/la2033725 | Langmuir 2012, 28, 4301−4308
