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Anti-biofilm activity of zinc oxide and hydroxyapatite nanoparticles as dental implant coating materials
Accepted Manuscript Title: Anti-biofilm activity of zinc oxide and hydroxyapatite nanoparticles as dental implant coating materials Author: Elham H. Abdulkareem K. Memarzadeh R.P. Allaker J. Huang J. Pratten D. Spratt PII: DOI: Reference:
S0300-5712(15)30057-9 http://dx.doi.org/doi:10.1016/j.jdent.2015.10.010 JJOD 2537
To appear in:
Journal of Dentistry
Received date: Revised date: Accepted date:
3-6-2015 28-9-2015 15-10-2015
Please cite this article as: Abdulkareem Elham H, Memarzadeh K, Allaker RP, Huang J, Pratten J, Spratt D.Anti-biofilm activity of zinc oxide and hydroxyapatite nanoparticles as dental implant coating materials.Journal of Dentistry http://dx.doi.org/10.1016/j.jdent.2015.10.010 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Anti-biofilm activity of zinc oxide and hydroxyapatite nanoparticles as dental implant coating materials
Anti-biofilm activity of zinc oxide and hydroxyapatite nanoparticles as dental implant coating materials Elham H. Abdulkareema, K. Memarzadehb,c*, R.P. Allakerb, J. Huangc, J. Prattena, D. Spratta aDepartment
of Microbial Diseases, UCL Eastman Dental Institute, University College London, UK; bInstitute of Dentistry, Barts and the
London School of Medicine and Dentistry, Queen Mary University of London, UK; cDepartment of Mechanical Engineering, University College London, UK.
Both authors contributed equally to this research
Short title: Anti-biofilm activity of ZnO/HA nanoparticles for dental implants
* Corresponding Author: Kaveh Memarzadeh (Post-doctoral researcher) Present address: Queen Mary, University of London, Barts and The London School of Medicine and Dentistry, Institute of Dentistry, 4 Newark Street, London, E1 2AT Tel: +44-02078822204
E-mails: [email protected] or [email protected]
Graphical abstract 1
Abstract Objectives: Dental implants are prone to failure as a result of bacterial biofilm accumulation. Such biofilms are often resistant to traditional antimicrobials and the use of antimicrobial nanoparticles as implant coatings may offer a means to control infection over a prolonged period. The objective of this study was to determine the antibiofilm activity of nanoparticulate coated Ti discs using a film fermenter based system.
Methods: Metal oxide nanoparticles of zinc oxide (nZnO), hydroxyapatite (nHA) and a combination (nZnO + nHA) were coated using electrohydrodynamic deposition onto titanium (Ti) discs. Using human saliva as an inoculum, biofilms were grown on coated discs for 96 h in a constant depth film fermenter under aerobic conditions with artificial saliva and periimplant sulcular fluid. Culture viability assays and biofilm thickness measurements were used to assess antimicrobial activity.
Results: Following 96 h, reduced numbers of facultatively anaerobic and Streptococcus spp on all three nano-coated surfaces were demonstrated. The proportion of non-viable microorganisms was shown to be higher on nZnO and composite (nZnO + nHA) coated surfaces at 96 h compared with nHA coated and uncoated titanium. Biofilm thickness comparison also demonstrated that nZnO and composite coatings to be the most effective.
Conclusions: The findings support the use of coating Ti dental implant surfaces with nZnO nanoparticles to provide an antimicrobial function.
Clinical Significance: Current forms of treatment for implant associated infection are often inadequate and may result in chronic infection requiring implant removal and resective / 2
regenerative procedures to restore and reshape supporting tissue. The use of metal oxide nanoparticles to coat implants could provide osteoconductive and antimicrobial functionalities to prevent failure.
Key words: Nanoparticles; Peri-implantitis; Zinc Oxide; Coatings; Biofilm; Antimicrobial.
1. Introduction Despite measures taken to prevent bacterial infection, complications with dental implants persist including those associated with antimicrobial treatment (1, 2). Upon contact, bacteria adhere, multiply and form a biofilm on the untreated surface of an implant (3). To help overcome this issue, implants can either be coated or impregnated with antimicrobial agents. However, many such approaches have been unsuccessful and resistance to those antimicrobials employed is often observed. In these cases, it is the complex structure of the associated microbial community that makes it difficult for antimicrobial agents to penetrate (4). Resistance can also involve other mechanisms (5), including microbial efflux systems that pump antimicrobial agents out of the cell (6) and the production of exopolymers which may additionally prevent penetration of agents and thus lead to the persistence of the microbial community (7). Failure or loosening of an osseointegrated implant will allow for further microbial growth, possibly resulting in peri-implantitis with associated soft and hard tissue damage (8). While the microbiota associated with oral health, including that around implants consists of predominantly Gram positive species that associated with oral infection and infected implants is predominantly Gram negative and includes Porphyromonas gingivalis, Fusobacterium spp. and Treponema denticola (9). The presence of such pathogenic bacteria in the oral cavity and other biofilms has led to the development of prophylactic approaches that
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involve the use of single or multiple antibiotics (10). However, such antibiotic treatments provide opportunities for resistance to develop when inappropriate doses are used to prevent biofilm formation (11-13). In particular, antibiotics with a single target will encourage resistance and therefore reduced efficacy of the drug. There is thus a need to investigate alternative approaches as opposed to the use of traditional antimicrobial agents (9). This may be possible by exploitation of the antimicrobial ability of metal based antimicrobial nanoparticles (NPs) to control biofilm formation. Their unique physiochemical properties should help avoid antimicrobial resistance, as activity is thought to involve multiple targets thus hindering rapid microbial adaptation and evolution (10-12). Metallic NPs possess bactericidal effects against a variety of species (13), withstand high temperatures required for fixation and possess low toxicity to mammalian cells when stabilised onto surfaces (14, 15). In order to utilise such antimicrobial capability, appropriate coating methods are required. Generally, coatings used for dental implants employ extreme conditions such as very high temperatures that can alter the properties of the coatings or surfaces (16). These conditions are often intended for stability and biological enhancement of the implant’s surface, however with respect to NPs, it is more suitable to use lower temperatures to retain beneficial physiochemical properties. Electrohydrodynamic atomisation (EHDA), a simple and economical spraying technique allows for the implementation of such conditions (17). This method produces a uniform and stable coating at room temperature whilst covering a large area and so enables sufficient functionality without compromising the original state of the coating (17). Zinc oxide NPs (nZnO) are currently being investigated for their antimicrobial activity and show much promise (18, 19). Studies suggest that the antimicrobial effects of these NPs is due to ion release or via the production of reactive oxygen species (ROS) (20). It is likely that the antimicrobial properties of NPs will differ according to their shape, surface area, size and
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relative chemical properties and thus it is these characteristics that lead to the overall synergistic activity (21). Recent studies indicate that the utilisation of nZnO also allows for the promotion of bone growth in addition to enhanced osteoblast proliferation. (22, 23). Furthermore, investigations involving HA as a coating material indicate a stable interface can be formed between bone and implant (24) . With respect to dentistry, calcium phosphates have played a key role due to their physical and chemical similarities to bone and mineralised tissues, such as enamel and dentine (25). Hence, nano-scale HA (nHA) closely resembles the size and properties of HA crystals in natural bone (26). Furthermore, studies suggest that nHA has the ability to inhibit the growth of both Gram-negative and Gram-positive bacteria, including Staphylococcus aureus and Staphylococcus epidermidis (27). The current investigation was performed using a constant depth film fermenter (CDFF) in order to improve our understanding of the antimicrobial activity of nZnO, nHA and their composite. It is hypothesized that such coating materials will be able to prevent infection as a cause of early or late dental implant failure.
2. Materials and Methods 2.1 Nanoparticles The NPs tested were: nZnO, nHA and a combination of 50% nZnO and 50% nHA. Nano-ZnO particulates were synthesised using flame pyrolysis as carried out by Johnson Matthey plc (JMTC) and nHA were synthesised as previously described (28). The surface area for nZnO was determined by Brunauer Emmett Teller (BET) analysis at JMTC.
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2.2 Ti samples Grade 23 moderate smooth machined Ti discs (Hyundai Hit 8S - A C Service Group, Fordingbridge, UK) 5 mm with a diameter and 2 mm height were used in all experiments. The Ti discs were sonicated with acetone for 20 min at 50 Hz in order to remove adherent debris or dust and then autoclaved before coating.
2.3 EHDA spraying Nano-HA, nZnO and their combination of equal proportions were used as suspensions for electrohydrodynamic spraying. This method uses a fine jet containing NPs which are deposited onto the surface of substrates. In the present study, NPs were suspended in 100% ethanol and jetted via a needle under an electric field. A stainless steel needle with a diameter of 300 μm was used to spray the content of a 1 mL syringe onto the surface of the sample. The outcome is a symmetrical cone shaped jet spray which delivers an optimal coating to ensure coverage of the sample. The distance between the substrate and the needle was maintained at 30 mm at all times. All experiments were performed with a freshly prepared suspension (10,000 µg/mL) of NPs. The voltage used was between 4.5 – 5 kV with Ti discs coated for a period of 1 min at a flow rate of 5 µL/min. All coated and uncoated samples were subjected to heat treatment (600°C) for 1 h. Samples were gradually cooled to maintain mechanical integrity of the coating (1°C/min). 2.4 Saliva samples In order to produce microcosm biofilms, saliva (unstimulated) was collected aseptically from ten members of the Department of Microbial Diseases, Eastman Dental Institute, UCL. These individuals had no significant oral disease (Ethical approval obtained from the UCL Ethics Committee; Project no. 1364/001). The pooled samples were resuspended in 10% (v/v) 6
glycerol and vortex mixed vigorously for 1 min to homogenise. Aliquots of 1 mL were then dispensed and stored at -80°C for subsequent use.
2.5 Constant depth film fermenter A vial of stored pooled saliva was thawed and 900 μL was used to inoculate 500 mL of artificial saliva. To make 1 L of artificial saliva 0.5 g 'Lab-Lemco' powder (Oxoid, UK), 1 g yeast extract (Oxoid, UK), 2.5 g protease peptone (Oxoid, UK), 1.25 g hog gastric mucin type III, partially purified from porcine stomach (Sigma Chemicals Co., Poole, UK), 0.18 g sodium chloride (BDH Chemicals Ltd, Poole, UK), 0.1 g calcium chloride (BDH) and 0.1 g potassium chloride (BDH) was added to deionised water. After autoclaving at 121°C, 0.625 ml of a 40% urea (Sigma-Aldrich) solution was added. This was mixed and pumped into the CDFF for 6 h at a rate 1.38 mL/min. A CDFF (29, 30) was used to grow microcosm biofilms on nano-coated and uncoated Ti samples. Briefly, the CDFF apparatus consists of a glass vessel with a top stainless steel endplate that allows for entry of nutrient medium, gas and sampling in addition to a bottom endplate designed for medium outlet. Subsequently, the inoculation flask was disconnected and the CDFF fed from a medium reservoir of sterile artificial saliva. The artificial saliva was delivered via a peristaltic pump (Watson-Marlow, Falmouth, UK) at a rate of 0.5 mL/min (0.72 L/day). The experiments were performed using two separate runs. Coated and uncoated Ti samples were positioned on PTFE plugs (located within a rotating turntable), held in place using vacuum silicone grease (Dow Corning) and recessed to a depth of 600 µm. Whole human saliva was used as the inoculum to provide microcosm biofilms. The nutrient source used for the experiments contained artificial saliva and PISF (40 µL/ min) to mimic the conditions associated with a dental implant under aerobic conditions for 5 days (37ºC).
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The rotating turntable holds 15 PTFE sampling pans rotating at 3 rpm. The rotation occurred underneath a PTFE scraper blade which allowed for sufficient medium distribution on the surface of the samples. The entire apparatus was autoclaved at 121°C for 15 min prior to use. After incubation, individual CDFF sampling pans were removed and placed into sterile universal tubes. All Ti discs were aseptically removed and placed into 1 mL of reduced transport fluid (RTF) containing five sterile glass beads (425-600 μm, Sigma-Aldrich) and then vortexed (IKA® MS 1 mini shaker - Sigma-Aldrich) for 1 min at 2200 rpm in order to disrupt the biofilm. The bacterial suspensions were serially diluted (1:10) and plated onto blood agar. Streptococcus spp. were isolated on Mitis Salivarius Agar (Sigma-Aldrich, UK); while both aerobic and anaerobic bacteria were identified using Columbia Blood Agar (Oxoid, UK) and Fastidious Anaerobe Agar (LabM, UK) respectively. All plates were incubated aerobically for 48 h with 5% CO2 at 37ºC, with the exception of fastidious anaerobic agar which were incubated for 4 to 5 days under anaerobic conditions (80% v/v N2, 10% v/v H2 and 10% v/v CO2). Finally, the colonies on the agar plates were counted and CFU/biofilm calculated. 2.6 Microscopy 2.6.1 Confocal laser scanning microscopy (CLSM) Biofilms were visualised for viable and non-viable bacteria using a LIVE/DEAD® BacLight™ kit (Life Technologies, UK). Briefly 1 µL of each of the component A and B of BacLightTM Live/Dead viability stain (Molecular Probes, Invitrogen) was added to 5 mL of reduced transport fluid. Furthermore 5 mL of this solution was carefully added to the biofilm in each well and incubated in room temperature at dark. The LIVE/DEAD BacLight Bacterial Viability Kits utilize mixtures of our SYTO® 9 green-fluorescent nucleic acid stain and the redfluorescent nucleic acid stain, propidium iodide. All prepared samples were transported into the microscope (Zeiss LSM710 Meta) and processed. 3D images were created from the green (viable bacteria) and red (non-viable 8
bacteria) colour channels using the 3D project tool of Image J and combined in order to create a single RGB (red-green-blue) stack allowing the spatial visualisation of live and dead bacteria within the biofilm structure. All images were analysed by Image J in combination with Image analysis software for investigation of microbial biofilms (31). A cell counting function (BioImage_L) was used to estimate the total bacterial biomass and the distribution of the viable and non-viable bacteria within a single image. Colours such as cream/yellow were considered to represent dying bacteria.
2.6.2 Transmission Electron Microscopy (TEM): NP characterisation NPs were characterised using TEM. ZnO and HA NPs were placed into 3mLs of deionised water and sonicated thoroughly to provide a homogenous suspension. 5 µL of each suspension was placed onto individual carbon film TEM grids (400 Mesh Cu, Agar Scientific) and air dried at room temperature. Grids were then analysed using a TEM device (Philips CM 12).
2.6.3 Scanning Electron Microscopy (SEM): Nano-coated Ti Samples All coated samples were attached to aluminum stubs, sputter coated with gold/palladium (Polaron E5000, Quorum Technology, UK) and examined using a SEM (JEOL UK Hertfordshire, UK). Samples were compared to an uncoated Ti sample (Control).
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2.7 Statistical analysis The mean values were analysed by one way-ANOVA and post hoc test for differences between controls and coating surfaces at different time points (GraphPad Prism 5). Statistical significance was established when p < 0.05.
3. Results 3.1 Characterisation of NPs using Transmission Electron Microscopy (TEM) Zinc Oxide NPs were shown to be mostly rods/hexagonal shaped with sharp angled edges and a size range of 20-50 nm in width and 20-100 nm in length (Figure 1 A and B). Similarly, nHA particles were rod-like with dimensions of 20-30 nm in width and 50-80 nm in length (Figure 1 C and D).
3.2 Characterisation of coated surfaces using Scanning Electron Microscopy (SEM) Electron micrographs demonstrate the nature of the EHDA as a coating method. ZnO and HA NPs along with their composite mixtures were uniformly coated onto the surface of Ti samples. The distribution of a fine NP layer that is homogenous can be visualised on all coated surfaces (Figure 2). 3.3 Biofilm structure analysis using CLSM Biofilms (on all surfaces and at each time point) were removed and prepared for visualisation using CLSM. All biofilms were stained with live/dead stain. At 24 h, the confocal micrographs revealed cells that were randomly distributed over the uncoated Ti samples, while on coated surfaces the cells were more likely to form chain-like structures. Biofilm communities were initially dominated by rods (24 h) and subsequently by fusiform bacteria (96 h) (Figure 3). 10
On average the biomass volume of the all surfaces was highest on the control (ca. 15,000 µm3) and lowest on the nZnO coated surface (ca. 5,000 µm3) at 24 h. The dead and dying cells were highest on the control surface (39%) and lowest on the combined nZnO + nHA surfaces (1%). Furthermore, the total biomass volume was highest on the control surface (ca. 30,000 µm3) and lowest on the HA surface (ca. 6,000 µm3) at 24 h. While at 96h, the dead and dying cells were highest on the combined nZnO + nHA surfaces (80%) and lowest on the nHA coated discs (4%). Compared with the 24 h samples, the dead or dying cells in the 96 h samples made up a greater proportion of the biofilm on the combined nZnO + nHA surfaces (80% vs. 1%) and nZnO surfaces (57% vs. 37%). The proportion of dead and dying cells decreased for controls (38% vs. 13%) and nHA coated discs (16% vs. 4%).
3.4 Biofilm analysis Biofilm samples were removed from the CDFF at 6, 24 and 96 h and processed (n =4). Total Streptococcus spp., anaerobic and aerobic colony forming units (CFUs) were quantified (Figure 4). Analysis of the pooled saliva inoculum revealed two thirds of the bacterial population were Gram-positive, half of which were Streptococcus spp. (1/3 of the total amount). Biofilms sampled at 6 h and 24 h showed a reduction of Streptococcus spp. on nZnO coated samples as compared with uncoated surfaces. Furthermore analysis of the data at 6 h indicated populations of Streptococcus spp. were reduced by 74%, 29% and 33% on nZnO , nHA and nZnO + nHA coated surfaces respectively (p < 0.05) when compared with uncoated samples. At 24 h, the CFU counts on nZnO surfaces were reduced by 96%, 76% and 89% for Streptococcus spp., anaerobic and aerobic species respectively as compared with uncoated surfaces. A significant increase in antimicrobial activity was observed at 96 h, when counts for Streptococcus spp. were reduced by 92%, 92% and 95% (reduction of 2 log fold) on nZnO, nHA and nZnO + nHA respectively (p < 0.05). Similarly the total population of anaerobic
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bacteria were reduced by 91%, 91% and 95% (reduction of 1.5 log fold) on nZnO, nHA and nZnO + nHA coated surfaces respectively (p < 0.01). Additionally the population of aerobes were reduced by 64%, 50% and 90% (reduction of 1 log fold) on nZnO, on nHA on nZnO + nHA surfaces respectively (p < 0.05). 3.5 Biofilm thickness analysis The surfaces of four Ti discs were examined by CLSM. Ten stacks of each nano-coated surface were analysed. Stacks were converted into JPEG format and processed through the image J software analysis with orthogonal views, which calculated the biofilm thickness (Figure 5). At 24 h, biofilm depth was measured at around 35 µm for uncoated Ti samples (Control), 13 µm for nHA, 14 µm for nZnO and 7 µm for combined nZnO + nHA. At 96 h biofilm depth was 41 µm for control, 35 µm for nHA, 24 µm for nZnO and 32 µm for combined nZnO + nHA surfaces. The thickest biofilm was present on control samples at 96 h (41 µm) and the thinnest biofilm was on nZnO surfaces at the same sampling time-point (24 µm). There was a significant difference between depth of biofilm and control surfaces of the nano-coated surfaces ( p < 0.05). 4. Discussion Upon insertion, a conventional Ti implant should integrate with the surrounding bone successfully. However this process may be interrupted if the surface of the implant is exposed to opportunistic pathogens with the subsequent formation of a multi-species biofilm and associated negative health-related consequences (8). The current study provides a unique insight into how such a biofilm may evolve over 4 days when in direct contact with surfaces coated with antimicrobial nanoparticles. This investigation also provided an opportunity to monitor the development of both viable and non-viable bacterial populations within a biofilm. During the initial 24 h, results indicated no significant antimicrobial activity for the coated titanium as compared to the control, with the exception of nZnO, while coatings of 50% nHA 12
+ 50% nZnO allowed 99% bacterial survival. This is in contrast to a 96h period of incubation, which demonstrated an increased antimicrobial activity for the composite coating. This ‘delayed’ antimicrobial activity was present with both the coated composites and nZnO only coated surfaces; with significant bactericidal activity against aerobes (p < 0.05), Streptococcus spp. (p < 0.01) and anaerobes (p < 0.01) compared to uncoated Ti surfaces. At 96 h the biofilm was expected to be in quasi steady state and it seems that in this case, the coatings were reducing the overall numbers of bacteria. It is possible that the coatings were inhibiting microbial growth and preventing the biofilm reaching a steady state. This was an unexpected finding as it suggests different modes of action driving the changes over the 96h period. The variation between different coatings may be attributed to surface area, which could enable bacteria to come into close proximity with the surface of nanoparticles. Furthermore, this may be attributed to the selectivity of given nanoparticles towards Gram-positive species rather than Gram negative spp. that could be directly related to the time allowed for a significant change in the composition of the biofilm. Mulligan et al, 2003 (32) have also shown that bacterial survival for silver containing phosphate glass constructs elicited antimicrobial effects using a CDFF based system for up to 96 h with a steady decline in bactericidal activity after this period. It is suggested that the nano-topography of surfaces can enhance or mitigate proliferation of bacteria. Puckett et al (33) have shown that nano-topographical changes to Ti surfaces can either enhance or decrease bacterial survival depending upon the chemical and physical properties of the surface alone, whereas other studies indicate that topography enhancement might lead to an increase in bacterial growth (34). In the current study, smooth machined titanium was used rather than the moderately roughened titanium as used in most implants to enhance bone integration. This may influence microbial adhesion and subsequent biofilm formation on the uncoated titanium surface (33). Surface topography may also significantly impact on the coating ability of NPs and any
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subsequent effects on biofilm formation (1, 21). The zinc oxide NPs used in this study were generally a mixture of square/rod shaped structures with sharp edges, whereas the HA NPs contained more rounded edges yet were similar in shape with fewer edges and generally homogenous in size. Our analysis of biofilm thickness of nano-coated surfaces indicates significant reduction for all surfaces but particularly for nZnO coated Ti surfaces. Biofilm reduction by utilisation of ZnO based coatings has been previously demonstrated in other nonoral situations (1, 35). A recent study by Eshed et al (36) demonstrated significant Streptococcus mutans biofilm reduction on ex vivo teeth coated with ZnO as compared to copper oxide (CuO) nanoparticles and uncoated teeth. A surprising outcome as CuO is often referred to as one of the most effective metal-based antimicrobial NPs (37). However, biofilms are likely to be composed of both viable and non-viable bacteria and while thickness is a useful marker of antimicrobial activity, the difference between the biofilm thickness and retrieved population of bacteria could potentially indicate that the most antimicrobial coatings can have an inverse effect on the thickness of the exo-polymers and therefore a reduction in biofilm thickness. Furthermore, experimental biofilms will contain micro-environments and concentration gradients within. For example, the concentration of nutrients and oxygen will decrease with depth into the biofilm, whilst the concentration of bacterial end products will increase. pH will also decrease with depth. These different micro-environments and gradients will affect the plaque ecosystem since the different conditions will favour or inhibit the growth of one species over another. The mechanisms by which nZnO causes bactericidal activity is currently under investigation and while the antimicrobial activity of these coatings may not fully persist over 96 h, multiple factors such as ion release, reactive oxygen species generation and direct physical contact are all thought to be responsible for these biocidal properties (38-40). Whether a single or a combination of these factors is involved in the present study is not apparent. Our findings
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indicate a highly effective coating that is capable of significant reductions in bacterial numbers for a sustained period of time in comparison to other antimicrobial coatings. Furthermore, these findings are coupled with growing evidence that nZnO based coatings are also capable of promoting osteoblast proliferation and differentiation (22, 41, 42), making them a suitable candidate for dental related implants. This is of particular importance because the current study model may represent the critical phase when infection might develop prior to full bone integration.
5. Conclusions The current study provides evidence for a novel nano-based antimicrobial implant coating which may have benefits in the prevention of implant associated infection. Furthermore, the findings show benefits in the use of composites that are comprised of multiple nanoparticles which could work in synergy to provide an antimicrobial effect against opportunistic pathogens and provide increased osseointegration. Further investigations are required in order to fully explore the effect of antimicrobial nanoparticles in the field of dentistry.
Conflict of interest: None declared
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Figure 1 – TEM analysis of nanoparticulates. A and B) micrographs indicate ZnO NPs exhibit rod-like and square-based structures with sharp edges (red arrows) that could play a role in their antimicrobial capability and action. C and D) Nano-based HA particulates are also predominantly rod-shaped and contain tiny nano-sized holes (indicated with green arrows) that allow for an increased surface area.
Figure 2 – Electron micrographs of nano-coated surfaces. A) Uncoated Ti sample, B) nZnO coated sample, C) nHA coated sample, D) Composite coatings of nHA + nZnO, E&F) Ti samples coated with nZnO have a uniform distribution throughout the sample (E), This distribution is mapped when the nZnO particulates and their aggregates are identified as white against a dark background (F).
Figure 3 – CLSM was utilised to compare viable and non-viable bacteria on different nanocoated Ti surfaces. The antimicrobial activity of the different coated surfaces is compared to uncoated samples over periods of 24 and 96 h. The percentages (%) indicate the relative populations of live (green), dead (red) and dying (yellow/cream) bacterial biomass in each specified region (Live/Dead Cell Assay). Each image represents the estimated overall effect of the coated regions on the bacterial biomass. N = 4 for each experiment at 24 & 96 h, Scale bars = 20 µm.
Figure 4 – Total colony counts of recovered bacteria from the coated surfaces (CFU/mL). The most effective antimicrobial activity can be observed at 96 h for all coated surfaces. A) Total Streptococcus spp., B) Total anaerobes and C) Total aerobes. Two independent experiments were performed for each set and four replicates of each were used. Error bars represent SD, N = 8, ** p < 0.01, * p < 0.05.
Figure 5 – Biofilm thickness (µm) comparison of coated samples at 6, 24 and 96 h. The collective biofilm thickness at each time point was measured by confocal microscopy and analysed by Image J. All samples were compared to uncoated samples as controls. Error bars represent SD, N = 10, *** p < 0.0001, * p < 0.01.
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S0300-5712(15)30057-9 http://dx.doi.org/doi:10.1016/j.jdent.2015.10.010 JJOD 2537
To appear in:
Journal of Dentistry
Received date: Revised date: Accepted date:
3-6-2015 28-9-2015 15-10-2015
Please cite this article as: Abdulkareem Elham H, Memarzadeh K, Allaker RP, Huang J, Pratten J, Spratt D.Anti-biofilm activity of zinc oxide and hydroxyapatite nanoparticles as dental implant coating materials.Journal of Dentistry http://dx.doi.org/10.1016/j.jdent.2015.10.010 This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
Anti-biofilm activity of zinc oxide and hydroxyapatite nanoparticles as dental implant coating materials
Anti-biofilm activity of zinc oxide and hydroxyapatite nanoparticles as dental implant coating materials Elham H. Abdulkareema, K. Memarzadehb,c*, R.P. Allakerb, J. Huangc, J. Prattena, D. Spratta aDepartment
of Microbial Diseases, UCL Eastman Dental Institute, University College London, UK; bInstitute of Dentistry, Barts and the
London School of Medicine and Dentistry, Queen Mary University of London, UK; cDepartment of Mechanical Engineering, University College London, UK.
Both authors contributed equally to this research
Short title: Anti-biofilm activity of ZnO/HA nanoparticles for dental implants
* Corresponding Author: Kaveh Memarzadeh (Post-doctoral researcher) Present address: Queen Mary, University of London, Barts and The London School of Medicine and Dentistry, Institute of Dentistry, 4 Newark Street, London, E1 2AT Tel: +44-02078822204
E-mails: [email protected] or [email protected]
Graphical abstract 1
Abstract Objectives: Dental implants are prone to failure as a result of bacterial biofilm accumulation. Such biofilms are often resistant to traditional antimicrobials and the use of antimicrobial nanoparticles as implant coatings may offer a means to control infection over a prolonged period. The objective of this study was to determine the antibiofilm activity of nanoparticulate coated Ti discs using a film fermenter based system.
Methods: Metal oxide nanoparticles of zinc oxide (nZnO), hydroxyapatite (nHA) and a combination (nZnO + nHA) were coated using electrohydrodynamic deposition onto titanium (Ti) discs. Using human saliva as an inoculum, biofilms were grown on coated discs for 96 h in a constant depth film fermenter under aerobic conditions with artificial saliva and periimplant sulcular fluid. Culture viability assays and biofilm thickness measurements were used to assess antimicrobial activity.
Results: Following 96 h, reduced numbers of facultatively anaerobic and Streptococcus spp on all three nano-coated surfaces were demonstrated. The proportion of non-viable microorganisms was shown to be higher on nZnO and composite (nZnO + nHA) coated surfaces at 96 h compared with nHA coated and uncoated titanium. Biofilm thickness comparison also demonstrated that nZnO and composite coatings to be the most effective.
Conclusions: The findings support the use of coating Ti dental implant surfaces with nZnO nanoparticles to provide an antimicrobial function.
Clinical Significance: Current forms of treatment for implant associated infection are often inadequate and may result in chronic infection requiring implant removal and resective / 2
regenerative procedures to restore and reshape supporting tissue. The use of metal oxide nanoparticles to coat implants could provide osteoconductive and antimicrobial functionalities to prevent failure.
Key words: Nanoparticles; Peri-implantitis; Zinc Oxide; Coatings; Biofilm; Antimicrobial.
1. Introduction Despite measures taken to prevent bacterial infection, complications with dental implants persist including those associated with antimicrobial treatment (1, 2). Upon contact, bacteria adhere, multiply and form a biofilm on the untreated surface of an implant (3). To help overcome this issue, implants can either be coated or impregnated with antimicrobial agents. However, many such approaches have been unsuccessful and resistance to those antimicrobials employed is often observed. In these cases, it is the complex structure of the associated microbial community that makes it difficult for antimicrobial agents to penetrate (4). Resistance can also involve other mechanisms (5), including microbial efflux systems that pump antimicrobial agents out of the cell (6) and the production of exopolymers which may additionally prevent penetration of agents and thus lead to the persistence of the microbial community (7). Failure or loosening of an osseointegrated implant will allow for further microbial growth, possibly resulting in peri-implantitis with associated soft and hard tissue damage (8). While the microbiota associated with oral health, including that around implants consists of predominantly Gram positive species that associated with oral infection and infected implants is predominantly Gram negative and includes Porphyromonas gingivalis, Fusobacterium spp. and Treponema denticola (9). The presence of such pathogenic bacteria in the oral cavity and other biofilms has led to the development of prophylactic approaches that
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involve the use of single or multiple antibiotics (10). However, such antibiotic treatments provide opportunities for resistance to develop when inappropriate doses are used to prevent biofilm formation (11-13). In particular, antibiotics with a single target will encourage resistance and therefore reduced efficacy of the drug. There is thus a need to investigate alternative approaches as opposed to the use of traditional antimicrobial agents (9). This may be possible by exploitation of the antimicrobial ability of metal based antimicrobial nanoparticles (NPs) to control biofilm formation. Their unique physiochemical properties should help avoid antimicrobial resistance, as activity is thought to involve multiple targets thus hindering rapid microbial adaptation and evolution (10-12). Metallic NPs possess bactericidal effects against a variety of species (13), withstand high temperatures required for fixation and possess low toxicity to mammalian cells when stabilised onto surfaces (14, 15). In order to utilise such antimicrobial capability, appropriate coating methods are required. Generally, coatings used for dental implants employ extreme conditions such as very high temperatures that can alter the properties of the coatings or surfaces (16). These conditions are often intended for stability and biological enhancement of the implant’s surface, however with respect to NPs, it is more suitable to use lower temperatures to retain beneficial physiochemical properties. Electrohydrodynamic atomisation (EHDA), a simple and economical spraying technique allows for the implementation of such conditions (17). This method produces a uniform and stable coating at room temperature whilst covering a large area and so enables sufficient functionality without compromising the original state of the coating (17). Zinc oxide NPs (nZnO) are currently being investigated for their antimicrobial activity and show much promise (18, 19). Studies suggest that the antimicrobial effects of these NPs is due to ion release or via the production of reactive oxygen species (ROS) (20). It is likely that the antimicrobial properties of NPs will differ according to their shape, surface area, size and
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relative chemical properties and thus it is these characteristics that lead to the overall synergistic activity (21). Recent studies indicate that the utilisation of nZnO also allows for the promotion of bone growth in addition to enhanced osteoblast proliferation. (22, 23). Furthermore, investigations involving HA as a coating material indicate a stable interface can be formed between bone and implant (24) . With respect to dentistry, calcium phosphates have played a key role due to their physical and chemical similarities to bone and mineralised tissues, such as enamel and dentine (25). Hence, nano-scale HA (nHA) closely resembles the size and properties of HA crystals in natural bone (26). Furthermore, studies suggest that nHA has the ability to inhibit the growth of both Gram-negative and Gram-positive bacteria, including Staphylococcus aureus and Staphylococcus epidermidis (27). The current investigation was performed using a constant depth film fermenter (CDFF) in order to improve our understanding of the antimicrobial activity of nZnO, nHA and their composite. It is hypothesized that such coating materials will be able to prevent infection as a cause of early or late dental implant failure.
2. Materials and Methods 2.1 Nanoparticles The NPs tested were: nZnO, nHA and a combination of 50% nZnO and 50% nHA. Nano-ZnO particulates were synthesised using flame pyrolysis as carried out by Johnson Matthey plc (JMTC) and nHA were synthesised as previously described (28). The surface area for nZnO was determined by Brunauer Emmett Teller (BET) analysis at JMTC.
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2.2 Ti samples Grade 23 moderate smooth machined Ti discs (Hyundai Hit 8S - A C Service Group, Fordingbridge, UK) 5 mm with a diameter and 2 mm height were used in all experiments. The Ti discs were sonicated with acetone for 20 min at 50 Hz in order to remove adherent debris or dust and then autoclaved before coating.
2.3 EHDA spraying Nano-HA, nZnO and their combination of equal proportions were used as suspensions for electrohydrodynamic spraying. This method uses a fine jet containing NPs which are deposited onto the surface of substrates. In the present study, NPs were suspended in 100% ethanol and jetted via a needle under an electric field. A stainless steel needle with a diameter of 300 μm was used to spray the content of a 1 mL syringe onto the surface of the sample. The outcome is a symmetrical cone shaped jet spray which delivers an optimal coating to ensure coverage of the sample. The distance between the substrate and the needle was maintained at 30 mm at all times. All experiments were performed with a freshly prepared suspension (10,000 µg/mL) of NPs. The voltage used was between 4.5 – 5 kV with Ti discs coated for a period of 1 min at a flow rate of 5 µL/min. All coated and uncoated samples were subjected to heat treatment (600°C) for 1 h. Samples were gradually cooled to maintain mechanical integrity of the coating (1°C/min). 2.4 Saliva samples In order to produce microcosm biofilms, saliva (unstimulated) was collected aseptically from ten members of the Department of Microbial Diseases, Eastman Dental Institute, UCL. These individuals had no significant oral disease (Ethical approval obtained from the UCL Ethics Committee; Project no. 1364/001). The pooled samples were resuspended in 10% (v/v) 6
glycerol and vortex mixed vigorously for 1 min to homogenise. Aliquots of 1 mL were then dispensed and stored at -80°C for subsequent use.
2.5 Constant depth film fermenter A vial of stored pooled saliva was thawed and 900 μL was used to inoculate 500 mL of artificial saliva. To make 1 L of artificial saliva 0.5 g 'Lab-Lemco' powder (Oxoid, UK), 1 g yeast extract (Oxoid, UK), 2.5 g protease peptone (Oxoid, UK), 1.25 g hog gastric mucin type III, partially purified from porcine stomach (Sigma Chemicals Co., Poole, UK), 0.18 g sodium chloride (BDH Chemicals Ltd, Poole, UK), 0.1 g calcium chloride (BDH) and 0.1 g potassium chloride (BDH) was added to deionised water. After autoclaving at 121°C, 0.625 ml of a 40% urea (Sigma-Aldrich) solution was added. This was mixed and pumped into the CDFF for 6 h at a rate 1.38 mL/min. A CDFF (29, 30) was used to grow microcosm biofilms on nano-coated and uncoated Ti samples. Briefly, the CDFF apparatus consists of a glass vessel with a top stainless steel endplate that allows for entry of nutrient medium, gas and sampling in addition to a bottom endplate designed for medium outlet. Subsequently, the inoculation flask was disconnected and the CDFF fed from a medium reservoir of sterile artificial saliva. The artificial saliva was delivered via a peristaltic pump (Watson-Marlow, Falmouth, UK) at a rate of 0.5 mL/min (0.72 L/day). The experiments were performed using two separate runs. Coated and uncoated Ti samples were positioned on PTFE plugs (located within a rotating turntable), held in place using vacuum silicone grease (Dow Corning) and recessed to a depth of 600 µm. Whole human saliva was used as the inoculum to provide microcosm biofilms. The nutrient source used for the experiments contained artificial saliva and PISF (40 µL/ min) to mimic the conditions associated with a dental implant under aerobic conditions for 5 days (37ºC).
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The rotating turntable holds 15 PTFE sampling pans rotating at 3 rpm. The rotation occurred underneath a PTFE scraper blade which allowed for sufficient medium distribution on the surface of the samples. The entire apparatus was autoclaved at 121°C for 15 min prior to use. After incubation, individual CDFF sampling pans were removed and placed into sterile universal tubes. All Ti discs were aseptically removed and placed into 1 mL of reduced transport fluid (RTF) containing five sterile glass beads (425-600 μm, Sigma-Aldrich) and then vortexed (IKA® MS 1 mini shaker - Sigma-Aldrich) for 1 min at 2200 rpm in order to disrupt the biofilm. The bacterial suspensions were serially diluted (1:10) and plated onto blood agar. Streptococcus spp. were isolated on Mitis Salivarius Agar (Sigma-Aldrich, UK); while both aerobic and anaerobic bacteria were identified using Columbia Blood Agar (Oxoid, UK) and Fastidious Anaerobe Agar (LabM, UK) respectively. All plates were incubated aerobically for 48 h with 5% CO2 at 37ºC, with the exception of fastidious anaerobic agar which were incubated for 4 to 5 days under anaerobic conditions (80% v/v N2, 10% v/v H2 and 10% v/v CO2). Finally, the colonies on the agar plates were counted and CFU/biofilm calculated. 2.6 Microscopy 2.6.1 Confocal laser scanning microscopy (CLSM) Biofilms were visualised for viable and non-viable bacteria using a LIVE/DEAD® BacLight™ kit (Life Technologies, UK). Briefly 1 µL of each of the component A and B of BacLightTM Live/Dead viability stain (Molecular Probes, Invitrogen) was added to 5 mL of reduced transport fluid. Furthermore 5 mL of this solution was carefully added to the biofilm in each well and incubated in room temperature at dark. The LIVE/DEAD BacLight Bacterial Viability Kits utilize mixtures of our SYTO® 9 green-fluorescent nucleic acid stain and the redfluorescent nucleic acid stain, propidium iodide. All prepared samples were transported into the microscope (Zeiss LSM710 Meta) and processed. 3D images were created from the green (viable bacteria) and red (non-viable 8
bacteria) colour channels using the 3D project tool of Image J and combined in order to create a single RGB (red-green-blue) stack allowing the spatial visualisation of live and dead bacteria within the biofilm structure. All images were analysed by Image J in combination with Image analysis software for investigation of microbial biofilms (31). A cell counting function (BioImage_L) was used to estimate the total bacterial biomass and the distribution of the viable and non-viable bacteria within a single image. Colours such as cream/yellow were considered to represent dying bacteria.
2.6.2 Transmission Electron Microscopy (TEM): NP characterisation NPs were characterised using TEM. ZnO and HA NPs were placed into 3mLs of deionised water and sonicated thoroughly to provide a homogenous suspension. 5 µL of each suspension was placed onto individual carbon film TEM grids (400 Mesh Cu, Agar Scientific) and air dried at room temperature. Grids were then analysed using a TEM device (Philips CM 12).
2.6.3 Scanning Electron Microscopy (SEM): Nano-coated Ti Samples All coated samples were attached to aluminum stubs, sputter coated with gold/palladium (Polaron E5000, Quorum Technology, UK) and examined using a SEM (JEOL UK Hertfordshire, UK). Samples were compared to an uncoated Ti sample (Control).
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2.7 Statistical analysis The mean values were analysed by one way-ANOVA and post hoc test for differences between controls and coating surfaces at different time points (GraphPad Prism 5). Statistical significance was established when p < 0.05.
3. Results 3.1 Characterisation of NPs using Transmission Electron Microscopy (TEM) Zinc Oxide NPs were shown to be mostly rods/hexagonal shaped with sharp angled edges and a size range of 20-50 nm in width and 20-100 nm in length (Figure 1 A and B). Similarly, nHA particles were rod-like with dimensions of 20-30 nm in width and 50-80 nm in length (Figure 1 C and D).
3.2 Characterisation of coated surfaces using Scanning Electron Microscopy (SEM) Electron micrographs demonstrate the nature of the EHDA as a coating method. ZnO and HA NPs along with their composite mixtures were uniformly coated onto the surface of Ti samples. The distribution of a fine NP layer that is homogenous can be visualised on all coated surfaces (Figure 2). 3.3 Biofilm structure analysis using CLSM Biofilms (on all surfaces and at each time point) were removed and prepared for visualisation using CLSM. All biofilms were stained with live/dead stain. At 24 h, the confocal micrographs revealed cells that were randomly distributed over the uncoated Ti samples, while on coated surfaces the cells were more likely to form chain-like structures. Biofilm communities were initially dominated by rods (24 h) and subsequently by fusiform bacteria (96 h) (Figure 3). 10
On average the biomass volume of the all surfaces was highest on the control (ca. 15,000 µm3) and lowest on the nZnO coated surface (ca. 5,000 µm3) at 24 h. The dead and dying cells were highest on the control surface (39%) and lowest on the combined nZnO + nHA surfaces (1%). Furthermore, the total biomass volume was highest on the control surface (ca. 30,000 µm3) and lowest on the HA surface (ca. 6,000 µm3) at 24 h. While at 96h, the dead and dying cells were highest on the combined nZnO + nHA surfaces (80%) and lowest on the nHA coated discs (4%). Compared with the 24 h samples, the dead or dying cells in the 96 h samples made up a greater proportion of the biofilm on the combined nZnO + nHA surfaces (80% vs. 1%) and nZnO surfaces (57% vs. 37%). The proportion of dead and dying cells decreased for controls (38% vs. 13%) and nHA coated discs (16% vs. 4%).
3.4 Biofilm analysis Biofilm samples were removed from the CDFF at 6, 24 and 96 h and processed (n =4). Total Streptococcus spp., anaerobic and aerobic colony forming units (CFUs) were quantified (Figure 4). Analysis of the pooled saliva inoculum revealed two thirds of the bacterial population were Gram-positive, half of which were Streptococcus spp. (1/3 of the total amount). Biofilms sampled at 6 h and 24 h showed a reduction of Streptococcus spp. on nZnO coated samples as compared with uncoated surfaces. Furthermore analysis of the data at 6 h indicated populations of Streptococcus spp. were reduced by 74%, 29% and 33% on nZnO , nHA and nZnO + nHA coated surfaces respectively (p < 0.05) when compared with uncoated samples. At 24 h, the CFU counts on nZnO surfaces were reduced by 96%, 76% and 89% for Streptococcus spp., anaerobic and aerobic species respectively as compared with uncoated surfaces. A significant increase in antimicrobial activity was observed at 96 h, when counts for Streptococcus spp. were reduced by 92%, 92% and 95% (reduction of 2 log fold) on nZnO, nHA and nZnO + nHA respectively (p < 0.05). Similarly the total population of anaerobic
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bacteria were reduced by 91%, 91% and 95% (reduction of 1.5 log fold) on nZnO, nHA and nZnO + nHA coated surfaces respectively (p < 0.01). Additionally the population of aerobes were reduced by 64%, 50% and 90% (reduction of 1 log fold) on nZnO, on nHA on nZnO + nHA surfaces respectively (p < 0.05). 3.5 Biofilm thickness analysis The surfaces of four Ti discs were examined by CLSM. Ten stacks of each nano-coated surface were analysed. Stacks were converted into JPEG format and processed through the image J software analysis with orthogonal views, which calculated the biofilm thickness (Figure 5). At 24 h, biofilm depth was measured at around 35 µm for uncoated Ti samples (Control), 13 µm for nHA, 14 µm for nZnO and 7 µm for combined nZnO + nHA. At 96 h biofilm depth was 41 µm for control, 35 µm for nHA, 24 µm for nZnO and 32 µm for combined nZnO + nHA surfaces. The thickest biofilm was present on control samples at 96 h (41 µm) and the thinnest biofilm was on nZnO surfaces at the same sampling time-point (24 µm). There was a significant difference between depth of biofilm and control surfaces of the nano-coated surfaces ( p < 0.05). 4. Discussion Upon insertion, a conventional Ti implant should integrate with the surrounding bone successfully. However this process may be interrupted if the surface of the implant is exposed to opportunistic pathogens with the subsequent formation of a multi-species biofilm and associated negative health-related consequences (8). The current study provides a unique insight into how such a biofilm may evolve over 4 days when in direct contact with surfaces coated with antimicrobial nanoparticles. This investigation also provided an opportunity to monitor the development of both viable and non-viable bacterial populations within a biofilm. During the initial 24 h, results indicated no significant antimicrobial activity for the coated titanium as compared to the control, with the exception of nZnO, while coatings of 50% nHA 12
+ 50% nZnO allowed 99% bacterial survival. This is in contrast to a 96h period of incubation, which demonstrated an increased antimicrobial activity for the composite coating. This ‘delayed’ antimicrobial activity was present with both the coated composites and nZnO only coated surfaces; with significant bactericidal activity against aerobes (p < 0.05), Streptococcus spp. (p < 0.01) and anaerobes (p < 0.01) compared to uncoated Ti surfaces. At 96 h the biofilm was expected to be in quasi steady state and it seems that in this case, the coatings were reducing the overall numbers of bacteria. It is possible that the coatings were inhibiting microbial growth and preventing the biofilm reaching a steady state. This was an unexpected finding as it suggests different modes of action driving the changes over the 96h period. The variation between different coatings may be attributed to surface area, which could enable bacteria to come into close proximity with the surface of nanoparticles. Furthermore, this may be attributed to the selectivity of given nanoparticles towards Gram-positive species rather than Gram negative spp. that could be directly related to the time allowed for a significant change in the composition of the biofilm. Mulligan et al, 2003 (32) have also shown that bacterial survival for silver containing phosphate glass constructs elicited antimicrobial effects using a CDFF based system for up to 96 h with a steady decline in bactericidal activity after this period. It is suggested that the nano-topography of surfaces can enhance or mitigate proliferation of bacteria. Puckett et al (33) have shown that nano-topographical changes to Ti surfaces can either enhance or decrease bacterial survival depending upon the chemical and physical properties of the surface alone, whereas other studies indicate that topography enhancement might lead to an increase in bacterial growth (34). In the current study, smooth machined titanium was used rather than the moderately roughened titanium as used in most implants to enhance bone integration. This may influence microbial adhesion and subsequent biofilm formation on the uncoated titanium surface (33). Surface topography may also significantly impact on the coating ability of NPs and any
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subsequent effects on biofilm formation (1, 21). The zinc oxide NPs used in this study were generally a mixture of square/rod shaped structures with sharp edges, whereas the HA NPs contained more rounded edges yet were similar in shape with fewer edges and generally homogenous in size. Our analysis of biofilm thickness of nano-coated surfaces indicates significant reduction for all surfaces but particularly for nZnO coated Ti surfaces. Biofilm reduction by utilisation of ZnO based coatings has been previously demonstrated in other nonoral situations (1, 35). A recent study by Eshed et al (36) demonstrated significant Streptococcus mutans biofilm reduction on ex vivo teeth coated with ZnO as compared to copper oxide (CuO) nanoparticles and uncoated teeth. A surprising outcome as CuO is often referred to as one of the most effective metal-based antimicrobial NPs (37). However, biofilms are likely to be composed of both viable and non-viable bacteria and while thickness is a useful marker of antimicrobial activity, the difference between the biofilm thickness and retrieved population of bacteria could potentially indicate that the most antimicrobial coatings can have an inverse effect on the thickness of the exo-polymers and therefore a reduction in biofilm thickness. Furthermore, experimental biofilms will contain micro-environments and concentration gradients within. For example, the concentration of nutrients and oxygen will decrease with depth into the biofilm, whilst the concentration of bacterial end products will increase. pH will also decrease with depth. These different micro-environments and gradients will affect the plaque ecosystem since the different conditions will favour or inhibit the growth of one species over another. The mechanisms by which nZnO causes bactericidal activity is currently under investigation and while the antimicrobial activity of these coatings may not fully persist over 96 h, multiple factors such as ion release, reactive oxygen species generation and direct physical contact are all thought to be responsible for these biocidal properties (38-40). Whether a single or a combination of these factors is involved in the present study is not apparent. Our findings
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indicate a highly effective coating that is capable of significant reductions in bacterial numbers for a sustained period of time in comparison to other antimicrobial coatings. Furthermore, these findings are coupled with growing evidence that nZnO based coatings are also capable of promoting osteoblast proliferation and differentiation (22, 41, 42), making them a suitable candidate for dental related implants. This is of particular importance because the current study model may represent the critical phase when infection might develop prior to full bone integration.
5. Conclusions The current study provides evidence for a novel nano-based antimicrobial implant coating which may have benefits in the prevention of implant associated infection. Furthermore, the findings show benefits in the use of composites that are comprised of multiple nanoparticles which could work in synergy to provide an antimicrobial effect against opportunistic pathogens and provide increased osseointegration. Further investigations are required in order to fully explore the effect of antimicrobial nanoparticles in the field of dentistry.
Conflict of interest: None declared
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Figure 1 – TEM analysis of nanoparticulates. A and B) micrographs indicate ZnO NPs exhibit rod-like and square-based structures with sharp edges (red arrows) that could play a role in their antimicrobial capability and action. C and D) Nano-based HA particulates are also predominantly rod-shaped and contain tiny nano-sized holes (indicated with green arrows) that allow for an increased surface area.
Figure 2 – Electron micrographs of nano-coated surfaces. A) Uncoated Ti sample, B) nZnO coated sample, C) nHA coated sample, D) Composite coatings of nHA + nZnO, E&F) Ti samples coated with nZnO have a uniform distribution throughout the sample (E), This distribution is mapped when the nZnO particulates and their aggregates are identified as white against a dark background (F).
Figure 3 – CLSM was utilised to compare viable and non-viable bacteria on different nanocoated Ti surfaces. The antimicrobial activity of the different coated surfaces is compared to uncoated samples over periods of 24 and 96 h. The percentages (%) indicate the relative populations of live (green), dead (red) and dying (yellow/cream) bacterial biomass in each specified region (Live/Dead Cell Assay). Each image represents the estimated overall effect of the coated regions on the bacterial biomass. N = 4 for each experiment at 24 & 96 h, Scale bars = 20 µm.
Figure 4 – Total colony counts of recovered bacteria from the coated surfaces (CFU/mL). The most effective antimicrobial activity can be observed at 96 h for all coated surfaces. A) Total Streptococcus spp., B) Total anaerobes and C) Total aerobes. Two independent experiments were performed for each set and four replicates of each were used. Error bars represent SD, N = 8, ** p < 0.01, * p < 0.05.
Figure 5 – Biofilm thickness (µm) comparison of coated samples at 6, 24 and 96 h. The collective biofilm thickness at each time point was measured by confocal microscopy and analysed by Image J. All samples were compared to uncoated samples as controls. Error bars represent SD, N = 10, *** p < 0.0001, * p < 0.01.
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