Patent Application
20150079174
The present invention relates to antimicrobial complexes and methods of their preparation, and, in particular to an antimicrobial complex comprising a nanomaterial and a surfactant. Antimicrobial-nanomaterial complexes and suspensions thereof can be used to form antimicrobial coatings with particular utility in industrial settings.
The development of antimicrobial coatings has received increasing attention in recent years, due to the need to prevent propagation of microbes on surfaces. The preparation of antimicrobial coatings for industrial purposes represents a particular challenge, due to the broad spectrum of antimicrobial activity required, the varying dimensions of the surfaces to be coated, as well as the different materials to which the surface coatings must be applied. In order to prove applicable to industrial settings, therefore, antimicrobial coatings should preferably be versatile, easy and relatively inexpensive to produce, and exhibit long-lasting antimicrobial characteristics.
Known antimicrobial coatings can be prepared by impregnating, adsorbing or covalently attaching antimicrobial agents to the relevant surface in order to provide a coating layer. Research has previously focused on the development of non-leaching surfaces capable of killing microorganisms on contact (Klibanov, J. Mater. Chem. 2007 (17) 2479-2482; Ferreira et al., J. Mater. Chem. 2009 (19) 7796-7806). Non-leaching surfaces are often considered preferable, because microbes are exposed to high surface concentrations of the antimicrobial agent compared with slow release surfaces. Known non-leaching antimicrobial coatings and preparation methodologies suffer from a number of drawbacks, however.
Methods for the permanent immobilization of synthetic polymers to the surfaces of materials are known. For example, the covalent attachment of polymers via surface-initiated polymerization and the use of coupling-reactions to immobilize polymer chains to functional groups on surfaces to be treated have been described in the art.
However, these methods involve multi-step procedures in the preparation of the antimicrobial surface, making large-scale production and commercialisation of the coatings impractical. Furthermore, the attachment of the polymers requires specific chemistry, meaning that tailored methodologies and/or surface modification are required with regards to the surface of each object to be coated.
Alternative approaches to the preparation of antimicrobial coatings include the use of coatings that are non-covalently linked to the surface, for example as described by Klibanov et al (J. Mat. Chem.' Permanently microbicidal materials coatings' 2007 (17) 2479-2482). However, in many cases, the methodologies described require multiple synthetic steps and are difficult to use in objects with different chemistries.
Therefore, despite active research in this area, there remains a need for novel antimicrobial complexes which exhibit broad spectrum antimicrobial activity and which can be readily immobilized to surfaces of a broad range of materials, without the requirement for specific surface chemistries. In addition, it would be desirable to provide antimicrobial complexes or coatings which expose microbes to high surface concentrations of the antimicrobial agent, and which exhibit sufficient life-span to make them viable for use in industrial settings. It would also be useful to have a versatile and inexpensive process for preparing surface coatings on a commercial scale.
The applicant has now discovered that surfactants can be complexed with nanomaterials, to form antimicrobial nanomaterial complexes which exhibit effective antimicrobial activity and which address some of the disadvantages of the prior art materials.
Accordingly, in an aspect of the invention there is provided an antimicrobial nanomaterial complex comprising a surfactant and a nanomaterial.
In the antimicrobial nanomaterial complex, the surfactant functions as an antimicrobial agent.
Although some surfactants are known to possess antibacterial properties, the activity of these surfactants is relatively low, making them generally unsuitable for use as antimicrobial agents due to the significantly high loading that would be required in order to obtain a useful surface-coating material. In addition, although mild antibacterial activity has been demonstrated, surfactants are not known to exhibit broad spectrum antimicrobial activity.
However, the applicant has surprisingly discovered that when an antimicrobial surfactant is complexed with a nanomaterial, significantly lower minimal inhibitory concentration is exhibited compared with the soluble surfactant alone. The lower minimal inhibitory concentration of antimicrobial surfactants in the nanomaterial complexes mean that surfactants can be effectively employed in the formation of antimicrobial preparations and coatings.
The “minimal inhibitory concentration” or “MIC” is defined as the lowest drug concentration that results in complete inhibition of visible growth of a microorganism.
The term “antimicrobial agent” as used herein is taken to mean a substance that kills (microbicidal) or inhibits (microbistatic) the growth of microorganisms. The term “antimicrobial agent” thus encompasses antibacterials, antivirals, antifungals and antiparasitics.
The antimicrobial nanomaterial complex may be a broad-spectrum antimicrobial complex and may have antibacterial, antiviral and antifungal activity.
In an embodiment of the invention, the antimicrobial nanomaterial complex is antiviral.
In the antimicrobial nanomaterial complex, the surfactant may be attached to the nanomaterial by any conventional means of attachment that allows the surfactant to be presented at the surface of the nanomaterial. For example, the surfactant may be adsorbed or immobilized on the surface of the nanomaterial. The surfactant may be coated on the surface of the nanomaterial and may interact electrostatically with the nanomaterial surface. Advantageously, the surfactant is attached directly to the nanomaterial in the complex or conjugate and is not attached to the nanomaterial by means of a linker molecule, such as a polymer.
The surfactant may be an amphiphilic polycation. The surfactant may be single-chained or multi-chained. For example the surfactant may be single-chained, di-chained or tri-chained. The chains may be of variable chain length. For example, the surfactant may include at least one hydrocarbon chain comprising between 12 and 20 carbon atoms. In particular, the surfactant may include at least one hydrocarbon chain comprising 12, 14, 16 or 18 carbon atoms.
In an embodiment of the invention, the amphiphilic polycation is a quaternary ammonium cationic surfactant. The quaternary ammonium cationic surfactant may be selected from the group consisting of dodecyltrimethylammonium bromide (DTAB), didodecyldimethylammonium bromide (DDAB), tridodecylmethylammonium chloride (TMAC), ditetradecyldimethylammonium bromide (DTDAB), dihexadecyldimethylammonium bromide (DHDAB) and dioctadecyldimethylammonium bromide (DODAB).
The term “nanomaterial” as used herein is taken to mean materials comprising particles having one or more dimensions at the nanometre scale and preferably those particles having one or more dimensions of 1 to 500 nm. Thus the term “nanomaterial” encompasses materials having both ultrafine particles (1 to 100 nm) and fine particles from 100 to 500 nm.
The nanomaterial may be a nanoparticle (NP). Suitable nanoparticles for use in the complexes of the invention include, but are not limited to gold nanoparticles, silver nanoparticles, silica nanoparticles and organic nanoparticles such as polylactic acid-co-glycolic acid), polyethyleneimine, chytosan and poly(hyaluronic acid).
In an embodiment of the invention, the nanoparticle is a silica nanoparticle (SNP). Silica nanoparticles are particularly advantageous for the formation of stable antimicrobial complexes due to the non-cytotoxicity, low price, high stability, durability, and ease of modification by organosilane chemistry allowing the incorporation of an array of different functional groups.
Nanoparticles for immobilization in the complex can be of any suitable diameter. In particular, the average diameter of the nanoparticles can be from 2 to 500 nm; from 10 to 400 nm; from 20 to 290 nm. In an embodiment, the average diameter of the nanoparticle can be 100 nm or below, or from 20 to 100 nm.
The nanomaterial may be a carbon nanotube (CNT). Carbon nanotubes are promising for the formation of antimicrobial complexes because they are known to possess inherent antimicrobial properties (Kang et al. ‘Single-walled carbon nanotubes exhibit strong antimicrobial activity’ Langmuir 2007 (23) 8670-8673).
Carbon nanotubes for immobilization in the complex may be of any suitable diameter and length. In particular, the average diameter of the nanotubes can be from 0.8 to 500 nm. The average diameter may be about 1.5 nm. The average length can be from 50 nm to 10 μm.
The carbon nanotube may be a single-walled (SWCNT) or multi-walled nanotube (MWCNT). In some embodiments single-walled nanotubes are preferred as they are known to exhibit mild antimicrobial activity. In other embodiments, multi-walled carbon nanotubes, which are less expensive, may be preferred.
In an aspect of the invention there is provided a process of preparing an antimicrobial complex as described above comprising the steps of:
The “critical micelle concentration” or “CMC” is defined as the concentration of surfactant at which phase separation occurs.
Typically, a solution of surfactant is mixed with a suspension of the nanomaterial and allowed to complex. After complexation, the mixture is centrifuged to wash away unconjugated surfactant. The method of the invention therefore represents an easy and low-cost route to the preparation of antimicrobial complexes.
Use of a surfactant in the preparation of an antimicrobial complex is envisaged. In an embodiment, an amphiphilic polycation, and preferably a quaternary ammonium cationic surfactant, can be used in the preparation of an antimicrobial complex. In this embodiment, the nanomaterial can be a silica nanoparticle or a carbon nanotube.
In an aspect of the invention there is provided use of an antimicrobial complex as described above as an antimicrobial coating.
The complex may be utilised in the form of a suspension of the nanomaterial complex in any suitable solution.
According to an aspect of the invention there is provided a method of preparing an object with an antimicrobial coating, wherein the method comprises coating at least one surface or part of a surface of the object with an antimicrobial complex as described herein.
The at least one surface or part of a surface of the device or material can be coated by any suitable means known in the art such as, for example, spray-drying, brushing or dipping.
In an embodiment of the invention, the at least one surface or part of a surface is treated with an adhesive prior to coating. Treating the surface with an adhesive prior to coating can ensure efficient immobilization of the antimicrobial nanoparticle complex. The adhesive may be a chemical adhesive.
In an embodiment, the adhesive is formed by a polyelectrolyte of opposite charge to the antimicrobial nanomaterial complex which is adsorbed or chemically reacted with the substrate. In this case, assembly of the nanoparticles into large-area uniform thin films can be achieved by layer-by-layer techniques such as described in Lee D et al.′ Multilayers of oppositely charged SiO2 nanoparticles: Effect of surface charge on multilayer assembly′, Langmuir 2007, 23, 8833-8837.
In an alternative embodiment, the adhesive is formed by nanoparticles of opposite charge to the antimicrobial nanomaterial conjugate. These nanoparticles can interact with the substrate by electrostatic interactions and physically immobilize the antimicrobial nanomaterial complexes of opposite charge.
In an embodiment, the adhesive is a chemical adhesive formed by polydopamine. Dopamine, a biomolecule that contains catechol and amine functional groups, found also in high concentration in mussel adhesive proteins, polymerizes at alkaline pHs to form thin adherent polydopamine films that exhibit latent reactivity toward amine and thiol groups (Lee H et al., ‘Facile conjugation of biomolecules onto surfaces via mussel adhesive protein inspired coatings’: Advanced Materials 2009, 21, 431).
The method of the invention can be used to apply an antimicrobial coating to any suitable surface such as, for example, metals ceramics, polymers, fibres and glass. Suitable metals include, for example, titanium and titanium alloys such as nitinol, nickel-titanium alloys and thermo-memory alloy materials; stainless steel; tantalum; nickel-chrome alloys and cobalt alloys such as cobalt-chromium alloys Elgiloy® and Phynox®. Suitable ceramic materials include, for example, oxides, carbides or nitrides of the transition elements such as titanium oxides, hafnium oxides, iridium oxides, chromium oxides, aluminium oxides and zirconium oxides. Suitable polymers include styrene and substituted styrenes, ethylene, propylene, poly(urethanes), acrylates, methacrylates, acrylamides, methacrylamides, polyesters, polysiloxanes, polyethers, poly(orthoester), poly(carbonates), poly(hydroxylalkanoates), and copolymers thereof. Attachment of the antimicrobial nanomaterial complex to a surface using an adhesive lends the method of the invention versatility as specific surface chemistries are not required. The method therefore has broad utility, and is particularly useful in industrial settings where a wide variety of materials and objects require antimicrobial protection.
Surfaces which may be coated can be in the form of films, particles, beads and fibres.
In an aspect of the invention there is provided an object with an antimicrobial coating, obtainable by the method as described herein.
In an embodiment of the invention, the antimicrobial nanomaterial complex is a surfactant-SNP complex. In an embodiment, the antimicrobial nanomaterial complex is a quaternary ammonium cationic surfactant-SNP complex. In this embodiment, the quaternary ammonium cationic surfactant preferably has two carbon chains. In a preferred embodiment, the antimicrobial nanomaterial complex is a DDAB-SNP complex.
In an embodiment of the invention, the antimicrobial nanomaterial is a surfactant-CNT complex. In an embodiment, the antimicrobial nanomaterial complex is a quaternary ammonium cationic surfactant-CNT complex. In this embodiment, the quaternary ammonium cationic surfactant preferably has two carbon chains. In a preferred embodiment, the antimicrobial nanomaterial complex is a DDAB-CNT complex. In an embodiment, the nanomaterial is a SWCNT. Alternatively the nanomaterial may be a MWCNT. In an embodiment, the antimicrobial nanomaterial complex is a DDAB-SWCNT complex.
Advantageously, the inventors have also discovered that the complexed nanomaterials can be encapsulated in order to control their spatial location in surfaces. This can be beneficial, for example, to minimise the cost of expensive nanomaterials such as carbon nanotubes while maintaining the surfactant at the surface of a coating material. As an example, the nanomaterial complexes can be encapsulated in a bead or a carrier. Specifically, the nanomaterial complexes can be encapsulated in alginate beads. In order to ensure presentation of the nanomaterial complex at the surface of the object to be coated, magnetic nanoparticles can be incorporated in the encapsulated product to direct the spatial location of the complex by use of a magnet.
The invention is now further described with reference to the following figures:
The invention is directed to antimicrobial complexes comprising nanomaterials complexed with surfactants. By way of illustration only, the invention will now be fully described in the context of a didodecyldimethylammonium bromide-silica nanoparticle complex (DDAB-SNP) and a didodecyldimethylammonium bromide-carbon nanotube complex (DDAB-CNT).
Commercially available silica nanoparticles of 5 nm (SNP5) and 80 nm (SNP80) in diameter were purchased from Eka Chemicals AB (Bohus, Sweden) and PlasmaChem GmbH (Berlin, Germany), respectively. The nanoparticles were characterised prior to conjugation by scanning electron microscopy, transmission electron microscopy and dynamic light scattering as detailed below. The specific surface area and the density of the particles were also determined.
1.2.1 Scanning Electron Microscopy (SEM) Analysis
SNP5 suspended in distilled water (5 μl, 0.5 mg/ml) were deposited on 0.5 cm2 glass slides. The solvent was allowed to evaporate and the slides mounted on a SEM sample stub using conductive carbon cement. The samples were then carbon coated by plasma vapour deposition and analyzed by a Hitachi SU-70, with a STEM detector at 4 kV. Results of the SEM analysis are shown in
1.2.2 Transmission Electron Microscopy (TEM) Analysis
For TEM analysis, a suspension of SNP80 (10 μl, 5 mg/ml in Phosphate Buffered Saline (PBS)) was spray coated on a TEM 400 mesh grid. The SNPs were then observed by TEM on a FEI microscope (model TECNAI G2 20 S-TWIN) at 200 kV. Results of the TEM analysis are shown in
1.2.3 Dynamic Light Scattering (DLS) Analysis
SNP size was determined using dynamic light scattering (DLS) via Zeta PALS Zeta Potential Analyzer and ZetaPlus Particle Sizing Software, v. 2.27 (Brookhaven Instruments Corporation). An aliquot (20 μl) of SNPs suspended in water (2 mg/ml) was added to 2 ml of PBS pH 7.4, vortexed and sonicated. All sizing measurements were performed at ca. 25° C., and all data were recorded at 90° C., with an equilibration time of 5 min. and individual run times of 60 s (5 runs per measurement). The average diameters are number-weighted, and were collected from 3 independent measurements. The zeta potential of the previous SNP suspensions was recorded in at least 6 runs with a relative residual value (measure of data fit quality) of 0.03.
1.2.4 Specific Surface Area Measurement
The specific surface area was assessed on an ASAP 2000 instrument, using the BET isotherm model with a relative pressure range of 0.05-0.15.
1.2.5 Density Measurement
Density of the nanoparticles was measured using helium pycnometry.
The results of the SNP characterisation are illustrated in Table 1 below.
SNP5 were found by DLS to have a diameter of 7.5±1.7 nm, corresponding to a diameter of 5.0±2.0 by SEM. SNP80 were found by DLS to have a diameter of 78.9±12.6 nm, corresponding to a diameter of 58.3±3.8 by TEM. BET analysis established a surface area of 293.0±3.4 m2g−1 for SNP5 and a surface area of 218.5±68 m2g−1 for SNP80. The densities of the SNPs were 2.18 g/cm3 (SNP5) and 1.20 g/cm3 (SNP80) measured by helium pycnometry.
Critical Micelle Concentration (CMC) was determined for a range of surfactants, following the method described by Brito et al. ‘Determination of the critical micelle concentration of surfactants using the fluorescent probe N-phenyl-1-naphthylamine.’ Anal. Biochem 1986 (152) 250-255.
The surfactants tested were:
dodecyltrimethylammonium bromide (DTAB, TCI), didodecyldimethylammonium bromide (DDAB, Sigma-Aldrich),
tridodecylmethylammonium chloride (TMAC, Sigma-Aldrich),
ditetradecyldimethylammonium bromide (DTDAB, TCI),
dihexadecyldimethylammonium bromide (DHDAB, TCI) and dioctadecyldimethylammonium bromide (DODAB, Sigma-Aldrich).
Briefly, the surfactants were dissolved in PBS (pH 7.4), serially diluted and mixed with N-phenyl-1-naphthylamine (NPN; TCI). The fluorescence of each sample was quantified (excitation at 350 nm; emission at 420 nm) and plotted against the concentration of each surfactant. The CMC was found at the intersection of two straight lines traced on the emission intensity plot, defining the aqueous and micellar environments. The CMC values for the tested surfactants are given in Table 2 below.
To evaluate the effect of the molecular structure of quaternary ammonium cationic surfactants on their antimicrobial activity, a range of surfactants were tested against fungi and bacteria in order to determine the minimal inhibitory concentration. The surfactants tested were single-chained (DTAB), di-chained (DDAB) and tri-chained (TMAC), having 12-C length hydrocarbon chains. In addition, to evaluate the effect of chain length on antimicrobial activity, dialkyldimethylammonium bromide surfactants with variable chain lengths including 12 (DDAB), 14 (DTAB), 16 (DHDAB) and 18 (DODAB) carbon atoms were tested. These surfactants are shown in
Growth kinetics of Candida albicans ATCC 10231 (C. albicans, fungi), Staphylococcus aureus ATCC 6538 (S. aureus, gram-positive bacteria) and Escherichia coli ATCC 25922 (E. coli, gram-negative bacteria) (1×105 cells/ml) exposed to variable concentrations (from 62.5 to 500 μg/ml for fungi and from 15.6 to 500 μg/ml for S. aureus and E. coli) of the surfactants was evaluated by absorbance at 600 nm. Results are shown in
Results are shown in Table 2.
C.
albicans
S. aureus
E. coli
With regard to E. Coli, the toxicity ranking of all surfactants studied in this work was DTAB>DDAB>DHDAB>TMAC=DTDAB; for C. albicans was DDAB>DHDAB>DTAB>TMAC>DTAB>DODAB, and finally for S. aureus was DDAB>DTAB>DHDAB>DODAB≈DTDAB>TMAC. DDAB is the most effective in the inhibition of C. albicans (125 μg/ml) and S. aureus (31.3 μg/ml), while DTAB is the most effective in the inhibition of E. coli (125 μg/ml). Results suggested that antimicrobial activity is higher for surfactants having two hydrocarbon chains formed by 12 carbons than for those with C-14, C-16 or C-18 chains.
Based on the results of example 3 above, DDAB was chosen for further investigation. SNPs were mixed with aqueous solutions of DDAB at different concentrations and pHs, for varying lengths of time to facilitate absorption. Concentrations of DDAB above its CMC (32.4 μg/ml, example 2 above) were tested (i.e. 200, 500, 1000 and 2000 μg/ml for DDAB50-SNP, DDAB125-SNP, DDAB250-SNP and DDAB500-SNP, respectively). The concentrations used were 50, 125, 250 and 500 μg of DDAB per mg of SNPs (suspension of 5%, w/v) and therefore per 0.293 m2 or 0.218 m2 in case of SNP5 and SNP80, respectively (taking into account the surface areas of 293.0±3.4 m2g−1 and 218.5±68 m2g1 for SNP5 and SNP80, respectively).
Specifically, aqueous suspensions of SNP5 or SNP80 (5% w/v, in (i) distilled water; (ii) 0.1M citrate/sodium citrate buffer pH 3.0 and (iii) 0.1M borate/NaOH buffer pH 9.0) were added to an aqueous solution of DDAB (50, 125, 250 and 500 μg per mg of SNPs), for variable time (30 min, 1 h, or 3 h), under vigorous magnetic stirring (800 rpm) at room temperature.
After the adsorption period, the suspension of the SNPs was centrifuged (30 min, 40,000 rpm for SNP5; 20 min, 10,000 rpm for SNP80), washed two times with distilled water and finally freeze-dried. Importantly, aqueous solutions of DDAB (i.e. without SNPs, forming vesicles with an average diameter of 57 nm and a zeta potential of 17.11±2.15 mV) centrifuged at the same velocities, did not form a pellet.
The adsorption of DDAB to SNPs was determined by conventional and high resolution modulated Thermogravimetric Analysis (TGA).
The thermal behavior of the samples was evaluated by simultaneous thermal analysis (STA; heat-flux DSC and TGA), by using a TA Instruments SDT Q600 equipment (thermobalance sensitivity: 0.1 μg), which was previously calibrated in the range 25-1000° C. by running tin and lead as melting standards, at a heating rate of 10° C. min−1, using open alumina crucibles and a dry nitrogen purge flow of 100 ml min−1. The mass loss measured in the temperature range from 200 to 350° C. was used to calculate the percentage of surfactant adsorbed to a certain mass of SNPs.
The mass loss process was further studied in detail by high-resolution modulated thermogravimetric analysis (HiRes-M-TGA) in a TA Instruments Q500 thermogravimetric apparatus (thermobalance sensitivity: 0.1 μg). The temperature calibration was performed in the range 25-1000° C. by measuring the Curie point of nickel standard. Open platinum crucibles and a dry nitrogen purge flow of 100 ml min−1 were used. The experiments were performed under a dynamic rate mode with a (maximum) heating rate of 2° C. min−1, a modulation period of 200 s, and a temperature amplitude of ±5° C.
The results of the thermogravimetric analysis are shown in
Depending on the initial concentration of DDAB, SNP5 was found to immobilize between 57 and 210 μg of DDAB per mg of SNP, while SNP80 was found to immobilize between 45 to 275 μg of the conjugate per mg of SNP (
The derivatives of the TGA plots in DDAB-SNP5 indicate the presence of two major peaks: the first one between 150° C. and 195° C., and the second one between 218° C. and 255° C. (
The influence of the concentration, pH and time on the physical immobilization of DDAB to SNPs was also assessed and the results are shown in
According to
The range of times (30 min, 1 hr, 3 hr) (
To further characterize the adsorption of DDAB to SNPs, Diffuse Reflectance Infrared Fourier Transform Spectroscopy (DRIFT) analysis was carried out. DRIFT analysis of lyophilized DDAB vesicles and of the bare and DDAB-coated SNPs was performed using a Mattson RS1 FTIR spectrometer with a Specac Selector, in the 400-4000 cm−1 range (wide band MCT detector), at 4 cm−1 resolution. The spectra were the result of 500 co-added scans for each sample, ratioed against the same number of scans for the background (ground KBr, FTIR grade from Aldrich). The samples were previously ground and mixed with KBr in appropriate proportions to obtain spectral absorbance in the range of applicability of the Kubelka-Munk transformation (Kubelka et al. ‘Ein Beitrag zur Optik der Farbanstriche.’ Z. Tech. Phys. 1931 (10) 593-601).
6.1 Bare SNP5 and Bare SNP80
The results of the DRIFT analysis are shown in
6.2 SNP-DDAB
To further characterize the adsorption of DDAB to SNPs, DRIFT analysis of the coated SNPs was performed.
The maximum of the υO—H band shifts considerably to lower wavenumbers upon DDAB adsorption and as DDAB concentration increases up to 250 μg/mg silica, indicating that the fraction of the more interacting hydroxyl groups increases with the DDAB load. This effect is clearer for SNP80 and suggests the involvement of OH groups as anchor sites for DDAB. However, the shift becomes very small when increasing the DDAB load to 500 μg/mg silica, which shows that further modifications at the SNP surface are negligible (
ν—wavenumber of maximum absorption, cm1−; % A percentual area; LO (SiO)x—longitudinal-optic component of siloxane x-rings; TO (SiO)x—transverse-optic component of siloxane x-rings; δ(SiO)x—LO-TO splitting of siloxane x-rings; %(SiO)x—total percentual area of siloxane x-rings.
In
Taking into account that the SNP5 are much richer in OH groups, a higher adsorption yield would be expectable for low DDAB concentrations, while the first layer is being formed (up to 125 μg/mg silica). This was confirmed by the TGA results and also by DRIFT, comparing the relative intensities of the CH2 and CH3 deformation bands versus the main silica band.
Based on the concentrations of DDAB adsorbed to SNPs and on the efficiency of the adsorption process, the ratio 250 μg of DDAB per mg of SNPs was selected (SNP-DDAB250) for further experiments.
Glass coverslips were coated as described in Paulo C. et al. ‘Antifungal nanoparticles and surfaces’. Biomacromolecules 2010 (11) 2810-2817. Basically, round glass coverslips (Ø=12 mm) were initially cleaned by ultrasonication in successive 10 min steps in acetone, methanol:H2O (1:1) and chloroform. The coverslips were then placed in 24-well plates, one per well, immersed in 0.5 ml dopamine hydrochloride solution (2 mg/ml in 10 mM Tris buffer pH 8.5), shaken overnight (150 rpm orbital shaking) and finally rinsed with distilled water. The coverslips were dried and covered with a suspension of SNP5 or DDAB250-SNP5 (20 μl each, 5 mg/ml in 10 mM Tris buffer pH 8.5), dried and immersed in 0.5 ml Tris buffer overnight. This process was repeated three times. The washing water was analysed by DLS via Zeta PALS Zeta Potential Analyzer in order to assess the number of particles removed from the surface (given by the counts per second). It was possible to immobilise 222.8 and 243.9 μg/cm2 of SNP5 and DDAB250-SNP5, respectively. Because no amine or sulfhydryl groups exist in DDAB250-SNP5, π-π interactions and hydrogen bonding likely mediate the immobilization of SNPs to the polydopamine-coated surface.
SNP5 or SNP80 coated with DDAB (SNP5-DDAB250 or SNP80-DDAB250) were tested in suspension against C. albicans, Aspergillus oryzae ATCC 46244 (A. oryzae; mold), Penicillium ochrochloron ATCC 9112 (P. ochrochloron; mold), S. aureus (bacteria gram-positive) and E. coli (bacteria gram-negative). Yeast Peptone Dextrose (YPD) and Tryptone Soy Yeast (TSY) media were used to culture yeast and bacteria, respectively.
DDAB250-SNP5 or DDAB250-SNP80 suspended in culture medium (500μg/ml) was added to (i) 1 ml YPD media containing 1×105 cells of C. albicans, (ii) 1 ml TSY medium containing 1×106 cells of E. Coli or S. Aureus, or (iii) 1 ml water containing 1×105 cells/l of A oryzae or P. ochrochloron spores. Cells were incubated for 6 h at 30° C. (fungi) or 37° C. (bacteria) with orbital shaking (150 rpm). Then, an aliquot of the medium was serially diluted in sterile water and plated on YPD agar plates (1% yeast extract, 2% peptone, 2% dextrose, 2% agar) or TSY agar plates (3.3% tryptic soy broth, 0.3% yeast extract, 1.7% agar). Finally, the plates were incubated at 30° C. and 37° C. for 18 h and the number of colony forming units (CFU) was counted and compared with the controls (SNP5 or SNP80) and the inoculum (prokaryotic cells incubated without SNPs). Minimum inhibitory concentrations (MICs) were determined by the broth microdilution method according to the National Committee for Clinical Laboratory Standard (for yeasts; NCCLS; M27A2E) and the Clinical and Laboratory Standards Institute (for bacteria; CLSI; M07-A8) guidelines.
To evaluate the anti-mold activity of SNP, suspensions of spores from Aspogillus or Penicillium were prepared from grown cultures on Potato Dextrose Agar (PDA) plates at 30° C. DDAB250-SNP80 suspended in sterile distilled water (500 μg/ml) was incubated with a suspension of spores (1×105 spores/ml, in sterile distilled water) for 6 h, at 150 rpm and 30° C. At the end of the incubation, an aliquot of the medium was serially diluted in sterile distilled water and plated on PDA plates (3.3% potato dextrose broth, 1.1% agar). The plates were then incubated at 30° C. and CFU counted after 3 days and compared with the controls (SNP5 or SNP80) and the inoculum (spores without SNP).
To evaluate the biological activity of DDAB250-SNP5 or DDAB250-SNP80 against yeast, bacteria and molds in multiple challenges, the particle suspension (3 mg/ml) was centrifuged (20 min, 14,000 rpm) after the previous antimicrobial activity assay and the SNPs resuspended in YPD medium containing C. albicans (1×105 cells/ml), TSY medium containing S. aureus or E. coli (1×106 cells/ml), or distilled water containing spores (1×106 spores of A. oryze or P. ochrochloron). The suspension was incubated in an orbital shaker for 6 h at 30° C. (fungi and molds) or 37° C. (bacteria) and the number of CFU was determined as described above.
Growth kinetics of C. albicans, S. aureus and E. coli (1×105 cells/ml) exposed to variable concentrations of DDAB250-SNP5 or DDAB250-SNP80 (from 3.1 to 50 μg/ml for fungi and S. aureus and from 150 to 1000 μg/ml for E. coli) were evaluated by absorbance at 600 nm. Inoculum corresponds to prokaryotic cells incubated without SNPs. The samples were prepared as described above.
As described above, after 6 h of exposure an aliquot of the cell suspension was plated on agar plates for 18 h and the number of yeast, bacteria or mold colonies was counted (
The MIC for SNP5-DDAB250 against C. albicans, S. aureus, and E. coli was 25, 12.5 and 500 μg/ml, respectively, while for SNP80-DDAB250 was 50, 25 and above 1000 μg/ml (Table 4). At these concentrations, coated SNP5 formulations have 4.5, 2.2 and 89.1 μg of DDAB, while SNP80 have 9.7, 4.9 and 194.4 μg of DDAB, respectively (Table 4). Importantly, with the exception of MIC values obtained against E. coli, the MIC values of SNPs coated with DDAB are much below the MIC of DDAB solutions (C. albicans: 125 μg/ml; S. aureus: 31.25 μg/ml; E. coli: 125 μg/ml).
To assess the relative contributions from surfactant release and SNP killing, SNP80-DDAB250 were incubated in YPD or TSY mediums or distilled water for 6 h, centrifuged and the supernatant collected and tested against microorganisms. Limited killing was observed (below 25%) suggesting that antimicrobial activity of the SNPs is mainly mediated by contact (
The antimicrobial activity of DDAB250-SNP80 over time was evaluated as follows. Samples of DDAB250-SNP80 (1 mg/ml) were maintained in 0.1 M citrate/sodium citrate buffer pH 3.0, PBS pH 7.4, 0.1 M borate/NaOH buffer pH 9.0 and YPD medium for 60 days. The buffers and the YPD medium were changed every 3 days. For that purpose, SNPs were centrifuged at 14,000 rpm for 20 min, and the buffer/medium replaced by a new one. After 60 days, the SNPs were centrifuged (same conditions as before), washed with distilled water, and freeze-dried before testing antifungal activity with C. albicans (1×105 cells/ml). After an incubation of 6 h, an aliquot of the medium was serially diluted in distilled water and plated in agar to assess the number of microorganisms. The results are shown in
Similar tests were performed with bacteria (E. coli and S. aureus) and molds (A. oryzae and P. ochrochloron) and the results are shown in
The antimicrobial activity due to the leaching of the SNPs was evaluated by washing the glass coverslips (sterilized for 30 min under UV light) coated with SNP5 and SNP5-DDAB250 for 6 h with 1 ml of YPD (at 30° C.) or TSY (at 37° C.), under orbital shaking (150 rpm). The washing medium was tested against 1×105 C. albicans cells, 1×106 E. coli or S. aureus cells. No measureable antimicrobial activity was detected (
After confirming the absence of antimicrobial activity in the washing solutions, the antimicrobial activity of the surfaces was tested by adding 1 ml of YPD with 1×105 C. albicans cells or 1 ml of TSY with 1×103 E. coli or 1×106 S. aureus cells to each coverslip and incubating at 30° C. (fungi) or 37° C. (bacteria) for 6 h, under orbital shaking (150 rpm). An aliquot of the medium was serially diluted in sterile distilled water and plated on YPD or TSY agar plates. The number of CFU was counted after incubation of the plates at 30° C. or at 37° C. for 24 h. The remaining medium was removed and the coverslips were rinsed twice with 1 ml of sterile water. After drying rapidly, the coverslips were plated with coated side down on YPD or TSY agar plates and incubated at 30° C. or at 37° C.
A 100% reduction in C. albicans, E. coli and S. aureus was observed in the media containing the coverslips coated with SNP5-DDAB250 relative to the control (coverslips coated with SNP5). Finally, the remaining medium was removed and the coverslips were rinsed with sterile water to remove non-adherent cells and plated upside down on YPD or TSY agar plates. After 24 h, no C. albicans, E. coli or S. aureus colonies were observed on the coverslips coated with SNP5-DDAB250, whereas fungi and bacteria colonized the control coverslips coated with SNP5 (
The size of the coated SNPs were determined using dynamic light scattering (DLS) as described in Example 1.2.3. The results of this analysis are shown in Table 4.
C. albicans
S. aureus
E. coli
aData corresponds to the average ± standard deviation of three independent measurements by DLS.
bThe value in parentheses is the amount of DDAB (μg) in the SNPs.
MDCK cells obtained from the European Collection of Cell Cultures agency (ECACC), were grown in Dulbecco's Modified Eagle's Medium supplemented with 10% fetal bovine serum (FBS), 100 units/ml penicillin and 100 μg/ml streptomycin, at 37° C. in a humidified-air atmosphere containing 5% CO2. Influenza A/PR/8/34 (H1N1) was obtained from Advanced Biotechnologies.
The antiviral activity of the coated glass coverslips was assessed by a methodology described in Halder et al. ‘Preparation, application and testing of permanent antibacterial and antiviral coatings’. Nat. Protoc 2007 (2) 2412-2417. Briefly, glass coverslips (glass, glass dopamine coated with SNP5 and glass dopamine coated with SNP5-DDAB250) were placed in 24-well plates and 20 μl of virus solution was deposited on the top of each sample and covered with parafilm to spread the drop. After 30 min. of incubation at room temperature, the samples were washed with 1.98 ml PBS and 2-fold serial dilutions were made. Then, confluent MDCK monolayers were washed twice with PBS and infected with 200 μl of each dilution for 1 h at room temperature. After incubation, the virus solution was aspirated and the cells were covered with agar medium (1:1 DME/F12 modified supplemented with 0.01% DEAE-dextran, 0.1% NaHCO3, 4 μg/ml trypsin, 100 units/ml penicillin, 100 μg/ml streptomycin and 0,6% agar) and incubated for 4 days at 37° C. in a humidified-air atmosphere containing 5% CO2. Finally, the agar overlay was removed and the cells were fixed with paraformaldehyde and stained with crystal violet (0.1% in 20% v/v aqueous methanol).
The antiviral activity of the surfaces due to the leaching of SNPs was evaluated by washing the coverslips with PBS (1 ml) for 6 h under orbital shaking and incubating the washing with 20 μl of virus solution for 1 h at room temperature. The number of viral colonies was determined as before.
No measurable antiviral activity was observed (
SWCNT were chosen for use in this study as they have been shown to exhibit antimicrobial activity (Kang et al. ‘Single-walled carbon nanotubes exhibit strong antimicrobial activity.’ Langmuir 2007 (23) 8670-8673) which is stronger than for MWCNTs (Arias et al ‘Inactivation of Bacterial Pathogens by Carbon Nanotubes in Suspensions. Langmuir 25 (2009) 3003-3012)
Carboxylated single-wall carbon nanotubes of 1.5 nm in diameter and 1-5 μm in length (>95% purity; 2-7 wt % COOH by titration) were purchased from NanoLab (USA) and used without further processing. SWCNTs and DDAB (Sigma-Aldrich) were mixed in a mass ratio from 0.1 to 100 (e.g. 1500 μg/ml DDAB is mixed with 100 μg/ml SWCNT suspension yielding a mass ratio of 15) in PBS and then sonicated. A 6 mm probe was used with sonication cycles of 1 h with 5:1 seconds of pulse (on:off). The amplitude used was typically 50%. After each cycle the released energy was registered and the optical density at 600 nm was measured. Typically, centrifugation cycles were performed on a centrifuge at 10,000 g for 20 min, to remove the aggregates.
The evolution of SWCNTs resuspension was performed by optical density measurement (at 600 nm) on a microplate reader. N-phenyl-1-naphthylamine (NPN) (Sigma-Aldrich) was used to determine the DDAB critical micelle concentration (CMC) and the presence of micelles on the SWCNT suspension. A solution of NPN of 2×10−1 M in ethanol was prepared and further diluted in PBS to the working concentration of 2×10−4 M. DDAB or SWCNT-DDAB suspensions were mixed in equal volumes on a opaque 96-well plate and the fluorescence measured with the assistance of a SpectraMax Fluoremeter using an excitation and emission wavelength of 320 and 450 nm, respectively.
In order to be able to accurately assess the antimicrobial activity of SWCNT:DDAB it was imperative to determine whether free surfactant was co-existing in solution with the complex SWCNT:DDAB. A centrifugation step to separate SWCNT:DDAB from free DDAB is not a reliable methodology since it requires extensive centrifugation velocities and the nanotubes tend to aggregate. It was observed that increasing the mass ratios of DDAB to SWCNTs from 2.5 to 100 decreased substantially the recovered pellet (
Alginic acid and calcium chloride were purchased from Sigma-Aldrich. Calcium-alginate microparticles were prepared at room temperature with a Nisco Nozzle Var-J1 (Nisco, Switzerland) and coaxial nitrogen air flow. Briefly, an alginate solution (0.5-2%) was extruded with controlled flow (10-200 ml/h) through a variable internal diameter needle (ID ˜200-450 μm). The controlled coaxial flow of nitrogen (0.5-15 L/min) cuts the alginate flow on the other edge of the needle, projecting the alginate drop against a calcium chloride solution of variable concentration (0.05-1 M) with or without magnetic stirring. The needle length (1-5 mm) and nozzle height (2-20 cm) to the calcium solution were also screened. Each sample was visualized and photographed under the contrast phase microscope and the sizes were assessed with ImageJ.
Magnetic nanoparticles were acquired from PlasmaChem (Germany). To prepare the alginate microparticles-loaded with DDAB-SWCNT and magnetic nanoparticles defined standard concentrations for each of the components were determined. The typical extruded composite suspension is composed of 2% alginate, 10-100 μg/ml of magnetic nanoparticles (PlasmaChem, Germany), 25 μg/ml of SWCNT and DDAB. The latter amount depends on the DDAB/SWCNT mass ratio chosen. This mix was typically extruded against a 0.1 M CaCl2 solution at a flow rate of 10-20 ml/h (needle with 200 μm i.d.) or 50-100 ml/h (needle with ˜420 μm i.d.). A nitrogen coaxial flow rate of 5-8 l/min was used to generate beads.
Typically, SWCNTs coated with DDAB or other surfactants were diluted taking in account the initial surfactant concentration being the SWCNT concentration estimated by the corresponding mass ratio used. The antimicrobial activity of the coated SWCNTs was tested in suspension against Candida albicans ATCC 10231 (C. albicans, fungi), Staphylococcus aureus ATCC 6538 (S. aureus, bacteria gram-positive) and Escherichia coli ATCC 25922 (E. coli, bacteria gram-negative). Yeast Peptone Dextrose (YPD) and Tryptone Soy Yeast (TSY) media were used to culture yeast and bacteria, respectively. Growth kinetics of the different microorganisms (1×105 cells/ml) exposed to variable concentrations of DDAB-SWCNT suspended in PBS (500-0.3 μg/ml of DDAB which corresponds to a 33-0.002 and to a 5-0.0003 μg/ml of SWCNT under a mass ratio of 15 and 100, respectively) was evaluated by absorbance at 600 nm. Inoculum corresponds to cells incubated without CNTs. The samples were prepared in a 96-well plate and incubated for 18 h at 30° C. (fungi) or 37° C. (bacteria) with constant shaking in a Biotek Synergy Mx spectrophotometer.
DDAB-SWCNTs (mass ratio of DDAB:SWCNT) of 15 and 100 showed high antimicrobial activity against C. albicans, S. aureus and E. coli, exhibiting minimal inhibitory concentrations (MIC) of approximately 2, 0.5 and 31 μg/ml, respectively (Table 4). These MIC values compare favourably to the ones obtained for silica nanoparticles coated with DDAB. Also, these values compare favourably to the ones reported in the literature using CNTs. The lowest concentration ever reported for an effective antimicrobial activity of SWCNTs in suspension (although in this case forming aggregates) was 1 μg/ml (Kang et al. ‘Single-walled carbon nanotubes exhibit strong antimicrobial activity’. Langmuir 2007 (23) 8670-8673).
C. albicans
S. aureus
E. coli
Advantageously, DDAB acts simultaneously as a dispersant agent of the SWCNTs and as an antimicrobial agent.
It is apparent that the antimicrobial complexes of the invention demonstrate effective antimicrobial activity both in suspension and when immobilized on surfaces. The surfactants exhibit low minimal inhibitory concentrations compared with soluble surfactants. Thus, the complexes of the invention can be used to prepare highly active antimicrobial coatings with a broad spectrum of activity. The inventors have shown that when the nanomaterial complexes are applied to a surface, a non-leaching surface coating is obtained such that a long-lasting antimicrobial effect can be achieved.
The invention also encompasses methods of preparing the nanomaterial complexes and of applying the complex to a surface to create an antimicrobial coating.
| Number | Date | Country | Kind |
|---|---|---|---|
| 12398001.3 | Apr 2012 | EP | regional |
| Filing Document | Filing Date | Country | Kind |
|---|---|---|---|
| PCT/EP2013/058166 | 4/19/2013 | WO | 00 |
