Antibacterial textiles functionalized by layer-by-layer assembly of polyelectrolytes and TiO₂ photocatalyst

Abstract

The sprayed layer-by-layer assembly of TiO₂ photocatalyst has been efficiently used for functionalizing polyester textiles and elaborating solar light active antibacterial textiles. Polyethylenimine (PEI) and polyanionic poly(styrene sulfonate) (PSS) polyelectrolytes have been used as polycation and polyanion layers, in alternation with dispersions of negatively- and positively-charged TiO₂ particles, respectively. A synergistic effect resulting from both biocidal photocatalytic activity of TiO₂ layers and biocidal activity of PEI layers within the multilayer system was put forward for explaining the high self-decontaminating activity of (PEI/TiO₂)_n functionalized textiles, 5 log of cell viability reduction being achieved with a single TiO₂ layer at 50 μg cm⁻². The sole solar light photocatalytic antibacterial activity of multilayer textiles was determined by masking the biocidal activity of PEI layers through the controlled assembly of PEI/(PSS/TiO₂)_n multilayers.

---

1. Introduction

Textile substrates are known to be suitable environments for microbial growth under certain conditions. Numerous microorganisms can rapidly multiply at the textile surface, so that they can be responsible for fabric deterioration,¹ staining or unpleasant odors,^2,3 and even can play a role in acquisition and transmission of pathogens.^4–8 Depending on operating parameters and on the inoculum load, survival of Gram negative bacteria such as e.g. Escherichia coli, Klebsiella pneumoniae or Pseudomonas aeruginosa on textiles has been observed from 2 h until more than 60 days, while periods of months have been reported for Staphylococci and Enterococci on polyester.^9–11 Persistence and lifespan of microorganisms on textiles depend on multiple factors, including the textile composition, the type of bacterial species and the environment.

Thereby, within a global approach dealing with the design of smart advanced textiles, the elaboration of antimicrobial or self-disinfecting textiles is especially receiving growing interest, aiming at preventing bacterial growth on the textiles.

Thus, many antimicrobial agents have been used for providing bactericidal properties to textiles, such as quaternary ammonium, triclosan, polyhexamethylene biguanide, zinc pyrithione, Ag nanoparticles or salts,^12–15 the most studied approach dealing nowadays with the use of Ag nanoparticles.^16,17 However, developing new disinfection approaches is required due to fast adaptation of bacterium and development of metal-resistant strains,¹⁸ and applying anti-bacterial inorganic nanoparticles would be an alternative for opening up new opportunities for antimicrobial textiles.^19,20

Oxidative heterogeneous photocatalysis is a promising disinfection approach based on the irradiation of a semi-conductor catalyst by light with energy matching or greater than its band gap. The resulting light absorption enables the generation of electron and hole carriers and subsequently of highly oxidative species – Reactive Oxygen Species (ROS) – at the catalyst surface, responsible for inducing damages to bacterial cells leading to cell death.^21–27 Titanium dioxide (TiO₂) is the most used photocatalyst under UV light, thanks to high quantum yield, both chemical and light stability, non toxicity as well as low cost for commercial standards.^21,24,25

Even if finishing techniques used in textile industry can to a certain extent be adapted for immobilizing TiO₂ onto textiles, they suffer from using additives in large amount like surfactants or thickening agents, that block the reactant access to the TiO₂ nanoparticles. Alternative approaches have thus been developed for synthesizing and immobilizing nanostructured oxides such as TiO₂ onto textile fibers.^28–34 One can cite e.g. sol–gel methods, direct current reactive magnetron sputtering, deposition of TiO₂ through linking with spacer agents, as well as deposition from TiO₂ suspensions. In some of them, TiO₂ was associated to Ag as antibacterial agent.^35–37

Layer-by-Layer assembly (LbL) is a multilayer approach based on electrostatic interaction of alternative layers of oppositely-charged materials used for building polyelectrolyte and multi-material layered films on substrates via a single process.^38–40 It has been used recently for building TiO₂ photo-catalytic films for environmental or detoxification purpose.^41–43 This approach was scarcely applied to textiles,^44,45 and only few works reported on LbL functionalization of commercial textiles by TiO₂ for providing them self-cleaning efficiency towards chemicals.^46–50 In particular, we proved that cotton/polyamide textiles with photocatalytic activity in the degradation of chemicals, could be obtained by substituting the polyanion layer by a dispersion of negatively-charged TiO₂ particles in alternation with polyethylenimine (PEI) as polycation layer.⁴⁷

Thus, we report herein on new LbL-functionalized photocatalytic multilayer textiles designed for exhibiting high self-decontaminating activity towards bacteria. The underlying strategy implemented here was to take advantage of both biocidal photocatalytic activity of TiO₂ nanoparticle layers and biocidal activity of alternative PEI polyelectrolyte layers – known to have good antibacterial properties⁵¹ – for providing a synergistic effect within the multilayer system resulting in the elaboration of high activity anti-bacterial textiles.

2. Experimental

2.1. Materials and textiles

Aeroxide® TiO₂ P25 was supplied by Evonik (Germany, 20% rutile – 80% anatase crystalline form, with a specific surface area of 50 m² g⁻¹).⁵² Synthetic 100% polyester textiles (PES) were delivered by Ouvry (France). They were washed with hot water, rinsed with distilled water and dried at 100 °C overnight.

The raw PES fiber textile was first submitted to a mild hydrolysis using KOH solution for generating negatively charged surface oxygen species, allowing to strengthen the interaction with the first positively charged PEI layer and consequently improving the adhesion of the first layer on the fibers. The textile was immersed into 0.5 M KOH aqueous solution for 30 min under soft orbital shaking (150 rpm). The fabric was then rinsed with distilled water until neutral pH is reached and then dried at 100 °C for 5 min.

In this study we used polyester textiles which are frequently found in a hospital environment e.g. drapes. Moreover, as compared to other textiles (cotton or nylon clothes), polyester has binding affinity towards bacteria.⁵³

2.2. Sprayed layer-by-layer coatings

The (PEI/TiO₂)_n layer-by-layer functionalization consists of alternate layers of polycationic PEI (BASF Lupasol/Water free, MW ∼ 25 000 g mol⁻¹) in aqueous solution (8 g L⁻¹) and TiO₂ P25 at 5 g L⁻¹ in water/ethanol (50 : 50 in volume) at pH 9 (adjusted with a few drops of NaOH solution to ensure negatively charged surface of TiO₂ as its isoelectrical point is around 6.5). The polyelectrolyte solution and the TiO₂ dispersion were filled into spray bottles (Air-boy from Carl Roth, 400 μm nozzle) and manually pressurized to about 1 bar. For the deposition, the dried PES fabrics were horizontally fixed onto metallic grids, and PEI solutions and TiO₂ nanoparticle dispersions were sprayed on from about 15 to 20 cm distance in crossed lines. Each spraying duration was about 20 s. After each layer, the fabric was allowed to equilibrate with the reactant for 10 min while still laying and then rinsed abundantly vertically with distilled water to eliminate any excess. After rinsing, the fabrics were dried at 100 °C for 5 min before the next bilayer was applied. This sequence was repeated until the number of PEI/TiO₂ bilayers was achieved. In this study, we used n = 1, 5 and 10 bilayers. After the last TiO₂ layer, the functionalized textiles were dried at 130 °C for 30 min.

Anionic poly(styrene sulfonate) (PSS) was also used as polyelectrolyte for functionalizing the PES textiles with TiO₂. In that case, an interfacial layer of PEI was added before building the (PSS/TiO₂)_n bilayers. After the first PEI layer, the textile was rinsed with distilled water and dried for 5 min at 100 °C. Then, the (PSS/TiO₂)_n bilayers consisted of alternate layers of PSS (MW ∼ 70 000 g mol⁻¹, Sigma-Aldrich) at 1 g L⁻¹ in distilled water rectified at pH 2.5 with HNO₃, and TiO₂ P25 at 10 g L⁻¹ in water/ethanol (50 : 50 in volume) at pH 2.5 (adjusted with HNO₃) to ensure positively charged surface of TiO₂. Our previous works have reported that pH of 2.5 was ideal for maintaining a strong positive charge on the nanoparticles and sufficient dissociation of the PSS.⁴³ This (PSS/TiO₂)_n sequence was repeated until the number of bilayers was achieved. In this study, we used n = 1, 5 and 10 bilayers.

2.3. Characterization techniques

The TiO₂ loading on the textiles was determined by Inductively Coupled Plasma Atomic Emission Spectroscopy (ICP-AES). Scanning electron microscopy (SEM) was performed on a Jeol JSM-6700F (1–10 kV), with metallization of samples with gold.

2.4. Bacterial strains and growth media

Antimicrobial activity of functionalized textiles has been determined against two Gram negative bacteria, E. coli and P. aeruginosa. Before each experiment, one loop full of E. coli ATCC 8739 or P. aeruginosa CIP A22 was seeded on a slant of tryptic soy agar (TSA) (BioRad), and grown aerobically at 37 °C for 24 h. Bacterial inoculum was monitored by setting the culture optical density at an absorbance wavelength of 620 nm (OD₆₂₀) at 0.156, corresponding to 10⁸ CFU mL⁻¹. These populations were diluted in Mueller-Hinton broth (MHB) to obtain an inoculum of 10⁶ CFU mL⁻¹.

2.5. Assessment of the bactericidal activity of PEI and PSS polyelectrolytes against E. coli and P. aeruginosa bacteria

2.5.1. Determination of the minimum inhibitory concentration (MIC). MIC of PEI and PSS polyelectrolytes against E. coli and P. aeruginosa bacteria is defined as the lowest concentration of the polyelectrolyte enabling the visual inhibition of bacterial cultivation after 18 h of incubation. They were determined against E. coli and P. aeruginosa bacteria by broth dilution method using MHB as recommended by National Committee for Clinical Laboratory Standards.⁵⁴ The 96-well plates were prepared by dispersing into each well 50 μL of MHB, 100 μL of bacteria suspension at 1 × 10⁶ CFU mL⁻¹ supplemented with 50 μL of PEI or PSS solution at different concentrations. So, 10⁵ bacterial cells in MHB were treated with serial twofold dilutions of PEI (in a concentration range from 0.625 g L⁻¹ to 40 g L⁻¹) or PSS (in a concentration range from 0.0625 g L⁻¹ to 8 g L⁻¹) in distilled water. Controls (without any polyelectrolytes) were performed with pure distilled water. The plates were incubated at 37 °C for 18 h. The minimum concentration at which there was no turbidity by measuring at OD₆₂₀ was taken as the MIC of polyelectrolytes.

2.5.2. Determination of the minimum bactericidal concentration (MBC). MBC of PEI and PSS polyelectrolytes against E. coli and P. aeruginosa bacteria is defined as the lowest concentration of polyelectrolyte that reduced the initial inoculums by at least 3 log. Thus, the initial inoculum at 10⁶ CFU mL⁻¹ was diluted in sterile physiological water to obtain a 3 log reduction. 10 μL of each dilution was deposited on Mueller Hinton agar (MHA) and incubated for 24 h at 37 °C. This plate was then kept in the fridge during 24 h. It used to observe the 3 log reduction in order to determine MBC.

Bacterial cultures treated by PEI or PSS polyelectrolytes showing no growth in the MIC tests were used for assessing MBC. 10 μL bacterial cultures treated by polyelectrolytes showing no growth were plated onto MHA plate and incubated at 37 °C for 24 h. This plate was compared with the one previously made for the initial inoculum. The MBC plate (at the lowest dilution) containing equal or less than number colony compared to the 3 log reduction plate was defined as the MBC.

2.6. Determination of the antimicrobial efficiency of functionalized textiles

The antibacterial properties of functionalized textiles against E. coli and P. aeruginosa were assessed according to a modified protocol derived from that of the BS ISO27447 test method.⁵⁵

Self-disinfecting tests under solar light were performed within an ATLAS Suntest XLS+ testing chamber equipped with Xe arc lamp and daylight filter simulating outdoor solar light, inside which 2 × 2 cm² sized textiles pieces were located and submitted to a total irradiance of 242 W m⁻² including 226 W m⁻² of visible light and 16 W m⁻² of UV-A. The spectral distribution is reported in ESI S1.† The temperature was regulated at 33–35 °C – an adequate temperature to survival of the selected bacteria – through a water-cooled plate. To avoid any desiccation, the textiles were pre-humidified with 70 μL of sterile physiological water (NaCl, 9 g L⁻¹). 100 μL of bacterial inoculum at 10⁸ CFU mL⁻¹ were deposited on each textile, corresponding to a bacteria load of 10⁷ CFU per textile.

After test under light or in the dark, each textile was introduced into a stomacher bag containing 10 mL of sterile physiological water supplemented with 2% of Tween 80 (Sigma-Aldrich, France) – acting as non-ionic surfactant and solubilizing agent – and was kneaded for 2 min (Stomacher Mayo, Homogenius HG400 V). After this step serial decimal dilution were made in physiological water. 1 mL of each dilution were added to 19 mL of nutrient agar media in surfusion (55 °C) containing 3 g L⁻¹ of beef extract (Difco), 5 g L⁻¹ of pancreatic digest of casein (Biokar diagnostics) and 15 g L⁻¹ of bacteriologic agar type A (Biokar diagnostics). The plates were incubated for 24 h at 37 °C and bacteria included in the agar media or growth at the surface were numerated.

The antibacterial efficiency of the textiles was assessed by numerating the bacteria in terms of Colony Forming Units (CFU) after solar light exposure, and determining the bacteria survival over the textiles. Controls have been performed in the dark as well. The data are presented as the mean ± standard error of the mean of at least three independent experiments. Statistical analysis was performed by the Student's t test (Prism 5, GraphPad Software). A value of p < 0.05 (*), p < 0.01 (**), p < 0.001 (***) or p < 0.0001 (****) was considered to be statistically significant.

3. Results and discussion

3.1. Characterization of functionalized textiles

Fig. 1 shows SEM images of PES–KOH textiles functionalized with PEI/TiO₂ and PSS/TiO₂ multilayers at the different steps of the film building, as well as the influence of the number of bilayers applied for both multilayer systems, on the TiO₂ wt content and the TiO₂ surface density.

Fig. 1 SEM images of (A and C) bare PES–KOH, (B) PES–KOH/PEI/(PSS/TiO₂)₁₀, (D) PES–KOH/(PEI/TiO₂)₁, (E) PES–KOH/(PEI/TiO₂)₅, (F) PES–KOH/(PEI/TiO₂)₁₀ textiles (G) PES–KOH/PEI/(PSS/TiO₂)₅ and (H) PES–KOH/PEI/(PSS/TiO₂)₁₀; (I) influence on the TiO₂ wt content and the TiO₂ surface density, of the number of bilayers applied on the PES–KOH/(PEI/TiO₂)_n and the PES–KOH/PEI/(PSS/TiO₂)_n textiles. Error bars were calculated from measurements over three samples.

Previous works evidenced that, at full coverage of the substrates, the average thickness of the first PEI layer determined by ellipsometry was 11.5 Å.⁵⁶

Fig. 1-I shows that the TiO₂ wt content – and thus the TiO₂ surface density – linearly increased with the number of bilayers applied on the fibers in the case of the PEI/(PSS/TiO₂) multilayer system, with a TiO₂ wt content of 2.5 wt% for 10 bilayers. By contrast, the TiO₂ wt content saturated more rapidly at 1.8 wt% for 10 bilayers in the case of the (PEI/TiO₂) system. Although not studied on textiles, a linear increase in the film thickness in the 20–30 bilayer range is usually observed for PSS/TiO₂ ^42,57 as well as PEI/TiO₂ ⁵⁸ photocatalytic systems on glass model substrates. The saturation observed here for PEI/TiO₂ bilayer systems differed from the linearity reported by Rojas-Blanco et al.⁵⁸ and could result from a possibly different building up of the film on woven irregular and curved PES fibers vs. flat model substrates when using PEI for structuring and elaborating the layered film rather than only as first anchorage/glue layer on the PES fibers.

3.2. Antimicrobial activity of polyelectrolytes in solution

The antimicrobial activity of PEI polycation and PSS polyanion was determined against E. coli and P. aeruginosa bacteria by measuring growth inhibition. It is well admitted that compounds with a MBC/MIC ratio ≤ 4 or >4 are considered as bactericidal or bacteriostatic, respectively.^54,59 For both bacteria, the PEI polycation displayed similar values of MIC and MBC, at 2.5 g L⁻¹ (Fig. 2a and c), so that PEI with MW ∼ 25 000 g mol⁻¹ was evidenced as bactericidal agent against E. coli and P. aeruginosa. This confirmed works performed with PEI over Gram negative and Gram positive bacteria such as E. coli, P. aeruginosa, S. aureus and S. epidermidis.^60–65 It was proposed that the bactericidal effect results from the fact that hydrophobic polycationic chains were able to traverse and irreparably damage the bacterial cellular membrane, by interacting with and disrupting negatively charged bacterial cell membrane followed by release of K⁺ ions and other cytoplasmic constituents.^61,62

Fig. 2 Antibacterial effect of PEI (a and c) and PSS (b and d) in g L⁻¹ against E. coli (a and b) and P. aeruginosa H21 (c and d) bacteria determined by measuring the optical density at 620 nm.

By contrast, MBC and MIC values for the PSS polyanion were higher than 8 g L⁻¹ for both bacteria (Fig. 2b and d for MIC, data not shown for MBC). Thus, neither a bacteriostatic nor a bactericidal effect was evidenced for PSS.

3.3. Antibacterial activity of textiles functionalized with (PEI/TiO₂)_n

Fig. 3 shows the antibacterial effect against E. coli achieved after 60 min under solar light over the PES textile functionalized with PEI/TiO₂ multilayers at the different steps of the multilayered film building, i.e. with one single PEI layer (Fig. 3C), and with 1 (Fig. 3D) or 5 bilayer(s) (Fig. 3E) of (PEI/TiO₂). Controls have been performed in the dark.

Fig. 3 Antibacterial effect against E. coli achieved after 60 min under solar light and in the dark over the PES textile functionalized with PEI/TiO₂ multilayers at the different steps of the film building, i.e. PES (A), PES–KOH (B), PES–KOH/PEI (C), PES–KOH/(PEI/TiO₂)₁ (D), PES–KOH/(PEI/TiO₂)₅ (E).

First, 2.4 × 10⁶ and 6.8 × 10⁶ CFU of E. coli have been numerated over the textiles in the dark, as-it (Fig. 3A) and after KOH pre-treatment (Fig. 3B), respectively, corresponding to about 20–70% of the 10⁷ CFU inoculums seeded on the textiles. This confirmed the expected neutrality against E. coli survival of the KOH treatment of the fiber surface, and further evidenced the efficiency of the bacteria recovery method used in comparison to usual surface sampling methods such as e.g. the RODAC contact plate method, swabbing techniques or wash-off methods.⁶⁶ Indeed, many studies reported that the amount of microorganisms recovered through the RODAC plate technique was much lower than the initial applied concentration, as evidenced by Rabuza et al. with recovery ratios lower than 10⁻⁵.⁶⁶ The method adapted from the BS ISO27447 standard method is thus adequate for extracting and further quantifying microorganisms which may penetrated deep into the structure of PES fiber textile, in contrary to the RODAC method. In addition, Fig. 3A and B confirmed the expected neutrality against E. coli of the exposure to solar light for 60 min of contaminated bare textiles, whether the textiles were pretreated or not by KOH.

The functionalization of the textile with a single layer of PEI resulted in a significant reduction of the cell viability in the dark as well as under light, with 1.5 × 10⁵ CFU per textile being measured in both cases (Fig. 3C), and corresponding to a bactericidal activity close to 2 log as compared to the control textile (Fig. 3A). A stronger reduction of the cell viability with a bactericidal activity close to 5 log was recorded in the dark and under light over the textile functionalized with a single PEI/TiO₂ bilayer (Fig. 3D). Further, the textile functionalized with 5PEI/TiO₂ bilayers exhibited a reduction of E. coli viability greater than 6 log in the dark as well as under light (Fig. 3E), the CFU quantity being relatively close to the detection threshold which is around 1.0 × 10¹ CFU per textile.

We proposed here that this high activity resulted from a synergistic antimicrobial activity between PEI and TiO₂. First, it has been reported that TiO₂ exhibited an antibacterial activity already in the dark,⁶⁷ by impacting key cellular components of bacteria. This effect was amplified under illumination through the production of ROS such as O₂˙⁻ superoxide radical, causing lipid peroxidation enhancing membrane fluidity and disrupting cell integrity. Already during the cytotoxic treatment in the dark with TiO₂ – and amplified under UV-A illumination – two-dimensional electrophoresis proteomic analysis associated performed on the whole proteome of E. coli and coupled to mass spectrometry protein analysis evidenced that the main degraded proteins were porins, proteins involved in oxidative stress response, chaperone proteins, and to a lesser extent proteins involved in the transport and metabolism of different molecules such as inorganic ions, lipids, carbohydrates and amino acids. To a lesser extent, proteins involved in energy production and conversion, as well as in translation regulation were also identified.⁶⁷ Thus, it is likely that the cytotoxicity of TiO₂ could directly enhance the biocidal effect of the PEI polyelectrolyte by strongly reducing the ability of the cells to respond to oxidative stress or to regulate environmental stress.

This synergy effect could also be envisaged inversely, considering that the biocidal activity of PEI against E. coli could directly enhance the impact of TiO₂ in the dark by modifying the organization of the bacterial outer membrane. It could also be enhanced under illumination. Indeed, PEI – like more generally polycationic agents – induces the reorganization of the bilayer structure with the alignment of lipid headgroups perpendicular to the membrane.⁶⁸ This increases the permeability of the bacterial outer membrane to solutes that are normally unable to cross the outer membrane,^62,69,70 like this was already proposed for antibiotics,⁶⁴ so that this could also increase its permeability to the O₂˙⁻ superoxide radical ROS. Among the different ROS generated at the surface of TiO₂, it has been proposed that the O₂˙⁻ superoxide radical ROS was playing an important role⁷¹ due to a longer lifetime at the water–TiO₂ interface when compared to other active ROS such as OH˙ hydroxyl radicals.⁷² This extended lifetime would allow O₂˙⁻ to cross the membrane for damaging directly inner cellular components (or for further reacting to form other active ROS), while, by contrast, ROS such as the highly active OH˙ will attack first the outer cellular components due to a shorter lifetime.

This bactericidal activity of the textile functionalized with a single bilayer of PEI/TiO₂ turns out to be very promising from an applicative point of view because it requires only the application of a single layer of TiO₂ nanoparticles to get 5 log of cell viability reduction. In this case, the textile was functionalized with only 0.4 wt% of TiO₂, corresponding to a TiO₂ surface density of 50 μg cm⁻². Such a so small content confers to the functionalized textile a very good stability, necessary for targeting a realistic application as self-disinfecting textiles.

3.4. Antibacterial activity of textiles functionalized with PEI/(PSS/TiO₂)_n

Truly considered, the investigation of the PEI/TiO₂ systems on the textile revealed that the high bactericidal activity shown by the multilayers built themselves with bactericidal PEI as positively charged polyelectrolyte did not allow to evidence the real contribution of photocatalysis on light-activated TiO₂ within the overall activity of the multilayer system. So, the sole bactericidal activity due to the photocatalysis was isolated by substituting positively charged PEI by positively charged PSS within the multilayers after the first bactericidal PEI layer was applied on the textile.

Fig. 4a shows the antibacterial efficiency against E. coli bacteria achieved after 60 min under solar light over the PES textile functionalized with PEI/(PSS/TiO₂)_n multilayers at the different steps of the film building, i.e. with one single PEI layer, subsequently with one additional PSS layer, and with 1, 5 or 10 bilayer(s) of (PSS/TiO₂). Controls have been also performed in the dark.

Fig. 4 Antibacterial effect against (a) E. coli and (b) P. aeruginosa bacteria achieved after 60 min under solar light and in the dark over the PES textile functionalized with PEI/(PSS/TiO₂)_n multilayers at the different steps of the film building, i.e. PES–KOH (A), PES–KOH/PEI (B), PES–KOH/PEI/PSS (C), PES–KOH/PEI/(PSS/TiO₂)₁ (E), PES–KOH/PEI/(PSS/TiO₂)₅ (F); PES–KOH/PEI/(PSS/TiO₂)₁₀ (G).

It shows that the application of a single layer of PSS (Fig. 4a-C) on the first PEI layer allowed a partial restoration of the cell viability to be obtained, whether the textiles was irradiated or not. Indeed, the PSS polyelectrolyte did not evidence any cytotoxic effect, so that applying an additional PSS layer partly masked the cytotoxicity of the first PEI layer (Fig. 4a-B), and resulted in an increase of about 1 log of the cell viability compared to the textile functionalized only with the cytotoxic PEI layer. Increasing the number of PSS layers – and thus of PSS/TiO₂ bilayers – led to an increase in the E. coli viability in the dark, with a complete viability restoration achieved already for 5 bilayers PSS/TiO₂ (Fig. 4a-F).

By contrast, masking the cytotoxicity of the first internal PEI layer by applying additional PSS/TiO₂ bilayers, allowed to evidence the antibacterial effect resulting under solar light solely from the photocatalytic TiO₂ layers. Indeed, an increase in the antimicrobial effect under solar light was dependent on the number of PSS/TiO₂ bilayers applied. 9.7 × 10⁵, 1.9 × 10⁵ and 9.2 × 10³ CFU per textile were measured after 60 min test under solar light on the textiles functionalized with 1 (Fig. 4a-E), 5 (Fig. 4a-F) and 10 (Fig. 4a-G) bilayers of PSS/TiO₂, respectively, so that the E. coli viability was progressively reduced by increasing the number of bilayers, and that a antibacterial photocatalytic activity close to 3 log was achieved for 10 PSS/TiO₂ bilayers.

This behavior with increasing the number of PSS/TiO₂ bilayers was confirmed by studying the antibacterial activity of the PEI/(PSS/TiO₂)_n multilayer system against P. aeruginosa bacteria (Fig. 4b). A single layer of PEI induced more than 2 log reduction whether the textile was irradiated or not (Fig. 4b-B). Addition of a single layer of PSS on the first layer of PEI (Fig. 4b-C) allowed the partial restoration of the cell viability (7 × 10⁵ CFU per textile vs. 2.6 × 10⁴ CFU per textile on the PES textile with one PEI layer). The higher restoration of cell viability was then obtained when applying one bilayer of PSS/TiO₂ (Fig. 4b-E), i.e. 1.5 × 10⁶ CFU per textile vs. 7.3 × 10⁶ CFU per textile on the naked textile. Increasing further the number of PSS/TiO₂ bilayers applied resulted in a progressive decrease of the cell viability on the solar light irradiated textiles, and 1.4 × 10⁵, 2.3 × 10³ and 1.0 × 10¹ CFU per textile were measured on the textiles functionalized with 1 (Fig. 4b-E), 5 (Fig. 4b-F) and 10 (Fig. 4b-G) bilayers of PSS/TiO₂, respectively, after 60 min of irradiation, corresponding to antibacterial activities close to 2, 4 and 6 log, respectively. So, similarly to the case of E. coli, the cytotoxicity of the first internal PEI layer could be masked by applying additional PSS/TiO₂ bilayers, so that the antibacterial effect resulting solely from the solar light active photocatalytic TiO₂ layers – directly dependent on the number of PSS/TiO₂ bilayers – was evidenced.

The results obtained against E. coli and P. aeruginosa bacteria demonstrated the solar light photocatalytic antibacterial efficiency of the textiles functionalized with PSS/TiO₂ bilayers. In the case of E. coli, this antibacterial activity could thus in a first approximation correspond to the sole photocatalytic anti-bacterial activity of PES textile functionalized with PEI/TiO₂ bilayers, so that the PEI/(PSS/TiO₂)_n multilayer system was suitable for presenting the kinetic of the photocatalytic self-disinfection of functionalized textiles against E. coli bacteria (Fig. 5).

Fig. 5 (a and b) Kinetic of the antibacterial effect against E. coli bacteria under solar light of the multilayered PES–KOH/PEI/(PSS/TiO₂)₁₀ textile (G). (A) Corresponds to control performed over the bare PES–KOH textile. Kinetic has been determined in the dark as well.

4. Conclusions

Solar light active antibacterial textiles have been prepared by functionalization of polyester textiles with sprayed layer-by-layer assembly of TiO₂ photocatalyst using PEI polyelectrolyte as polycation layer in alternation with a dispersion of negatively-charged TiO₂ particles, or PSS polyelectrolyte as polyanion layer in alternation with a dispersion of positively-charged TiO₂ particles.

The high self-decontaminating activity of (PEI/TiO₂)_n functionalized textiles was assigned to a synergistic effect within the multilayer system resulting from the association of both biocidal photocatalytic activity of TiO₂ layers and biocidal activity of PEI polyelectrolyte layers. The application of a single layer of TiO₂ at 50 μg cm⁻² to get 5 log of cell viability reduction turns out to be very promising for targeting a realistic application as self-disinfecting textiles.

PEI/(PSS/TiO₂)_n functionalized textiles were of high interest for evidencing the sole solar light photocatalytic antibacterial activity of multilayered textiles by masking the biocidal activity of PEI.

Acknowledgements

The authors are grateful to DGA (Direction Générale de l'Armement) and Alsace regional council for financially supporting this work in the frame of the PhD grant of G. Carré. The authors acknowledge the support of the French Agence Nationale de la Recherche (ANR) under reference ANR-09-SECU-10. We thank Th. Romero for performing SEM analysis.

Notes and references

1. J. Szostak-Kotowa, Int. Biodeterior. Biodegrad., 2004, 53, 165.
2. C. Callewaert, E. De Maeseneire, F. M. Kerckhof, A. Verliefde, T. Van de Wiele and N. Boon, Appl. Environ. Microbiol., 2014, 80, 6611.
3. C. Rodriguez, A. Di Cara, F. N. R. Renaud, J. Freney, N. Horvais, R. Borel, E. Puzenat and C. Guillard, Catal. Today, 2014, 230, 41.
4. J. M. Nordstrom, K. A. Reynolds and C. P. Gerba, Am. J. Infect. Control, 2012, 40, 539.
5. S. Fijan and S. Sostar-Turk, Int. J. Environ. Res. Public Health, 2012, 9, 3330.
6. A. Toomey, in Medical and Healthcare Textiles, ed. S. C. Anand, J. F. Kennedy, M. Miraftab and S. Rajendran, Woodhead Publishing, 2010, pp. 357–367.
7. A. Mitchell, M. Spencer and C. Edmiston, J. Hosp. Infect., 2015 DOI:10.1016/j.jhin.2015.02.017.
8. G. Borkow and J. Gabbay, Med. Hypotheses, 2008, 70, 990.
9. A. N. Neely, J. Burn Care Rehabil., 2000, 21, 523.
10. A. N. Neely and M. P. Maley, J. Clin. Microbiol., 2000, 38, 724.
11. V. J. Colclasure, T. J. Soderquist, T. Lynch, N. Schubert, D. S. McCormick, E. Urrutia, C. Knickerbocker, D. McCord and J. H. Kavouras, Am. J. Infect. Control, 2015, 43, 154.
12. K. Takai, T. Ohtsuka, Y. Senda, M. Nakao, K. Yamamoto, J. Matsuoka and Y. Hirai, Microbiol. Immunol., 2002, 46, 75.
13. L. Windler, M. Height and B. Nowack, Environ. Int., 2013, 53, 62.
14. F. Piccinno, F. Gottschalk, S. Seeger and B. Nowack, J. Nanopart. Res., 2012, 14, 1109.
15. M. S. Sataev, S. T. Koshkarbaeva, A. B. Tleuova, S. Perni, S. B. Aidarova and P. Prokopovic, Colloids Surf., A, 2014, 442, 146.
16. M. A. Radzig, V. A. Nadtochenko, O. A. Koksharova, J. Kiwi, V. A. Lipasova and I. A. Khmel, Colloids Surf., B, 2013, 102, 300.
17. W.-G. Kwak, M. H. Oh and M.-S. Gong, Carbohydr. Polym., 2015, 115, 317.
18. S. Koechler, J. Farasin, J. Cleiss-Arnold and F. Arsène-Ploetze, Res. Microbiol., 2015 DOI:10.1016/j.resmic.2015.03.008.
19. R. Dastjerdi and M. Montazer, Colloids Surf., B, 2010, 79, 5.
20. J. Bauer, K. Kowal, S. A. M. Tofail and H. Podbielska, in Biological Interactions with Surface Charge in Biomaterials, Royal Society of Chemistry, 2012, p. 193.
21. K. Nakata and A. Fujishima, J. Photochem. Photobiol., C, 2012, 13(3), 169.
22. P. C. Maness, S. Smolinski, D. M. Blake, Z. Huang, E. J. Wolfrum and W. A. Jacoby, Appl. Environ. Microbiol., 1999, 65, 4094.
23. Z. Huang, P.-C. Maness, D. M. Blake, E. J. Wolfrum, S. L. Smolinski and W. A. Jacoby, J. Photochem. Photobiol., A, 2000, 130, 163.
24. J.-M. Herrmann, J. Photochem. Photobiol., A, 2010, 216, 85.
25. M. R. Hoffmann, S. T. Martin, W. Choi and D. W. Bahnemann, Chem. Rev., 1995, 95, 69.
26. G. Gogniat, M. Thyssen, M. Denis, C. Pulgarin and S. Dukan, FEMS Microbiol. Lett., 2006, 258, 18.
27. S. Josset, N. Keller, M. C. Lett, M. J. Ledoux and V. Keller, Chem. Soc. Rev., 2008, 37, 744.
28. D. Wu, M. Long, J. Zhou, W. Cai, X. Zhu, C. Chen and Y. Wu, Surf. Coat. Technol., 2009, 203, 3728.
29. J. Kiwi and C. Pulgarin, Catal. Today, 2010, 151, 2.
30. O. Baghriche, S. Rtimi, C. Pulgarin, C. Roussel and J. Kiwi, Appl. Catal., B, 2013, 130–131, 65.
31. M. Montazer and E. Pakdel, J. Photochem. Photobiol., C, 2011, 12, 293.
32. R. Rahal, M. Le Bechec, R. Guyoneaud, T. Pigot, H. Paolacci and S. Lacombe, Catal. Today, 2013, 209, 134–139.
33. S. Matsuzawa, C. Maneerat, Y. Hayata, T. Hirakawa, N. Negishi and T. Sano, Appl. Catal., B, 2008, 83, 39–45.
34. K. Kowal, P. Cronin, E. Dworniczek, J. Zeglinski, P. Tiernan, M. Wawrzynska, H. Podbielska and S. A. M. Tofail, RSC Adv., 2014, 4, 19945.
35. E. Kasuga, Y. Kawakami, T. Matsumoto, E. Hidaka, K. Oana, N. Ogiwara, D. Yamaki, T. Sakurada and T. Honda, Int. J. Nanomed., 2011, 6, 1937.
36. G. Li, H. Liu, H. Zhao, Y. Gao, J. Wang, H. Jiang and R. I. Boughton, J. Colloid Interface Sci., 2011, 358, 307.
37. S. Rtimi, O. Baghriche, R. Sanjines, C. Pulgarin, M. Bensimon and J. Kiwi, J. Photochem. Photobiol., A, 2013, 256, 52.
38. G. Decher, Science, 1997, 277, 1232.
39. P. Podsiadlo, Z. Tang, B. S. Shim and N. A. Kotov, Nano Lett., 2007, 7, 1224.
40. G. K. Such, A. P. R. Johnston and F. Caruso, in Comprehensive Nanoscience and Technology, ed. D. L. Andrews, G. D. Scholes and G. P. Wiederrecht, Academic Press, Amsterdam, 2011, pp. 359–377.
41. T. Shibata, N. Sakai, K. Fukuda, Y. Ebina and T. Sasaki, Phys. Chem. Chem. Phys., 2007, 9, 2413.
42. D. N. Priya, J. M. Modak and A. M. Raichur, ACS Appl. Mater. Interfaces, 2009, 1, 2684.
43. D. Dontsova, V. Keller, N. Keller, P. Steffanut, O. Felix and G. Decher, Macromol. Rapid Commun., 2011, 32, 1145.
44. S. T. Dubas, P. Kumlangdudsana and P. Potiyaraj, Colloids Surf., A, 2006, 289, 105.
45. Q. Wang and P. J. Hauser, Carbohydr. Polym., 2010, 81, 491.
46. M. Grandcolas, A. Louvet, N. Keller and V. Keller, Angew. Chem., 2009, 48, 161.
47. M. Grandcolas, L. Sinault, F. Mosset, A. Louvet, N. Keller and V. Keller, Appl. Catal., A, 2011, 391, 455.
48. K. C. Krogman, N. S. Zacharia, D. M. Grillo and P. T. Hammond, Chem. Mater., 2008, 20, 1924.
49. S. S. Ugur, M. Sariisik and A. H. Aktas, Nanotechnology, 2010, 21, 325603.
50. B. Ding, J. Kim, E. Kimura and S. Shiratori, Nanotechnology, 2004, 15, 913.
51. N. Beyth, D. Kesler Shvero, N. Zaltsman, Y. Houri-Haddad, I. Abramovitz, M. P. Davidi and E. I. Weiss, PLoS One, 2013, 8, e78586.
52. B. Ohtani, O. O. Prieto-Mahaney, D. Li and R. Abe, J. Photochem. Photobiol., A, 2010, 216, 179.
53. M. Takashima, F. Shirai, M. Sageshima, N. Ikeda, Y. Okamoto and Y. Dohi, Am. J. Infect. Control, 2004, 32, 27.
54. A. L. Barry, W. A. Craig, H. Nadler, L. B. Reller, C. C. Sanders and J. M. Swenson, Methods for Determining Bactericidal Activity of Antimicrobial Agents; Approved Guideline, CLSI, 1999.
55. ISO 27447:2009, Fine ceramics (advanced ceramics, advanced technical ceramics) – Test method for antibacterial activity of semiconducting photocatalytic materials, ISO, Geneva, 2009.
56. O. Félix, Z. Zheng, F. Cousin and G. Decher, C. R. Chim., 2009, 12, 225.
57. D. S. Kommireddy, A. A. Patel, T. G. Shutava, D. K. Mills and Y. M. Lvov, J. Nanosci. Nanotechnol., 2005, 5, 1081.
58. L. Rojas-Blanco, M. D. Urzúa, R. Ramírez-Bon and F. J. Espinoza Beltrán, Appl. Surf. Sci., 2012, 258, 2103.
59. M. A. Pfaller, D. J. Sheehan and J. H. Rex, Clin. Microbiol. Rev., 2004, 17, 268.
60. N. Beyth, Y. Houri-Haddad, L. Baraness-Hadar, I. Yudovin-Farber, A. J. Domb and E. I. Weiss, Biomaterials, 2008, 29, 4157.
61. I. Yudovin-Farber, J. Golenser, N. Beyth, E. I. Weiss and A. J. Domb, J. Nanomater., 2010 DOI:10.1155/2010/826343.
62. J. Lin, S. Qiu, K. Lewis and A. M. Klibanov, Biotechnol. Bioeng., 2003, 83, 168.
63. C. Wiegand, M. Bauer, U. C. Hipler and D. Fischer, Int. J. Pharm., 2013, 456, 165–174.
64. H. Khalil, T. Chen, R. Riffon, R. Wang and Z. Wang, Antimicrob. Agents Chemother., 2008, 52, 1635.
65. E. Kiss, E. T. Heine, K. Hill, Y. C. He, N. Keusgen, C. B. Penzes, D. Schnoller, G. Gyulai, A. Mendrek, H. Keul and M. Moeller, Macromol. Biosci., 2012, 12, 1181.
66. U. Rabuza, S. Sostar-Turk and S. Fijan, Text. Res. J., 2012, 82, 2099.
67. G. Carre, E. Hamon, S. Ennahar, M. Estner, M. C. Lett, P. Horvatovich, J. P. Gies, V. Keller, N. Keller and P. Andre, Appl. Environ. Microbiol., 2014, 80, 2573.
68. A. M. Carmona-Ribeiro and L. D. de Melo Carrasco, Int. J. Mol. Sci., 2013, 14, 9906.
69. I. M. Helander, H. L. Alakomi, K. Latva-Kala and P. Koski, Microbiology, 1997, 143, 3193.
70. S. Brosel-Oliu, N. Abramova, A. Bratov, N. Vigues, J. Mas and F. X. Munoz, Electroanalysis, 2015, 27, 656.
71. K. I. Ishibashi, A. Fujishima, T. Watanabe and K. Hashimoto, J. Phys. Chem. B, 2000, 104, 4934.
72. G. Carre, D. Benhamida, J. Peluso, C. D. Muller, M. C. Lett, J. P. Gies, V. Keller, N. Keller and P. Andre, Photochem. Photobiol. Sci., 2013, 12, 610.

---

Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c5ra05541e

---