**1. Introduction**

Microbial resistance to antimicrobial agents is increasing worldwide and it represents a major threat to the successful treatment of infectious diseases [1]. Development of antimicrobial resistance is an inescapable consequence of natural selection and is associated with exposure to antimicrobial agents [1]. Efforts need to be made to decrease the unnecessary exposure of bacteria to antibiotics to reduce the selective pressure driving the development of resistance so that existing antibiotics retain their efficacy for as long as possible [1]. In part, this can be achieved by controlling the spread of pathogenic bacteria and therefore reducing the number of infections that require antibiotic treatment [2].

Healthcare-associated infections are a major contributor to patient morbidity and mortality, and occur in part due to bacterial contamination of hospital surfaces [3]. Surfaces in hospitals that come into contact with hands are regularly contaminated with nosocomial pathogens [4,5]. Infected patients shed pathogenic bacteria, including methicillin-resistant *Staphylococcus aureus* and vancomycin-resistant *Enterococcus* spp., into their immediate environment [5–10]. Surfaces near shedding patients, such as walls, door handles, bed frames, and light switches, tend to be touched frequently and therefore are more likely to be contaminated [3,10,11]. Once a surface is contaminated, a single hand contact event is sufficient to transmit bacteria from the surface to a person [7,8,12].

Bacteria that have been transferred to a surface can persist for a period of time or actively colonise to form a biofilm. Bacterial persistence on a surface is influenced by dynamic environmental conditions, including organic soiling, humidity, and temperature [4,13,14]. Biofilms are highly recalcitrant to antimicrobial treatments and facilitate the persistence of bacteria on surfaces resulting in surface associated pathogen reservoirs, which increase the risk of transmission [15–17]. The level of bacterial transfer that occurs between a contaminated surface and a hand following contact has been demonstrated to occur at a comparable level to direct contact with an infectious patient, which is a well-established transmission route [4,18,19]. Hand washing can help to control the spread of infection in hospitals; however, without the decontamination of surfaces, the reservoirs of pathogens will seed further spread. Nosocomial pathogens isolated from hospital surfaces are typically in the range of 100–10,000 colony forming units (CFU)/cm<sup>2</sup> [5,10]. For a microbial burden exceeding 250 CFU/100 cm2, transmission from the surfaces to health care workers and/or patients increases [10,20,21]. Therefore, despite the relatively low inocula present, any contamination of a hospital surface by a pathogen should be considered to be a transmission risk [10,20,21].

The involvement of contaminated surfaces in pathogen transmission pathways in hospitals necessitates the improved control of surface microbiology. Reduction of microbial contamination on hospital surfaces could disrupt transmission pathways and potentially reduce infectious disease incidence rates and the associated antibiotic usage [22]. Utilisation of antibacterial surfaces is a promising means of reducing microbial surface load as well as preventing formation of biofilms and surface associated pathogen reservoirs [2]. An ideal antimicrobial surface would be active against relevant bacteria at appropriate bacterial loads and active in environmental conditions relating to potential applications in terms of temperature, relative humidity, pH, exposure to cleaning products, and contaminating organic matter [23–25]. The time that is required for decontamination would need to be sufficiently short to be effective in breaking transmission pathways. Activity needs to be retained for sufficiently long periods of time, and after repeated bacterial challenges, to be cost effective [26,27]. Activity overtime is informed by whether the antimicrobial agen<sup>t</sup> is immobilised on the surface or if it has to be released to elicit an effect [28,29]. The release of an antimicrobial agen<sup>t</sup> over time means that the surface concentration of the agen<sup>t</sup> will fall below the threshold needed to exert antimicrobial activity [27]. An ideal antimicrobial surface would also need to be cheap and easy to make, suitable for large-scale production, and have regulatory approval for the intended use [26,30].

To create an antimicrobial surface, we can take one of two basic approaches. First, a coating may be applied to a material or a modification of the surface chemistry of the material made to provide an antimicrobial surface [31]. Alternatively, the material may be fabricated by incorporating an antimicrobial into the material, which can be challenging as manufacturing procedures can involve extreme environmental conditions, including high temperatures and shear forces, which can negatively impact on bactericidal activity [26,28,32]. Covalent attachment of an antimicrobial agen<sup>t</sup> to a surface may cause side reactions that result in conformational changes in the agent, ultimately causing a loss of activity [33]. Therefore, the method of antimicrobial surface production may affect the resulting surface activity.

The activity of an antimicrobial surface is also influenced by the nature of the surface. Surfaces can be absorbent allowing water droplets to move into the surface or they can be non-absorbent, in which water droplets sit on top of the surface [34,35]. These surface properties may affect the antimicrobial activity as a bacterium in a water droplet would have more contact with the antimicrobial agen<sup>t</sup> if it has absorbed into the surface. Non-absorbent surfaces in hospitals are frequently contaminated with pathogens, and include walls, door handles, and bed frames. Much of the focus of development of antimicrobial surfaces in the published literature is on model non-absorbent surfaces, such as metal coupons and plastic films [36–38]. Many absorbent surfaces in hospitals are fabric-based, such as apparel worn by healthcare workers and patient privacy curtains [36,39,40]. Privacy curtains are high-touch areas that are contacted by the hands of the healthcare worker before, during, and after patient care, and are infrequently changed [40,41]. It has been demonstrated that more than 90% of privacy curtains can become contaminated within a week of use [41]. Contaminated absorbent surfaces in hospitals may be involved in pathogen transmission [36,39]. Absorbent surfaces are harder to clean or disinfect than non-absorbent surfaces, while the latter facilitates a greater transfer of bacteria [8,41,42]. Therefore, the development of both absorbent and non-absorbent antimicrobial surfaces would help to curtail the spread of infection in hospitals [1].

Antimicrobial polymers are good candidates for immobilised biocides. These polymers can be either polymeric biocides (the repeating unit is a biocide) or biocidal polymers (the active principle is embodied by the whole macromolecule) [28,29]. In this article, we investigate the antimicrobial activity of polyaniline (PANI) and a functionalised derivative (fPANI), homopolymer poly(3-aminobenzoic acid) (P3ABA), as surface-immobilised biocidal polymers. Utilisation of PANI for potential applications is restricted because of its insolubility in common solvents, which renders it difficult to process [43,44]. fPANIs are easily and inexpensively synthesised using substituted aniline monomers, which improves the solubility, and thus the processability, of the resulting polymer [43,44]. PANI and P3ABA are good candidates for incorporation into surfaces because they have thermal stability up to 300 ◦C, environmental stability in the conducting form, simple and inexpensive synthetic procedures [45–48], and have been demonstrated to be biocompatible with mammalian cells [49–52], all of which increases their commercial viability. Surfaces containing PANI and P3ABA are non-leaching [45,53,54], which promotes activity over a longer period of time and reduces both personal and environmental safety concerns [27].

In this study, we investigated the potential of PANI and P3ABA as surface antimicrobial agents. Initial testing involved the challenge of target organisms in suspension, mirroring standard antimicrobial susceptibility testing methods [55,56]. The target organisms selected were the antimicrobial susceptibility testing strains, *Escherichia coli* 25922 and *S. aureus* 6538, which reflect bacteria that are commonly isolated from surfaces in hospitals [10,57]. Susceptibility to antimicrobial activity can be influenced by media composition through its effects on bacterial cell physiology [58]. Therefore, *E. coli* was challenged in Lennox Broth (LB)—a rich media on which cells grow at high rates—and in minimal A salts with 0.4% succinate as the carbon source, which only contains nutrients that are essential for growth [59–61]. The slow growth of bacteria in a minimal media environment is similar to what may occur on surfaces in nature [62].

Following confirmation of activity in suspension, PANI and P3ABA were incorporated into absorbent and non-absorbent surfaces. The effect of incorporation on antimicrobial activity was determined in 96 well plate based assays, which allowed for the the testing of many concentrations and treatment times against one inoculum [27]. Absorbent surfaces were modelled using agar mixed with varying amounts of PANI or P3ABA [63]. Drops of liquid containing bacteria absorbed into the solidified agar test surfaces [64]. Non-absorbent surfaces were established in the form of compression moulded Styrene Ethylene Butylene Styrene (SEBS) films [63]. The activity of non-absorbent surfaces containing PANI and P3ABA were then characterised in relevant environmental conditions, including challenging with a range of inocula and in the presence of organic matter [62]. The experimental strategy is summarised in Figure 1, and, taken together, the results presented demonstrate the activity of PANI and P3ABA in suspension and in surfaces, in application relevant settings. The efficacious activity of P3ABA supports the utilisation of this polymer to create contamination resistant surfaces.

**Figure 1.** Experimental strategy. Polyaniline (PANI) and functionalised derivative (fPANI) are tested according to the scheme presented. 1 Activity is measured as reduction in the number of viable cells recovered from surfaces. 2 The Gram positive bacterium *S. aureus*, and the Gram-negative bacterium *E. coli* were tested as model species.
