Enhanced Clearing of Wound Related Pathogenic Bacterial Biofilms Using Protease-Functionalized Antibiotic Nanocarriers

: Biofilms are prevalent in chronic wounds and once formed are very hard to remove which is associated with poor outcomes and high mortality rates. Biofilms are comprised of surface-attached bacteria embedded in an extracellular polymeric substance (EPS) matrix, which confers increased antibiotic resistance and host immune evasion. Therefore, disruption of this matrix is essential to tackle the biofilm embedded bacteria. Here we propose a novel nanotechnology to do this, based on protease-functionalized nanogel carriers of antibiotics. Such active antibiotic nanocarriers, surface coated with the protease Alcalase 2.4 L FG, "digest" their way through the biofilm EPS matrix, reach the buried bacteria and deliver a high dose of antibiotic directly on their cell walls which overwhelms their defenses. We demonstrated their effectiveness against six wound biofilm forming bacteria, Staphylococcus aureus, Pseudomonas aeruginosa, Staphylococcus epidermidis, Klebsiella pneumoniae, Escherichia coli and Enterococcus faecalis . We confirmed a six-fold decrease in the biofilm mass, and a substantial reduction in bacterial cell density using fluorescence, atomic force and scanning electron microscopy. Additionally, we showed that co-treatments of ciprofloxacin and Alcalase-coated Carbopol nanogels led to a 3-log reduction in viable biofilm forming cells when compared to ciprofloxacin treatments alone. Encapsulating an equivalent concentration of ciprofloxacin into the Alcalse-coated nanogel particles boosted their antibacterial effect much further, reducing the bacterial cell viability to below detectable amounts after 6 hours of treatment. The Alcalase-coated nanogel particles were non-cytotoxic to human adult keratinocyte cells (HaCaT), inducing a very low apoptotic response in these cells. Overall, we demonstrated that the Alcalase-coated nanogels loaded with a cationic antibiotic elicit very strong biofilm clearing effects against wound associated biofilm forming pathogenic bacteria. This nanotechnology approach has the potential to become a very powerful treatment of chronically infected wounds with biofilm forming bacteria.


INTRODUCTION
Biofilms are microbial communities surrounded by a structure referred to as extracellular polymeric substance (EPS), composed of biopolymers such as exopolysaccharides, nucleic acids, lipids, and proteins. 1 The EPS matrix is highly hydrated, with occasional hydrophobic regions, such as areas containing cellulose. 2 Combined, these substances form a structured matrix, allowing bacteria to effectively adhere to a range of surfaces. 3 The formation of biofilms begins when surface-attached bacterial cells secrete EPS which allows complex communal tertiary microcolony structures to form. 4 Biofilms are commonly surface-attached with one layer of cells in direct contact with the substratum, however, they can be found in flocs in which biofilms are formed around cells without contact with a substratum. 5 The EPS matrix provides effective protection of the bacteria within the biofilm to a wide array of environmental factors, such as antimicrobial agents and a host immune response molecules. 6,7 Biofilms confer benefits to the cells within, in quenching antimicrobial agents which would kill planktonic cells. 8 The biofilm is able to impair the diffusion of antimicrobial agents through the EPS and lower local concentration, thus increasing the tolerance of the cells to the most chemicals. 9 An example of tolerance is the neutralization of toxic metal ions such as copper which is complexed to the polysaccharides of the EPS. 10,11 Biofilm infections in humans with species which show high resistance to treatment may result in chronic infections. 16 Opportunistic pathogens such as, Staphylococcus, Pseudomonas and Klebsiella genus', can form biofilms on acute and chronic dermal wounds, impairing wound healing, and potentially leading to sepsis. 17 This effect can lead to increased patient mortality and morbidity in regard to biofilm infection. 18,19 Nanoparticles (NPs) have been investigated for their anti-biofilm properties. Ag-Au nanocomposite particles, CuONPs, Mg(OH)2 NPs, silica NPs This document is the Accepted Manuscript version of a Published Work that appeared in final form in ACS applied materials & interfaces, copyright © American Chemical Society after peer review and technical editing by the publisher. To access the final edited and published work see https://doi.org/10.1021/acsami.9b16119 and Au NPs have shown high efficacy in this regard. 21,22,12,64,13 Borovicka et al. 37,45,46 developed photothermal colloid antibodies based on bioimprints that can selectively kill microbial cells by cell shape recognition. Au NPs in conjunction with heat produced with laser irradiation can effectively disrupt biofilms. 23 Taking advantage of the antimicrobial effect of Ag has led to the novel use of bioglass which incorporates surface reacted Ag and has been shown to have effective anti-biofilm use. 24 Enzymatic biofilm degradation has also been investigated, e.g. the use of hydrolases, amylases, and proteases. 25 The principle of disrupting the biofilm network using enzymatic lysis, rather than directly attacking the cells themselves has the advantage of not relying on the biocidal action of tradition antimicrobials, and as such do not place a direct evolutionary pressure on the bacteria to develop resistance. 26 All components of the EPS matrix have been investigated for the removal of biofilms, these include DNases, glycosidases, and proteases. 27 An examples of a glycosidases is dispersin B produced by Aggregatibacter actinomycetemcomitans, has been demonstrated as being effective against S. epidermidis and S. aureus biofilms. 28,29 Glycoside hydrolase alginate lyase and DNase NucB has also proved effective against P. aeruginosa and E. coli biofilms respectively. 30,31 Proteases (EC 3.4) are present in all life forms, providing a wide variety of metabolic and physiological functions, including signal transduction, apoptosis, cell division, blood pressure regulation, and many more. 32 Protease have previously been investigated for their anti-biofilm potential, both as small molecules (e.g. ficin) and bacterial-derived Bacillus proteases (AprBp and GseBp). 33,34 Ironically, bacteria have become a useful source of anti-biofilm proteases. 35,36 Proteases have two unique properties which make them an excellent enzyme choice for degrading biofilms. Their ability to hydrolyze EPS matrix proteins and adhesins (allowing a biofilm to attach to a solid surface and bacteria-bacteria adhesion) give them a distinct advantage over other anti-biofilm enzymes. 1 Additionally, they may disrupt intercellular communication by lysis of type I signal peptidase, as shown in S. aureus. 39 Proteases have also been shown to have potential in improving wound healing by degrading the biofilm EPS which may be interfering with the normal wound healing process. 40,41 Serine Esp based protease obtained from S. epidermidis and elastase LasB from P. aeruginosa have been shown to reduce biofilm formation in S. aureus, providing further evidence of their efficacy and usefulness in biofilm treatment options. 42 In addition to the evidence that exogenous protease degrade biofilms, it has been shown that in the natural selfinduced disassembly of biofilms, bacteria induce expression of extracellular proteases to cleave matrix proteins. 44 Some of these extracellular proteases include sigB and sarA, which have all been shown to reduce biofilm capacity in S. aureus. 47 Novozymes® (Denmark) produce a wide array of enzymes produced by Bacillus spp. (Gram-positive, rodshaped, spore-forming bacteria) which have applications in industry and biopharmaceutical ingredients, many with useful activity properties as mentioned above. 38 It has been demonstrated that immobilization of proteases can increase their stability, reusability, and increase their tolerance of environmental stresses. 48 Covalent bonding, ionic interactions, entrapment/encapsulation in a polymer or inorganic matrices and hydrophobic adsorption have previously been demonstrated as methods to immobilize enzymes. 49 Although immobilization can improve the enzyme activity, stability and storage life, there is also the possibility of enzyme rigidification, particularly in strong covalent techniques, which reduce the effective activity of the enzyme. 50 Nanoparticle carriers such as chitosan beads and soybean fibers have been shown to be useful biocompatible carriers for proteins and also increase the activity of Alcalase. 51,52 In this study, we investigated the use of Carbopol Aqua SF1 nanogels as a substrate to immobilize a Serine endo-peptidase proteases, namely Alcalase 2.4 L FG (Figure 1 illustrates the process). Polyacrilic acid co-polymer nanogels have cross-linked polymer chains which due to the abundance of COOH groups confer an anionic charge upon their surface and interior. 53 These nanogel particles have shown good biocompatibility and have been found to be effective in encapsulating cationic antimicrobials such as chlorhexidine digluconate, and tetracycline and lincomycin hydrochloride. 54,55 Nanocarrier-based antibiotic delivery provides an opportunity to increase the effectiveness of therapeutic agents by specifically targeting bacteria due to their anionic cell wall 54,55 (e.g. utilizing cationic coatings of the nanogel such as PDAC and bPEI), localizing the drug whilst reducing the unintended damaging effect on neighboring tissue. Since Alcalase 2.4 L FG has a very high isoelectric point, it is positively charged at neutral pH. We took advantage of the negative surface charge of Carbopol Aqua SF1 nanogel particles to electrostatically immobilize Alcalase 2.4 L FG on their surface. We investigated the stability of Alcalase 2.4 L FG coated Carbopol Aqua SF1 nanoparticles, and evaluated their potential anti-biofilm use on skin associated pathogens which are known to produce biofilms. Additionally, we investigated the potential synergistic effect of antibiotic co-treatments and encapsulations within the nanogel using ciprofloxacin hydrochloride, combined with further surface coating of the loaded nanogel with Alcalase 2.4 L FG. Our study demonstrates that there is a strong synergistic clearing effect of these novel dual functionalized nanogels on biofilms of a range of pathogenic bacteria. . Millipore Express PLUS Membrane Filter paper (0.22 µm pore size, hydrophilic polyethersulfone, 47 mm diameter), Ciprofloxacin hydrochloride (≥98.0%), Acridine Orange base (75%) and Resazurin Sodium salt (75%) were purchased from Sigma-Aldrich, UK. Resazurin Sodium salt was used for cell metabolic in situ assays and diluted in Dulbecco's Phosphate Buffered Saline (DPBS) (Gibco, Fisher Scientific, UK) at a concentration of 0.015 wt/vol%. HaCaT cells were obtained from AddexBio, T0020001, USA. DMEM and FBS were obtained from Gibco, UK, L-Glutamine and Trypsin EDTA by BioWhittaker, UK. A Pierce TM Protease Assay Kit (Thermo Scientific, USA) was used to characterize the Alcalase 2.4 L FG activity. A MTT colorimetric survival and proliferation kit (Millipore Corp, USA) was used for Ha-CaT cell viability experiments and ANNEXIN V apoptosis detection Kit I (BD Pharmogen, USA) was used to measure apoptosis after treatments. Deionized water purified by reverse osmosis and ion exchange with a Milli-Q water system (Millipore, USA) was used in all our studies. Its surface tension was 71.9 mN m -1 at 25 °C, with measured resistivity higher than 18 MΩ cm -1 . Consumable plasticware used in the study were purchased from Sarstedt (UK), Thermo Scientific (UK), or CytoOne (UK) unless otherwise stated. Synthesis of Alcalase-coated nanogel particles. Carbopol Aqua SF1 (30 wt%) was diluted into 0.6 wt% by pipetting 2 mL of the stock suspension (30 wt%) and diluting with deionized water to a final volume of 100 mL. This suspension was sonicated for 15 min using 2 second on/off pulsing to allow effective dispersion. Droplets 0.25 M HCl was added to the suspension to reduce the pH to 5.5. A 0.6 wt% solution of Alcalase L FG 2.4 was created by diluting 0.6 mL of the stock liquid enzyme solution and diluting with deionized water to a final volume of 100 mL. The 0.6 wt% Alcalase solution was sonicated for 15 min to prevent aggregation. 25 mL of the 0.6 wt% Carbopol Aqua SF1 suspension and 25 mL of the 0.6 wt% Alcalase L FG 2.4 solution were mixed together for 30 min at pH 5.5 with constant stirring to allow the cationic enzyme to electrostatically bind to the anionic Carbopol NPs. After mixing the suspension was centrifuged at 4000 rpm for 15 min the pellet was washed trice with deionized water and then dispersed into 50 mL of fresh deionized water. Droplets of acetate buffer solution were used to maintain the dispersion at 5.5 pH. The supernatant was retained for analysis to determine the surface functionalization efficiency by UV-Vis absorption measurement. The particle size and zeta potential distribution of the Carbopol-Alcalase NPs were measured using a Malvern Zetasizer as described below. Prior to measurement, and use of the nanogels in treatments, the 0.6 wt% Carbopol-0.6 wt% Alcalase dispersion was sonicated for 5 min to remove aggregation and diluted into deionized water to the appropriate concentration. Preparation of ciprofloxacin-loaded nanogel. The principle of encapsulating a cationic antibiotic (as a hydrochloride salt) is based on the swelling and deswelling cycle of the Carbopol Aqua SF1 at different pH. 53 Briefly, a 100 mL of 0.6 wt% aqueous dispersion of the nanogel was prepared, this was then adjusted to pH 7.5 by adding droplets of 0.25 M NaOH whilst been stirred. The dispersion was then warmed to 37 °C. An aliquot of 0.0032 wt% ciprofloxacin hydrochloride aqueous dispersion was prepared by weighing 3.2 mg of the antibiotic (ABX = ciprofloxacin) powder, diluting into 100 mL of deionized water and then warming to 37 °C. The 100 mL ABX solution was then added to the pH 7.5 nanogel dispersion and shaken for 30 min at 37 °C to allow the antibiotic cations to diffuse into and electrostatically bind in the cores of the swollen nanogel particles. The pH of the ABX-Carbopol solution was then reduced to pH 5.5 using droplets of 0.25 M HCl whilst being stirred for another 30 min. The ABX-Carbopol suspension was then centrifuged at 4000 rpm for 15 min, and the supernatant was removed and retained for encapsulation efficiency analysis. The pellet was washed twice with deionized water and re-dispersed into 100 mL of deionized water. The pH was then increased to 7.5 by gradually adding droplets of 0.25 M NaOH and the solution was gently stirred overnight. The final ABX-Carbopol nanogel solution was reduced to pH 5.5 using acetate buffer solution. The particle size and zeta potential distribution of the ATX-Carbopol dispersion were measured using a Malvern Zetasizer Nano ZS as described above. Coating of the ciprofloxacin encapsulated Carbopol NPs with Alcalase 2.4 L FG was performed as described above. Characterization of free Alcalase, Alcalase-nanogel and Alcalase-coated Ciprofloxacin-loaded nanogel. All hydrodynamic diameter and zeta potential measurements were performed using a Malvern Zetasizer Nano ZS. The isoelectric point of Alcalase L FG 2.4; 10 mL of 0.02 wt% aqueous solution Alcalase aliquots were created at a range of pH 5 to 12 using droplets of either 0.25 M HCl or 0.25 M NaOH. Afterwards, the aliquots were sonicated for 15 min. 1 mL of each aliquot was added to a quartz cuvette and the zeta potential measured using a ZEN1002 dip cell at a refractive in-dex of 1.45 and absorption of 0.001 as per Malvern Instruments protein refractive index manual. Measurements were performed at 25 o C and data represented as the mean of 3 repeats. Hydrodynamic diameter and zeta potential of Alcalasenanogel and Alcalase-nanogel loaded with Ciprofloxacin. 50 mL of pH 5.5 0.6 wt% Carbopol-0.6 wt% Alcalase L FG 2.4 NPs and 0.6 wt% Carbopol-0.0032 wt% ciprofloxacin-0.6 wt% Alcalase-nanogel dispersions were prepared as described above. Aliquots of 0.6 wt% Carbopol were mixed with varying concentrations of Alcalase L FG 2.4 (range; 0.000625 wt% to 0.6 wt%). The hydrodynamic diameter and zeta potential were measured at 25 o C using a quartz cuvette at a RI of 1.336 and absorbance of 1.000 as determined by Al-Awady et al. 53 Each measurement was repeated 3 times and data presented as the mean. The UV-Visible spectrum of 0.05 wt% Carbopol, 0.05wt% Alcalase 2.5 L FG and 0.05 wt% Carbopol with 0.05 wt% Alcalase 2.4 L FG coating. Samples were measured at pH 5.5 by adjustment with acetate buffer solution. The absorbance was measured between 220 nm and 1000 nm using a FLUOstar Omega spectrophotometer, BMP Labtech. The Alcalase coating efficiency of the Carbopol NPs was examined by UV-Vis analysis of the supernatant produced during the synthesis as described above. The absorbance of the supernatant was measured at 500 nm using a spectrophotometer (FLUOstar Omega spectrophotometer, BMP Labtech). Protease activity of free Alcalase vs. nanogel-immobilized Alcalase. The activity of free Alcalase 2.4 L FG vs Alcalase 2.4 L FG-Carbopol Aqua SF1 NPs was investigated by measuring the cleavage of succinylated casein in the presence of 0.2 wt% Alcalase, or 0.2 wt% Alcalase-0.2 wt% Carbopol NPs at varying pH, temperature, and over time. 10 mg of lyophilized succinylated casein was dissolved in 1 mL of acetate buffer (pH 5.5), PBS (pH 7.5), or BupH Borate buffer (pH 8.5). For the purpose of the temperature and time experiments acetate buffer was used for all measurements, mimicking the buffer used in the NP preparation. 100 μL of this solution was added to a 96-well plate, and 100 μL of buffer was added to bring the total volume to 200 μL in each well. 100 μL of plane buffer was used as a blank. 50 μL of either free Alcalase 2.4 L FG vs. Alcalase 2.4 L FG-Carbopol Aqua SF1 NPs was added to the wells. The activity of the samples was measured at different pH (using buffers above) for 30 min at room temperature, over time (30 mins to 24 hr) at room temperature, or at different temperatures (30 o C, 37 o C, and 45 o C) for 30 min. 100 μL of trinitrobenzenesulfonic acid was dissolved in 14.9 mL of appropriate buffer to create a TNBSA working solution. Succinylated casein in the presence of proteases is cleaved at the peptide bonds revealing primary amine groups. TNBSA reacts with these groups to produce an orange-yellow product of which the absorbance can be measured at 450 nm informing of protease activity of the samples. The protease activity of the free protease vs nanogel-coated protease were measured at specific time points by adding 50 μL of the TNBSA solution to the wells and incubating for 20 min. Then, the absorbance of the wells was measured at 450 nm subtracting the absorbance from the blank wells. This data was used to assess the relative protease activity of the free Alcalase 2.4 L FG vs Alcalase 2.4 L FG-Carbopol Aqua SF1 nanogels at different experimental conditions. Bacterial culture and biofilm producing assessment. Frozen ATCC species were prepared onto MHA plates according to manufacturer's instructions. Overnight (O/N) cultures were prepared by incubating a single colony scraped from the MHA stock plates into 10 mL of MHB for 16 hr at 37 o C with 140 rpm shaking (Stuart Orbital Incubator S1500). For biofilm assays, O/N cultures were adjusted to 0.5 McFarland standard by diluting the O/N culture into 0.85 w/v% sterile saline until an optical density of 0.08-0.12 at 625 nm (1-2×10 8 CFU/mL) was obtained using a spectrophotometer (FLUOstar Omega spectrophotometer, BMP Labtech). These adjusted bacterial saline suspensions were then diluted 1:150 into MHB to yield starting concentrations between 5×10 5 -1×10 6 colony forming units per mL (CFU mL -1 ). All 6 ATCC species used in this study have previously been confirmed as biofilm producers, however, to validate this, a simple qualitative biofilm formation assessment was performed. 2 mL of MHB was placed into glass test tubes and inoculated with a single colony of bacteria from culture plates. The test tubes where incubated for 24 hr at 37 °C. Afterward, the culture media was decanted and the test tubes washed with PBS (pH 7.4), and left to dry for 30 min. 0.1 wt% Crystal Violet was used to stain cells adhered to the glass tubes for 5 min, before being washed away with deionized water. The test tubes where then left to dry in an inverted position for 2 hr. Biofilm formation was considered positive when a visible film lined the wall and the bottom of the tube. Ring formation at the liquid interface was not indicative of biofilm formation. The formation of biofilm by staining of the adhered cells was scored as either 0-absent, 1-weak, 2-moderate or 3-strong, as performed by Christensen et al. 56 Zeta potential of bacteria treated with free Alcalase and Alcalase-coated nanogel. Bacterial species were cultured as described in as described above. The bacterial suspension was then centrifuged for 10 min at 5000 rpm and the supernatant discarded. The pellet was washed twice with deionized water and finally resuspended into 10 mL of 0.2 wt% Alcalase or 0.2 wt% Alcalase-0.2 wt% Carbopol Aqua SF1 in deionized water. The samples were incubated at room temperature with gentle mixing for 10 min or 1 hrs. Afterwards, a 1 mL aliquot of this suspension was added to a quartz cuvette and the mean zeta potential distribution was measured using the Dipstick probe and a Malvern Zetasizer Nano ZS (Malvern Instruments, UK). The refractive index was 1.384, the absorption 1.000, and the temperature was 25 o C for all measurements. Biofilm staining with Crystal Violet and Congo Red. A standard 96-well microtiter biofilm formation plate assay was performed using crystal violet and Congo Red dyes (modified from O'Toole, 2011). 57 Crystal Violet staining: 1 mL of 5×10 5 -1×10 6 CFU/mL of O/N bacterial culture was pipetted into 48-well TC-treated plates and incubated at 37 o C for 24 hrs in static conditions. After incubation, the plates were washed twice by submerging in deionized water to remove any remaining media and suspended cells. The plate was then shaken dry and left to air dry for 15 mins at room temperature. 1 mL of MHB supplemented with Carbopol-Alcalase synthesized NPs treatment was added to the wells and incubated for 24 hrs at 37 o C in static conditions. The plates were then washed vigorously by submersion in deionized water to remove excess treated media and suspended cells. The plate was then shaken to remove any liquid droplets and left to dry for 15 mins at room temperature. 1 mL of 0.1 wt% Crystal Violet solution was added to each well and incubated at room temperature for 15 mins. The plates were then washed thoroughly by submersion in deionized water and blotted with paper towels to remove excess dye and water. The plate was left to dry for 2 hrs at room temperature. 1 mL of 30 wt% acetic acid was added to each well for 15 min to solubilize to the Crystal Violet. 125 μL of this solution was transferred to a fresh 96well plate and the absorbance measured at 570 nm with 30 wt% acetic acid used as a blank measurement. Congo Red staining: As described above, 48-well plate biofilms were formed and treated with 1 mL of MHB supplemented Carbopol-Alcalase nanogel suspension. After the treatment, the enzyme-infused MHB media was removed and the 48well plate washed thoroughly by submerging in water. The plate was then shaken to remove liquid droplets and left to dry for 15 mins at room temperature. 125 μL of 0.005 wt% Congo Red was added to each well and left to stain the amyloid protein complexes for 15 mins. The 96-well plate was then washed thoroughly again by submersion in deionized water and blotted with paper towels to remove excess Congo red dye and water. The plate was left to dry for 2 hrs at room temperature. 125 μL of 30% ethanol was added to each well for 15 mins to solubilize to the Congo Red dye. 125 μL of this solution was transferred to a fresh 96-well plate and the absorbance measured at 550 nm with 30% ethanol used as a blank sample. Membrane Colony Biofilm NP and Ciprofloxacin treatment. To test the effects of Alcalase-Carbopol nanogel and ciprofloxacin co-treatments on biofilms a filter membrane method was employed. 2 cm 2 Millipore Express PLUS Membrane Filter paper sections were placed on to MHA. An overnight culture was adjusted to 1×10 5 CFU/mL in saline and 20 μL of this culture added as a droplet to the center of the filter paper. The sample was then incubated for 24 hours at 37 o C to allow a colony biofilm to form. After 24 hrs the membranes were placed onto fresh agar plates (3mm deep) into a 6-well plate and 1 mL of treatment added to each well. The plate was gently shaken in the incubator during treatment. 1 mL of deionized water was considered the growth control. 0.6 wt% Carbopol, 0.6 wt% Alcalase 2.4 L FG in deionized water, and 0.6 wt% Carbopol-0.6 wt% Alcalase 2.4 L FG NPs were used as treatments to the measure their effect on the cell viability. Additionally, ciprofloxacin hydrochloride was added separately and as a co-treatment with the Carbopol-Alcalase NPs at a range of 0 to 32 μg/mL (0.0032 wt%). Equivalent concentrations of encapsulated ciprofloxacin tested as a comparison. After 24 hrs of treatment, the membrane filter paper was peeled from the agar wells and placed in to sample tube with 1 mL of fresh MHB and 2 cm of sterile glass beads. Each sample was vortexed for 30 secs at high speed to disintegrate the biofilm and inoculated the MHB with cells. The drop plate enumeration method was used to measure cell viability in CFU/mL. To measure the viability of cells within the biofilms, 10-fold dilutions were made in fresh MHB, 10 μL droplets were placed on to MHA plates and grown for 24 hrs at 37 o C. CFUs were counted from the last two droplets which contained a countable amount of colonies (3 to 30 colonies per 10 μL drop) and averaged. Resazurin Sodium Salt in situ bacterial metabolic activity assay. After the 24-hr membrane colony biofilm NP treatment, as described above, each membrane biofilm was carefully removed from the agar plate using sterile tweezers at the corner of the membrane. The membrane biofilm was allowed to air dry in sterile conditions for 15 min and then placed onto a dry sterile Petri dish. 20 μL of 0.015 wt% Resazurin Sodium Salt was added and incubated for 30 min at 37 o C to visualize the metabolically active cells within the biofilm (modified from Wilkinson et al. 58 ). Images were captured using an 8MP digital camera and the percentage of cell metabolic activity calculated using ImageJ v1.52a. Biofilm formation assay on a glass surface. An overnight culture of each species was adjusted to 0.5 McFarland standard. 10 μL of the adjusted culture was placed into a Petri plate containing a glass slide and 10 mL of MHB and incubated for 48 hrs at 37 o C allowing the bacterial species to form a glass slide adhered biofilm. After 24 hrs the glass slide was washed with deionized water to remove any remaining planktonic cells, and the media was replaced with 10 mL of fresh MHB supplemented with 0.6 wt% Carbopol, 0.6 wt% Alcalase L FG, and 0.6 wt% Carbopol-0.6 wt% Alcalase 2.4 L FG. Un-supplemented media was considered the control. The glass biofilms were incubated for a further 24 hrs at 37 o C. Post treatment, the glass slides were washed with deionized water. 100 μL 0.2 wt% Acridine Orange (AO) was added to the Petri dish and incubated at room temperature for 5 mins in darkness with gentle shaking to reduce photobleaching. After incubation, the slides were washed three times with deionized water and allowed to air dry for 15 mins. The slides were examined using an Olympus BX51 fluorescent microscope with DP70 digital camera. Atomic force microscopy (AFM). AFM images of the airdried biofilms were obtained using a Dimension Icon scanning probe microscope (Bruker, USA) using tapping mode. The biofilms were prepared and treated as described above in the glass surface biofilm formation assay. Post treatment, the glass slide biofilms were washed gently with deionized water, and the biofilm fixed in 1 wt% aqueous glutaraldehyde for 2 hrs by submersion. After fixation, the slide was washed three times with deionized water and left to air dry inside an enclosed container. TESPA-V2 probes (Bruker, USA) with a nominal length of 127 μm and a tip radius of 7 nm were used, the images obtained at 512 lines/scan at a 0.25 Hz scan rate. Images were acquired in height (topography) and the analyzed using Nanoscope Analysis v.1.7. (Bruker, USA). TEM images of Alcalase-nanogel and SEM images of the treated biofilms. SEM images of membrane colony biofilms were obtained. Post treatment the membrane was gently removed from the agar using sterile tweezers and placed onto a 7 mm diameter circular glass slide and adhered using Carbon discs. The excess membrane was trimmed from the edges of the glass slide using sterile scissors. The biofilm was gently washed with deionized water to remove excess media and treatment. The biofilms were then fixed in a 1 wt% glutaraldehyde PBS buffer solution for 1 hr at room temperature. After fixation, the biofilms were washed 3 times gently with deionized water to remove excess glutaraldehyde. The samples were then dehydrated in 50%/75%/90% and absolute ethanol solutions for 30 min per each ethanol concentration. The absolute ethanol was dried using liquid CO2 at its critical point using an E3000 Critical Point Dryer (Quorum Technologies, UK) and then coated in 10 nm carbon layer. Samples were imaged with variable pressure 100-micron aperture at 40 Pa. EHT 20 kV, probe current 100 pA. Images were captured with Zeiss smartSEM software (Zeiss Evo-60 SEM, Germany). TEM images of bare Carbopol Aqua NPs, free Alcalase 2.4 L FG protease, and Alcalase 2.4 L FG coated Carbopol Aqua SF1 nanogels were obtained by placing a droplet of the suspended sample onto carbon-coated copper grids (EM Solutions, UK) and allowed to adhere for 2 min. The grid was quickly rinsed with deionized water and negatively stained with 1 wt% aqueous uranyl acetate. This was again quickly rinsed with deionized water, allowed to dry in air and then imaged with a Gatan Ultrascan 4000 digital camera attached to the Jeol 2010 TEM 2010 electron microscope (Jeol, Japan) running at 120kV. EDX data was collected via an Oxford Instruments Nanoanalysis X-Max 65-T detector and INCA software. Cytotoxicity and apoptotic stasis of HaCaT spontaneously transformed keratinocytes from histological normal skin after treatment with nanogel, Alcalase and Alcalase-nanogels. HaCaT cells were cultured in T75 flask using DMEM supplemented with 10% FBS and 1% L-glutamine under humidified conditions at 37 °C, 5% CO2. When a confluency of 80% was achieved, determined by visualization with an optical microscope, passaging was performed to ensure that the cells remained in the exponential phase for experimentation. Passaging was performed by removing spent media, washing in DPBS and incubating with 1 × trypsin EDTA at 37 °C 5% CO2 for 5 min until the cells were detached in suspension. The trypsin EDTA was then neutralized with a 1 : 1 volumetric ratio of fresh DMEM and gently centrifuged at 400 rpm for 4 min, the supernatant was aspirated, and the pellet was re-suspended in DMEM (supplemented as above) at a 1 : 6 ratio and transferred into a fresh T75 flask. Experimental cells were diluted in fresh DMEM, supplemented with 2% FBS and 1% L-glutamine and seeded at 5 × 104 in 100 μL media, were placed into 96-well plate wells and incubated for 24 hours at 37 °C 5% CO2. The medium was then removed and the cells and replaced with 100 μL of treatment infused DPBS. The treatments with only DPBS were used as a control, 0.6 wt% Carbopol, 0.6 wt% free Alcalase, and 0.6 wt% Alcalase -0.6 wt% Carbopol nanogel were used as treatments. DPBS was used so DMEM peptones did not interfere with the true activity of the Alcalase. Treatments were performed for 1, 6, and 24 hrs time points. A colorimetric (MTT) cell survival and proliferation assay kit was used to measure cell viability. Treatment culture was aspirated away and the cells washed for 2 min with fresh DPBS. 100 μL of fresh DMEM was added after which 100 μL of MTT reagent (50% 3-(4,5-dimethylthiazol-2yl)2,5-diphenyl tetrasodium bromide and 50% PBS) was added. This was incubated in the same conditions for 2 hrs until intracellular purple formazan crystals were visible under a light microscope. After 2 hrs, 100 μL of color development reagent in isopropanol with 0.04 M HCl was added for 1 hr, allowing the cells to lyse and the formazan crystal to be solubilized to a homogenous blue solution. The absorbance of the blue solution was read at 570 nm on a plate reader and subtracted from a blank of media only. These data were calculated into cell count data using the Beer-Lambert extinction coefficient law using absorbance values from a fixed number of cells in media. The HaCaT cell viability assay was repeated in three independent experiments. The induction apoptosis was investigated using a FITC Annexin V staining kit according to manufactures instructions. Apoptosis was measured after 24 hrs exposure to 0.6 wt% Alcalase-0.6 wt% Carbopol nanogel. Briefly, post treatment the cells were washed twice with DPBS and detached from the 96-well surface using a mini-cell scraper and re-suspended in 1X binding buffer at a concentration of 1×10 6 cells mL -1 . 5 μL of FITC Annexin V and 5 μL of Propidium Iodide (PI) solution to 100 μL of the cell suspension. The cells were then incubated at room temperature for 15 mins in darkness with gentle shaking. After incubating, 400 μL of 1X binding buffer was added to the cell samples which were then placed quantified by flow cytometry (BD FACSCalibur, USA). The Annexin V fluorescence was measured using a FITC-FL1 detector, and PI measured using a FL2 phycoerythrin signal detector. The experiment was ended after 10,000 cells were counted, the data was analyzed by a dot plot scatter and percentages of live, dead, and apoptotic cells determined by cell quadrating.

RESULTS AND DISCUSSION
Coating and immobilization of Alcalase 2.4 L FG on to Carbopol Aqua SF1 nanogel particles. A colloidal suspension of Carbopol Aqua SF1 was chosen as an appropriate nanocarrier for investigating the immobilization Alcalase 2.4 L FG protease on their surface. The lightly cross-linked carboxylic groups confer a net anionic surface charge corresponding to a zeta-potential of approximately -30 mV at a pH of 5.5. This allows it to be used as a substrate to immobilize the cationic protein on its surface. We chose to limit this suspension to 0.6 wt% to prevent the aggregation of the nanogel particles. Alcalase 2.4 L FG has a diameter of ~ 8 nm as shown from the dynamic light scattering size distribution and TEM micrographs (see Figure 2A and 3A-B, respectively). Data in Figure 2D shows that Alcalase 2.4 L FG has an isoelectric point of ~ 9 and a pH 5.5 has a zeta potential of ~ +18 mV. This allows a successful ionic interaction between the anionic Carbopol NPs and the cationic Alcalase 2.4 L FG. Figure 2C shows the bare 0.6 wt% Carbopol NPs have a hydrodynamic diameter of approximately 110 nm, however, when coated with 0.6 wt% Alcalase 2.4 L FG (Figure 2D) the particles mean diameter is increased to approximately 120 nm. Figure 2B confirms this through measurement of the hydrodynamic diameter of the Alcalase-coated collapsed nanogel particles. Figure S2 (ESI) shows a comparison between the bare Carbopol and Alcalase coated Carbopol nanogels. It can be seen that by increasing the concentration of the Alcalase, the coated Carbopol NPs average particle size also increases. This can be seen in the TEM images on Figure 3D, where the corona of the collapsed nanogel particles can be seen coated with a "halo" of the Alcalase protein molecules. The successful coating of the Alcalase on the Carbopol NPs was also confirmed using UV-Vis spectrophotometry. Figure 2F shows that Alcalase has a distinct absorption peak at ~ 245 nm, however, the bare Carbopol shows no such peak. When the Alcalase has coated the Carbopol particles, a small peak also at ~ 245 nm can be seen, indicating the Carbopol and Alcalase have ionically interacted and formed a composite Alcalase-coated Carbopol particles. Figure S3 (ESI) shows each of the formulations visually. The stability of the Alcalase-coated Carbopol nanogel particles was also investigated. Figure 2E shows that over 24 hrs, the zeta potential was stable in the range between 16-20 mV. This indicates the Alcalase remains electrostatically deposited in the nanogel particles over 24 hrs, indicating that it is a stable carrier for immobilization of the enzyme during treatment. Additionally, we investigated the EDX spectra of the bare and coated Alcalase-nanogels using samples prepared for TEM. (see Figure 3E-F). The EDX spectra are similar, however, a sulfur peak can be seen in the coated sample spectra, which is not present in the uncoated nanogel. The bare nanogel formulation contains only C, O, and N, so the presence of S, an element commonly found in protein indicates that the protein has electrostatically bound to the Carbopol nanogel particles. Fourier Transform Infra-Red spectroscopy (FTIR) was used to investigate the composition of the Carbopol/Alcalase/Ciprofloxacin NPs at a range of 600 cm -1 to 4000 cm -1 ( Figure S4, ESI). Carbopol Aqua SF1 shows a strong peak at 3350 cm -1 . This indicates stretching vibrations of O-H and intermolecular hydrogen bonding. Strong peaks are also seen at 3005 cm -1 and 1698 cm -1 correlating to strong stretching of aliphatic C-H and strong vibration of carbonyl (C=O) bonds. Peaks at 1000 cm -1 and 1195 cm -1 indicate coupling between in-plane O-H bending and C-O stretching of neighboring carboxyl groups. Alcalase 2.4 L FG shows weak peaks at 2950 cm -1 and 3001 cm -1 , which are characteristic of amino acid with aromatic side chains e.g. tyrosine, tryptophan and phenylalanine. The peak at 1753 cm -1 indicates strong C=O stretching, attributed to C=O bond adjacent to secondary amide (peptide) bond. The peak at 1156 cm -1 shows strong C-O-H group stretching, present in the amino acids serine and tyrosine. Ciprofloxacin hydrochloride shows a peak at 1451 cm -1 attributed to the strong vibration of C-H bonds. The peak 1255 cm -1 is associated with the aromatic C-F group present on ciprofloxacin. Peak 801 cm -1 is interpreted as tetra-substituted benzene ring derivatives, present on the ciprofloxacin chemical structure. The tightly packed range of peaks between 600 cm -1 to 1450 cm -1 indicates an array of substituted groups associated with the ciprofloxacin chemical structure. Ciprofloxacin encapsulated Carbopol, coated with Alcalase contains peaks associated with the individual components. The peak at 3358 cm -1 shows stretching vibrations of O-H and intermolecular hydrogen bonding, this is analogous to Carbopol alone. The peak at 1721 cm -1 provides evidence strong C=O stretching, attributed to C=O bond adjacent to secondary amide (peptide) bond, present in this formulation due to the coating effect on the Carbopol surface. High density peaks at 600 -1000 cm -1 again are similar to ciprofloxacin alone, which indicates that a variety of substituted groups associated with the ciprofloxacin being present within the Carbopol. The elemental analysis of Carbopol Aqua SF1, Alcalase 2.4 L FG and Ciprofloxacin alone, and Ciprofloxacin-loaded Carbopol nanogel coated with Alcalase 2.4 L FG was performed using a CHN Analyzer (Carlo Erba 1108) (Table S6, ESI). The formulated nanogel particles where centrifuged for 15 min at 4000 rpm. The supernatant was discarded and the pellet washed 3 times with deionized water. The precipitate was further dried overnight at room temperature under vacuum. The data shows the % value of C, H, N and S for the above materials. Carbopol alone contained only C and H, which is consistent with polyacrylic acid polymers. Oxygen is present in all samples but not analyzed. Alcalase contained C, H, and N, present in all amino acids, additionally it contained 0.48% S, which is present in the amino acids methionine and cysteine. This is confirmed by the EDX results in Figure 3F. Ciprofloxacin contains C, H and N.
The formulation of 0.6 wt% Carbopol Aqua SF-1-0.0032 wt% Ciprofloxacin-0.6 wt% Alcalase 2.4 L FG nanogel contained all elements analyzed, confirming that the Alcalase had been successful retained on the Carbopol during preparation. EDX spectra of the 0.6 wt% Carbopol-0.0032 wt% Ciprofloxacin-0.6 wt% Alcalase nanogel, and the individual components was performed on SEM prepared samples (Figure S5, ESI). It can be seen that the formulation contains both sulfur and fluorine peaks, characteristic of Alcalase and ciprofloxacin respectively. This provides evidence the formulated nanogel has successfully encapsulated ciprofloxacin and is coated with Alcalase. Alcalase 2.4 L FG was chosen as a potential protease to degrade biofilms due to its favorable activities (see Figure S6, ESI). It is highly active and stable between pH 6-9 in wide temperature range between 30-65 o C. Table S1 (ESI) also shows its optimal usage conditions are between 30-65°C and pH 7-9 (Novozymes data). These properties make it an ideal choice of bacterial biofilms which are grown at 37 o C and can grow at either acidic or basic conditions. 59

Protease activity of free Alcalase vs. Alcalase-nanogel.
Enzyme catalytic potential is affected by several factors. We investigated the effect of pH, temperature and time on the free enzyme, and the surface immobilized Alcalase on the Carbopol nanogel particles ( Figure S7, ESI). The effect of pH was investigated at 5.5, 7.5, and 8.5. As seen in Figure S7A, a higher pH increases the activity of the free Alcalase, and Alcalase-nanogel substantially. There is only a marginal difference between the protease performances of each sample, which is likely due to the gentle electrostatic attraction between the anionic Carbopol nanogel particles, and the cationic protease in this pH range, having little effect on the enzymes tertiary structure. Covalent enzyme immobilization methods have been shown to cause a decrease in protease activity for this reason. 60 A similar trend is observed when measuring the effect of temperature ( Figure S7B) on the activity of the free and immobilized Alcalase. Its activity broadly increased at physiological and higher temperatures, a common feature of biologically derived enzymes, but there is very little difference in performance between the samples. This data is in agreement with the enzyme activity data provided by Novozymes® (see Figure S6). This provides further evidence the Carbopol nanogel immobilization technique is exerting little influence on the enzyme activity. Figure S7C examines the effect time has the activity of the free and immobilized samples. As like pH and temperature there is only a small difference in activity, particularly up until 6 hr. At the 24 hrs time point, the immobilized Alcalase-Carbopol nanogel show and increased activity compared to the free counterpart.  Table S3 (ESI). P <0.05 is considered significant. *P < 0.05, **P < 0.01, ***P < 0.001.
This indicated that over time the Alcalase-nanogel can outperform the free Alcalase at equivalent concentration. However, it is concluded from this data that the immobilization progress is not allowing the enzyme functionalized nanogel to perform better due to any intrinsic property the Carbopol is inferring, but rather the nanogel allow the enzyme to concentrate and localize at key anionic structures of the biofilm, such as the cell wall, or biofilm surface adherence proteins. 61 Qualitative biofilm producing capability assessment of representative ATCC bacterial species. Before assessment of the Alcalase 2.4 L FG coated Carbopol NPs on biofilms, the ability of the six types of bacteria to produce biofilms was investigated. A simple glass tube Crystal Violet staining assay was used to assess their adherence to a smooth glass surface after 24 hrs of growth. Table S2 (ESI) shows that all six species were able to adhere to the glass tube. The adhered cells and biofilm was relatively scored based on the concentration of Crystal Violet dye retained after washing with deionized water. Figure S8 (ESI) shows the blue/purple coating of the glass over the area in which the bacterial cells were cultivated and adhered to the glass wall in a biofilm. S. aureus, P. aeruginosa, and S. epidermidis were shown to have strong adherence. K. pneumoniae and E. coli showed moderate adherence, and finally, E. faecalis was shown to have a weaker adherence. Staphylococcal, Pseudomonas, Escherichia, and Enterococcus species have all been previously shown to form biofilms. A Congo Red agar culture method, first demonstrated by Freeman et al., 62 was also performed (see Figure S9, ESI). This uses a highly nutritional agar supplemented with sucrose, encouraging the production of exopolysaccharides. Biofilm producing species produce black colonies, which can be seen on the S. aureus, P. aeruginosa, and S. epidermidis species. However, this was not expressed so well on the K. pneumoniae, E. coli, and E. faecalis species, although there was some darkening of the colonies. The sensitivity and the exact mechanism of the production of the black colonies is not well understood, and as it has been suggested by Freeman et al. 62 that it may be due to the presence of secondary metabolites.  Table S4 (ESI). P<0.05 is considered significant. *P < 0.05, **P < 0.01, ***P < 0.001.
We therefore, have referred to the Crystal Violet tube method which shows that all six species are biofilm producers. Biofilm disruption by Alcalase-coated nanogel particles. Proteolytic enzymes have previously been used, particularly in wound healing applications, as agents to remove necrotic tissue and cell debris. 33,63,65,66 We investigated if Alcalase 2.4 L FG coated Carbopol nanogels were able to disrupt the biofilms formed by the species described above. Biofilms were formed by culturing the bacterial species in TC-coated 48-well plates for 24 hr at 37 o C in static conditions. After 24 hrs, the media was removed and the biofilms adhered to the well plates were washed to remove any remaining planktonic cells. Fresh culture media was supplemented with 0.2 wt% Carbopol, 0.2 wt% Alcalase, and 0.2 wt% Alcalase coated Carbopol nanogel, un-supplemented media was used as a control. 0.1 wt% Crystal Violet was used to stain the adhered cells and biofilm matrix, before being solubilized in 30% acetic acid. The absorbance was taken to quantify the residual biofilm mass. Figure 4 shows that bare 0.2 wt% Carbopol nanogel showed very little biofilm disruption capability, approximately a 5% reduction in biofilm mass. 0.2 wt% free Alcalase showed a marked improvement in biofilm mass reduction. This ranged from around 80% to 40% residual biofilm left, with P. aeruginosa and K. pneumoniae showing the least and most reduction in biofilm mass respectively. Interestingly, across all six species the 0.2 wt% Alcalase coated 0.2 wt% Carbopol nanogel showed a much greater reduction in biofilm mass compared to the free Alcalase enzyme. This ranged from around 35% to 15% residual biofilm mass compared to the control. In S. aureus, P. aeruginosa, S. epidermidis, E. coli, and E. faecalis there was an over 50% decrease in the biofilm mass compared to the free Alcalase alone. K. pneumoniae showed a 20% decrease in the biofilm mass compared to the free Alcalase. Figure S10 (ESI) shows visually the reduction in Crystal Violet staining on the biofilms after treatment. To test the hydrolysis of the biofilm matrix, particularly the protein content, the same experiment was performed using 0.005 wt% Congo Red to stain the biofilm. This was solubilized in 30% ethanol and the absorbance taken to quantify the intensity of the staining of the biofilms after treatment. Congo Red has previously been used to stain amyloidosis, an accumulation of proteins, in histology sections. Increased concentrations of protein result in a more intense red staining. Figure 5 shows a similar correlation between the reduction of the biofilm mass observed by Crystal Violet staining. 0.2 wt% Carbopol nanogel showed no reduction to very little reduction of the protein content (see Figure S11 (ESI) for visual images of Congo Red biofilm stains). Free 0.2 wt% Alcalase showed between a 40% to 60% residual protein content within the biofilms. However, 0.2 wt% Alcalase coated 0.2 wt% Carbopol shows a much more impressive reduction in protein content within the biofilm, between 25% and 15 % residual protein compared to the control. This shows that the Alcalase coated nanogels are causing the degradation of the protein backbone of the biofilm. The immobilized Alcalase on Carbopol nanogel particles are likely resulting in an increase in enzyme stability and reduction in aggregation, allowing them to outperform their free enzyme counterparts. Resazurin sodium salt in situ bacterial biofilm metabolic activity assay. Resazurin Sodium Salt assay was used to measure the metabolic activity of the bacterial cells within the biofilm. Biofilms of the six species were growth for 24 hrs on Membrane Filter paper sections placed on to MHA. After 24 hrs of growth a biofilm was formed on the filter paper. The membrane biofilm was then transferred onto fresh MHA in a 6-well plate and submerged into treatment infused deionized water (Un-supplemented deionized water was used as the control). Treatments were increased to 0.4 wt% and 0.6 wt% Alcalase to investigate if higher concentrations of free Alcalase enzyme, and Alcalase-coated Carbopol nanogels would affect the viability of the cells within the biofilm after 24 hrs 0.015 wt% Resazurin sodium salt was used to measure the metabolic activity of the cells within the biofilm by applying evenly to the surface of the biofilm. Resazurin sodium salt can be used as a metabolic activity assay by measuring the production of resorufin (fluorescent pink) created in the presence of cellular NADH. 67 Only metabolically activity cells are able to reduce resazurin to resorufin. The development of resorufin on the biofilm was quantified to measure the metabolic activity of the biofilm after treatments. Figure 6 shows that 0.6 wt% Carbopol has no effect on the metabolic activity of the bacterial cells. 0.4 and 0.6 wt% free Alcalase was able to reduce the metabolic activity of cells within the biofilm by on average 60% to 40% across the six species. S. epidermidis showed by far the most decreased metabolic activity, 30% to 20 % viability with 0.4 and 0.6 wt% free Alcalase treatment. P. aeruginosa and E. faecalis were able to tolerate the free enzyme treatments much better, reducing to approximately 60 to 50% after treatment with 0.4 and 0.6 wt% free Alcalase, respectively. In agreement with the 48-well biofilms described above, the Alcalase-coated Carbopol nanogel performed much better than the equivalent concentration of the free Alcalase enzyme. In all six species, the 0.6 wt% Alcalase -0.6 wt% Carbopol nanogel were able to reduce the viability of the biofilms to approximately 4%. Figure S13 (ESI) shows the intensity of the purple color of biofilm after treatment with 0.6 wt% Alcalase -0.6 wt% Carbopol. This contrasts the control image which shows a pink color indicating the biofilm contains metabolically active cells.

S. aureus biofilm cellular viability after treatment with
Alcalase-nanogel particles and co-treatment with free ciprofloxacin hydrochloride. Alcalase-coated Carbopol nanogel has been shown to reduce the biofilm mass, the protein concentration of biofilm matrix, and biofilm metabolic activity. We decided to investigate the viability of the cells with in the biofilms after treatment by CFU enumeration. S. aureus was chosen as the proxy biofilm producing bacterium for these experiments. S. aureus is considered one of the most prevalent pathogens in wound infection, 70 and has been widely studied making it a good choice for further investigation. Biofilms were growth on filter membrane atop MHA and treated with 0.6 wt% bare Carbopol, 0.6 wt% free Alcalase, and 0.6 wt% Alcalase coated 0.6 wt% Carbopol in the same fashion as above. Additionally, we investigated the effect of co-treatment with the antibiotic ciprofloxacin as a free agent and equivalent concentrations of ciprofloxacin encapsulated into Carbopol.  Table S8 (ESI). P<0.05 is considered significant. *P < 0.05, **P < 0.01, ***P < 0.001.
We have previously demonstrated that antibiotics can be encapsulated into Carbopol and this confers an increased efficacy of the antimicrobials against target cells. 55 Table S5 (ESI) shows that 75% of the ciprofloxacin encapsulated with the Carbopol was retained. The encapsulation efficiency was measured by analyzing the supernatant of the formulation immediately after centrifugation, the encapsulation efficiency was calculated using a standard calibration curve of absorption vs. known concentrations of ciprofloxacin ( Figure S14, ESI). Post-encapsulation, the mean particle diameter was marginally increased by approximately 1 nm post encapsulation and there was no detectable change in zeta potential of the nanogel particles ( Figure S15, ESI).
Previously, polyelectrolytes have been used to reverse the charge of nanogel and allowed an electrostatic adhesion to anionic surface of the bacteria. 14,15,20 In this study, we used Alcalase to accomplish this goal to create a dual purpose nanocarrier able to effectively degrade the biofilm and simultaneously deliver concentrated doses of antibiotic localized directly on the cell wall. This strategy is known to work extremely well for planktonic bacteria, but so far this has never been tried for bacteria in a biofilm. The release kinetics of the nanogel-loaded ciprofloxacin hydrochloride was measured over 24 hr (Figure S16, ESI) and as in previous studies the antibiotic was released slowly over a period of 24 hrs after dilution of the stock suspension. This facilitated the release of ciprofloxacin in close proximity to the S. aureus. Cell enumeration was performed after 1, 6, and 24 hr of treatment. The treated biofilms were glass bead beaten and enumerated using the drop plate method. Ciprofloxacin was chosen as an appropriate co-treatment as it can be administered topically, orally, and intravenously. Figure S17 (ESI) shows that the S. aureus strain used in this study was susceptible to ciprofloxacin and had a MIC of 0.08 g/mL (0.000008 wt%). We chose 0.0032 wt% as this is the maximum concentration measured to be effective against S. aureus using the ETEST® strips. This allowed a fair comparison of formulated vs. free ciprofloxacin at concentrations shown to be effective against S. aureus. Additionally, this concentration allows the nanocarrier colloidal stability to be maintained due to ciprofloxacin's low water solubility. Figure 7A shows that after seeding a membrane with 1×10 5 CFU/mL for 24 hr at 37 o C, the growth control continued to increase in cell density after a further 1, 6, and 24 hr of growth. Treatment with 0.6 wt% bare Carbopol showed little change in the CFU/mL of biofilm. After treatment with 0.6 wt% free Alcalase, there was a reduction in viable cells within the S. aureus biofilm. There was little difference after 1 hr, but after 24 hr the viable cell count was reduced to just under 1×10 5 CFU/mL, very close to the concentration of cells the original membrane was seeded with.Surprisingly, after treatment with 0.6 wt% Alcalase -0.6 wt% Carbopol NPs there was very little difference in the number of viable cells after each treatment time point. This is likely due to the Alcalase-Carbopol nanogel degrading the biofilm, but having very little effect on the bacterial cell viability, as there is no antibiotic in this treatment to act on the released bacteria. Treatment with 0.0032 wt% ciprofloxacin (higher than the MIC of 0.00008 wt%) was used to measure the number of viable cells, after 24 hrs of treatment the viable cells in the S. aureus biofilm had reduced from approximately 1×10 6 to 1×10 3 . The cells within the biofilm are much less susceptible to antibiotics due to their poor penetration through the biofilm matrix. There was no real improvement when 0.0032 wt% ciprofloxacin is treated with 0.6 wt% Carbopol. This result is unsurprising due to the Carbopol nanogel alone demonstrating no ability to degrade the biofilm matrix. Again with 0.00032 wt% ciprofloxacin and 0.6 wt% free Alcalase there was no difference between the CFU/mL compared to the antibiotic alone. However, when the antibiotic is administered as a co-treatment with 0.6 wt% Alcalase coated 0.6 wt% Carbopol NPs there was a reduction to 1×10 1 . This indicates the cells in the biofilm have been destroyed due to the degradation of the biofilm matrix by the Alcalase-Carbopol nanogel allowing the ciprofloxacin to reach and penetrate the bacterial cells. This synergistic effect was enhanced further when the ciprofloxacin was delivered to the S. aureus in an encapsulated method, rather than as a separate co-treatment. After 6 hr and 24 hr of treatment with 0.0032 wt% ciprofloxacin encapsulated in 0.6 wt% Carbopol with 0.6 wt% Alcalase coating there were no viable bacteria found in the samples. This provides evidence that the dual functionality of the Alcalase-nanogel mediate biofilm degradation and the localized delivery of ciprofloxacin of this formulation is able to effectively clear the biofilms and kill all released bacterial cells.  Figure 7B shows that there is a correlation between the increasing of the concentration of free ciprofloxacin in the presence of equivalent amount of Alcalase-Carbopol NPs and the residual bacterial cell viability after treatment. Different dilutions of the treatments were prepared to measure this effect. A very low treatment concentration, 0.075 wt% Alcalase -0.075 wt% Carbopol + 0.0004 wt% ciprofloxacin, showed very little effect after 1, 6, or 24 hrs. This is due to the likely ineffective degradation of the biofilm. However, as the concentration of the Alcalase-Carbopol NPs increased alongside the increased concentration of ciprofloxacin, CFU/mL was reduced to almost no viable cells. After 24 hrs at the higher concentration of the formulation, there was very little difference in the CFU at the 1, 6, and 24 hr time points, this indicates the Alcalase-coated nanogel particles are very fast acting, enabling the ciprofloxacin access to the cells quickly. Interestingly, at the mid-treatment concentrations CFU/mL increased between 1 hr and 6 hr, however a 24 hr it had fallen below the first time point. This indicates the lower concentrations of Alcalase-Carbopol NPs are slower to effect the biofilms. 0.6 wt% Alcalase -0.6 wt% Carbopol nanogel was found to be the most effective concentration to kill cells with in a biofilm when combined with the synergistic effect of ciprofloxacin. Figure 7C shows the results on the concentration dependence of the cell viability on the antibiotic treatment in experiments where the ciprofloxacin was encapsulated within the Carbopol nanogel, rather than being delivered separately as a co-treatment. We were able to show that there was a significant decrease in cell viability after 1, 6 and 24 hr compared to when ciprofloxacin was administered separately at the same concentration. We attribute this to the highly localized release of ciprofloxacin from the adhered nanogel enhancing the effect of the antibiotic. Alcalase-functionalized Carbopol nanogel particles can be seen attached to the bacterial cell after the biofilm has been cleared away ( Figure 9D) providing evidence of their ability of local delivery antibiotic directly to the bacterial cell wall.
Anti-biofilm efficacy of Alcalase-nanogel particles on S. aureus biofilm structure. We also investigated how 0.6 wt% Alcalase -coated 0.6 wt% Carbopol nanogel affects the biofilm structure of S. aureus using fluorescence microscopy. S. aureus was allowed to adhere to a glass slide which has been submerged in this culture for 24 hr at 37 o C. After 24 hr the media was removed and the glass slides washed to remove non-sessile cells. Supplemented fresh media containing 0.6 wt% Carbopol, 0.6 wt% free Alcalase, and 0.6 wt% Alcalase -0.6 wt% Carbopol NPs was incubated for a further 24 hrs covering the slides. Acridine Orange (AO) was used to visualize the remaining cells after treatment. AO is a cell-permeable nucleic acid selective fluorochrome dye. It can emit green light at 520 nm when intercalated with dsDNA. Figure 8A shows the growth control has a highly congested biofilm structure, the cells are highly packed which is consistent with biofilm formation. There is very little difference to the biofilm structure after treatment with 0.6 wt% bare Carbopol and 0.6 wt% free Alcalase (Figure 8B, 8C). The cells remain packed densely together, although there is a moderate reduction in the cell density when treated with free Alcalase. However, in Figure 8D, there is a marked reduction in the intensity of the AO fluorescence and a reduction in the cell density. The cells have much lower surface density. This suggests the disruption of the biofilm matrix by the Alcalase-Carbopol nanogel particles has allowed many of the cells to detach from the biofilm during treatment, or during the followed washing. For better understanding of the S. aureus biofilm structure, the same samples used for fluorescent analysis were imaged using AFM (Figure 8E-H). The AFM images again show how the 0.6 wt% Alcalase -0.6 wt% Carbopol NPs effectively remove the biofilm from the glass slide. Figure 8E shows the tightly packed biofilm with staphylococcal cells of thickness approximately 1 μm; the cells appear to be in a multilayer due to the height sensor recording a maximum height of 2.2 μm. There was no clear differences in structure between the treatments of 0.6 wt% bare Carbopol nanogel and 0.6 wt% free Alcalase treatments ( Figures  8F and 8G, respectively). There is only a very small reduction in the average thickness of the biofilm of 0.1 μm which is insignificant. The AFM scans again show staphylococcal layer of approximately 1 μm in a tightly packed fashion. In the control, 0.6 wt% bare Carbopol nanogel and 0.6 wt% free Alcalase there was no discernible difference in cellular morphology. The image in Figure 8H, however, shows the biofilm structure has largely been removed, in the scanned region there were no obvious signs of cell adherence and the peak height was 0.5 μm which is half the average diameter of an S. aureus cell. This results indicates that the cells have been removed during the Alcalase-Carbopol nanogel treatment, and washed away after treatment during the preparation of the glass slides for AFM microscopy. This correlates with the data obtained in the AO fluorescence microscopy images. Finally, we investigated the structure of the S. aureus biofilm using scanning electron microscopy (SEM). In this case, we used biofilms grown of filter membranes as this allowed feasible samples to be obtained for SEM analysis. The samples were prepared in the same manner as described in the cellular viability assays of biofilm grown S. aureus reported above. The biofilms were treated with 0.6 wt% bare Carbopol nanogel, 0.6 wt% free Alcalase, and 0.6 wt% Alcalase-coated 0.6 wt% Carbopol nanogel particles. Un-supplemented media was considered the growth control. After the treatment, the biofilm membranes were gently removed from the agar using serial tweezers, however, instead of enumeration the membrane was washed in deionized water, fixed for 1 hr in 1 wt% glutaraldehyde, dehydrated in ethanol and dried in liquid CO2 at critical point as described in the methods section. 10 nm of carbon coating was used to create a conductive layer over the biofilm, and reduce thermal damage during SEM imaging. Figure 9 shows the SEM images after the treatments. The control sample shows the highly congested S. aureus biofilm. At higher magnification, the EPS within the matrix can also be seen adhered to the surface of the coccus cells and linked to other cells around, consistent with biofilm formation. This morphology is similar in the samples with 0.6 wt% free Carbopol nanogel treatments. The cell density remains highly analogous to the control sample and EPS matrix can be observed around the cells. The cells appear to also be in multilayer with cells atop one and other. Interestingly, in the sample treated with free 0.6 wt% Alcalase, the cellular morphology and density remain largely unchanged, however, there appears to be a distinct lack of EPS matrix coating and surrounding the cells. This indicates the Alcalase may have digested the biofilm matrix but left the cells largely untouched. Finally, the sample treated with the Alcalase-Carbopol NPs there is a clear reduction the cell density and additionally the biofilm EPS matrix has been degraded. The Alcalase-Carbopol nanogel particles can also be seen adhered to the surface of the S. aureus cells, due to the cationic protease molecules on the nanogel surface having an electrostatic attraction to the anionic surface of the bacterial cell wall. This data correlates to the images seen using fluorescence and atomic force microscopy, and confirms that the Alcalase-Carbopol nanogel particles are degrading the biofilm glycoprotein network. Zeta potential of bacteria treated with free Alcalase and Alcalase-nanogel. We also investigated the effect the free Alcalase and the immobilized Alcalase on-Carbopol nanogel on the zeta potential of the bacterial species. The experiments on the cells were performed in suspension in allow zeta potential measurements by DLS. Figure S12 (ESI) indicates that all six species conferred a negative charge on their cells walls, with the Gram-negative generally having a stronger charge than the Gram-positive. Across all six species, the free Alcalase showed a marginal reduction in the negative charge after 10 mins and 1 hr. This was likely due to the accumulation of protease on the cell wall. The charge difference was only slight between 10 mins and 1 hr, indicating the Alcalase protease had not built up and saturated the surface of the cells. In contrast, the Alcalase-Carbopol nanogel particles showed an increased reduction of the negative charge of the bacterial cells. On average, this was reduced by approximately 7-10 mV across the six species. This indicated a buildup of the cationic protease-coated nanogel carriers on the surface on the bacterial cells. The SEM images of S. aureus in Figure 9D also support this data, showing the Alcalase-Carbopol particles adhered to the bacterial cell wall. The experiments where stopped after 1 hr due to the aggregation of the cells causing unreliable zeta potential measurements. Whilst these experiments are not indicative of the behavior of the cells in a biofilm, which will have a much more complex EPS matrix surrounding them, it does provide a valuable insight into the increased attraction of the Alcalase-Carbopol nanogel to the exposed bacterial cells, and therefore the likely increased affinity of these particles to protein structures within the biofilm matrix. Comparison with standard treatments for biofilm forming bacteria. Currently, there is no Gold Standard when it comes to topically treating wound biofilms. There is universal agreement that mechanical debridement is the most effective way of remove biofilm infection, but this is not always possible and often results in regrowth of the biofilm creating a chronic infection. 68,69,70  Long term antibiotic treatment and topical antimicrobial treatments are other common options in treating biofilms. After decades of use, iodophore-based formulations such as Poly(vinylpyrrolidone)-Iodine (PVP-I) remain prevalent in wound biofilm treatments. 71 Iodine based agents are effective antimicrobial agents, which are tolerated well by the body and offer a broad spectrum of activity. 72 Additionally, they are able to penetrate biofilms, and no clinical evidence presently shows they have a negative effect on wound healing. 73 Chlorohexidine is also often used to as a disinfectant on patients and is commonly found in formulations designed to reduce infection. However, it is not known to be effective against cells within biofilms. 74 Figure 10 shows a comparison between 0.6 wt% Carbopol-0.0032 wt% Ciprofloxacin-0.6 wt% Alcalase nanogels against Iodine, PVP-I and chlorhexidine using the membrane biofilm model. The results show that at the concentrations similar to active agent in our formulation, ciprofloxacin, the alternative treatments are ineffective. This is likely due requirement of higher concentrations needed to be effective and the lack of penetration into the biofilm. The quenching effective of the EPS is paramount in preserving cell viability within a biofilm. Additionally, we tested the effect of 0.2 wt% chlorhexidine, and 10 wt% PVP-I, concentrations which are commonly used clinically in wound cleaning and sterilization. 73,75 Again, the results follow the same trend as the low concentrations, with neither agent able to fully eliminate the biofilm bacteria viability. Cytotoxicity and apoptosis inducing ability of bare nanogel, free Alcalase, and Alcalase-nanogel on human cells. A preliminary human cell cytotoxicity experiment was performed on HaCaT cells to investigate if the bare Carbopol Aqua SF1, the free Alcalase 2.4 L FG, and the Alcalase 2.4 L FG-coated Carbopol Aqua SF1 nanogel particles were toxic to human cells. HaCaT cells are very good proxy for studying cytotoxicity of the treatment as they are human keratinocytes which are the same type of cells likely to be exposed to the formulation in chronic wound cells. Figure   11A shows that 0.6 wt% Carbopol showed only a marginal reduction in viable cells over 1, 6, and 24 hrs. This was also the case with 0.6 wt% free Alcalase 2.4 L FG, and Alcalase 2.4 L FG-coated 0.6 wt% Carbopol Aqua SF1 nanogels, the same formulations used in the anti-biofilm experiments. This indicates that the Alcalase-coated Carbopol nanogels confer only a slight increase in cytotoxicity towards the Ha-CaT cells, compared with the free enzyme and the bare nanogel. The data indicates that the enzyme concentration which provided effect anti-biofilm effect had very little impact on cell viability, providing evidence they may be appropriate for in vivo use in surface chronic wounds. Additionally, we investigated the effect the 0.6 wt% Alcalase-0.6 wt% Carbopol nanogel particles on inducing apoptosis after 24 hrs treatment. An Annexin V-FITC staining kit was used to measure the induction of apoptosis using FACS (Fluorescence-activated cell sorting). Figure 11B shows that after 24 hrs exposure to the Alcalase-coated Carbopol nanogel there was approximately a 10% residual viable cells compared to untreated cells, which correlates well with the MTT viability data shown in Figure 11A. There is only a marginal difference in induced apoptosis (cells staining positive for Annexin and negative for PI). This indicates that the Alcalase 2.4 FG protease is not triggering apoptosis by way of the caspases cascade apoptosis pathway.

CONCLUSIONS
Our results confirm that protease Alcalase 2.4 L FG can disrupt the bacterial biofilm EPS matrix and reduce its biomass. The surface functionalization of Carbopol Aqua SF1 nanogels with this enzyme further boosts this effect on six wound biofilm associated pathogens reducing the biomass 6-fold compared to untreated biofilms. The efficacy of the Alcalase-coated Carbopol nanogel particles was confirmed using fluorescence, atomic force and scanning electron microscopy. Additionally, the application of Alcalase-Carbopol nanogels in a co-treatment with ciprofloxacin resulted in a 3-log reduction in viable cells counts compared to using of equivalent concentration of ciprofloxacin alone. When ciprofloxacin was encapsulated within Carbopol at equivalent concentrations to the free ciprofloxacin, there was a very significant further boost to the effectiveness of the antibiotic in this dual functionalized nanocarrier. These dual purpose Alcalase-coated antibiotic loaded nanogel particles provided effective treatment against S. aureus biofilms, with no detectable CFU after 6 hr and 24 hr treatment times. The Alcalase-Carbopol nanogels also showed negligible cytotoxicity and apoptosis inducing influence towards adult human keratinocytes. We conclude that the Alcalase-coated and ciprofloxacin loaded Carbopol nanogel carrier are an effective and apparently safe anti-biofilm formulation which could provide an alternative approach to treatment of external wound biofilms. We believe further investigation of the practical administration of Acalase-coated Carbopol nanogels loaded with antibiotics on biofilm infected wounds is required, where these nanogels could provide an advantageous tool for therapeutic anti-biofilm treatments and fight against antimicrobial resistance.

ASSOCIATED CONTENT
In the enclosed electronic supplementary information (ESI) we present the following additional data: (i) Schematic of the enzymatic degradation of peptide bonds in proteins by    Figure S4. Fourier transform infra-red (FTIR) spectra of Carbopol Aqua SF1, Alcalase 2.4 L FG, Ciprofloxacin and Carbopol encapsulated Ciprofloxacin coated with Alcalase NPs. Samples were prepared by removing water using a Silica gel desiccant by in vacuum chamber for 1 day, then dried at 60 o C for 2 days. The spectra was measured between 4000-600 cm -1 using PIKE ATR diamond settings. A spectrum was obtained with a blank of just air to reduce transmittance interference with the samples.        Figure S12. The zeta potential of bacterial cells treated with either 0.2 wt% free Alcalase or 0.2 wt% Alcalase-0.2 wt% Carbopol NPs. The bacteria / treatment suspension was placed in a quartz cuvette and zeta potential was measured using the Dipstick probe and Malvern Zetasizer nano ZS. The refractive index was 1.384 and the absorption was 1.000. N = 3 with ±S.D.   Figure S14. Standard calibration graphs of the absorption vs concentrations of ciprofloxacin hydrochloride. Absorbance was measured at 280 nm. Aliquots were prepared by adding ciprofloxacin hydrochloride to 250 mL Milli-Q water (adjusted to pH 5.5 using acetate buffer) at specific concentrations.      Table S8. Statistical analysis of free ciprofloxacin vs encapsulated ciprofloxacin (0.6 wt% Alcalase-0.6 wt% Carbopol formulation) at 1, 6 and 24-hour time points. Data were expressed as average values ± standard deviations of the mean. P-values of less than 0.05 were considered significant. All Student's T-tests were performed in GraphPad v7.0.4.