New Hydrogels Nanocomposites Based on Chitosan, 2-Formylphenylboronic Acid, and ZnO Nanoparticles as Promising Disinfectants for Duodenoscopes Reprocessing

Abstract

New hydrogels nanocomposites, based on iminoboronate hydrogels and ZnO nanoparticles (ZnO–NPs), were obtained and characterised in order to develop a new class of disinfectants able to fight the nosocomial infections produced by duodenoscopes investigation procedures. The formation of the imine linkages between chitosan and the aldehyde was demonstrated using NMR and FTIR spectroscopy, while the supramolecular architecture of the developed systems was evaluated via wide-angle X-ray diffraction and polarised optical microscopy. The morphological characterisation of the systems via scanning electron microscopy revealed the highly porous structure of the materials, in which no ZnO agglomeration could be observed, indicating the very fine and homogenous encapsulation of the nanoparticles into the hydrogels. The newly synthetised hydrogels nanocomposites was proven to have synergistic antimicrobial properties, being very efficient as disinfectants against reference strains as: Enterococcus faecalis, Klebsiella pneumoniae, and Candida albicans.

1. Introduction

One of the most concerning outcomes of recent years has been the nosocomial infection outbreaks brought on by multidrug-resistant organisms (MDRO) after endoscopic retrograde cholangiopancreatography (ERCP) procedures [1]. Endoscopic retrograde cholangiopancreatography (ERCP), first performed in 1968, is an advanced medical treatment designed to perform a minimally invasive diagnosis and treat pancreaticobiliary disorders [2,3]. Due to their intricate construction, duodenoscopes are challenging for proper reprocessing [4,5,6].

After reprocessing, duodenoscope contamination rates range from 0.4% to 35.8% [1,7,8,9,10,11,12,13]. The majority of nosocomial infections display adhesion problems at duodenoscopes, making it challenging to access these surfaces for reprocessing due to their persistence. The most frequently encountered pathogens, such as Klebsiella pneumoniae, Pseudomonas aeruginosa, Escherichia coli, and Salmonella faecalis, are known for their ability to form an MDRO and adhere to surfaces and spaces that are difficult to access for reprocessing [14,15,16,17,18]. Our team has prior research experience in this area, with studies describing the effects of routine procedure use and reprocessing cycles on the duodenoscope, demonstrating usage-related changes to both coating and working channel polymers, and demonstrating that the external changes progress progressively from the distal to the proximal to the elevator sample [19,20]. According to reported duodenoscope contamination rates, present reprocessing techniques are insufficient. This highlights the need for additional research into novel classes of disinfectants to enhance reprocessing outcomes and prevent future outbreaks and infections linked to duodenoscopy.

Within this framework, the literature abounds in information related to the antimicrobial activity of ZnO–NPs against Escherichia coli [21,22,23], Pseudomonas aeruginosa [24], Staphylococcus aureus [25,26], Klebsiella pneumonia [22,27], Enterococcus faecalis [28,29,30,31], and Candida albicans [29,32]. ZnO–NPs activity against drug-resistant pathogens is considered to depend on size and doping amount; they can be cost-effective and are quite stable for long-term storage with a prolonged shelf-life [33]. Moreover, MeO–NPs can be subjected to sterilisation via methods of high temperature, gamma irradiation, or plasma treatment without losing their properties or inactivation [34]. In the future, they could be combined with various classes of substances for optimal antimicrobial activity due to their additive nature [35].

Within this context, this study aims to obtain a new class of disinfectants based on the synergic activity of chitosan, boronic aldehyde, and ZnO and able to surpass the difficulties of duodenoscope reprocessing and to decrease the incidence of nosocomial infections in this procedure. Previous studies on iminoboronate chitosan hydrogels demonstrated the very high antifungal activity of these systems, on both planktonic yeast and biofilm [36] on two Candida strains—C. glabrata and C. albicans. Moreover, hydrogels based on chitooligosaccharides and boronic aldehyde presented high activity against nine reference strains: Staphylococcus aureus, Escherichia coli, Candida albicans, Candida parapsilosis, Candida glabrata, Saccharomyces cervisiae, Penicillium crysogenum, Cladosporium cladosporioides, and Aspergillus brasiliensis [37]. ZnO–NPs, on the other hand, are known in the literature for their antimicrobial properties against various microbial strains [38].

The aim of the present study is to obtain new disinfectants in the form of hydrogels based on iminoboronate chitosan derivatives, in which the ZnO–NPs should be homogenously dispersed, with the goal of both reaching synergism and broadening the antimicrobial activity of the hydrogels to a large number of microorganisms through the presence of ZnO–NPs. This combination has not been reported elsewhere, as far as we know.

2. Materials and Methods

2.1. Materials

Chitosan (147 kDa, DD = 88%), 2-formylphenylboronic acid (2-FPBA), 95% zinc acetate dihydrate, sodium hydroxide, acetic acid, and ethanol were purchased from Sigma (St. Louis, MO, USA) and used as received.

2.2. The Synthesis of ZnO Nanoparticles (ZnO–NPs)

The synthesis of ZnO–NPs was performed following a previously reported study, using the co-precipitation method [39]. Briefly, 30 mL Zn(Ac)₂·2H₂O (0.1 M) solution and 25 mL NaOH (0.2 M) solution in H₂O were subjected to magnetic stirring for 2 h at a temperature of 60 °C. After 2 h, the initially clear solution turned into a milky white dispersion. The resulting white product was separated via centrifugation at 5000 rpm for 10 min and washed 3 times with ultrapure water. ZnO–NPs, in powdered form, were obtained by drying the product in a conventional laboratory oven at 35 °C for 48 h.

2.3. The Synthesis of the Hydrogels Containing ZnO–NPs

Three hydrogels containing ZnO–NPs were synthesised using an in situ hydrogelation method of chitosan with 2-formylphenylboronic acid in the presence of ZnO–NPs. The hydrogels contained the same amount of ZnO but had different crosslinking degrees, depending on the molar ratio used between chitosan and 2-FPBA, ranging from 1/1 to 2/1. At lower molar ratios between the reagents’ functionalities, the hydrogelation was not reached. For example, the experimental protocol for the obtaining of the S2 sample was as follows: to a solution of chitosan in acetic acid aqueous solution (see

Table 1. The composition of the obtained systems.

Sample Code	mchitosan (mg)	Vwater (mL)	Vac.acid (μL)	maldehyde (mg)	Vethanol (mL)	NH2/CHO Molar Ratio	mZnO (mg)
S1	60	6	42	44.8	4.48	1	18
S1.5	60	6	42	33.3	3.33	1.5	18
S2	60	6	42	22.4	2.24	2	18
R	60	6	42	-	2.24	-	18

), a mixture of ZnO and 2-FPBA in ethanol was added at 55 °C under continuous vigorous magnetic stirring (1500 rot/min). The hydrogelation was reached after different time intervals, depending on the NH₂/CHO molar ratio, from 5 min for sample S1 to 3 h for samples S2. A sample containing only ZnO dispersed into chitosan was also obtained and used as reference (R).

2.4. Characterisation

2.4.1. Hydrogels’ Lyophilisation

The hydrogels containing ZnO–NPs (S1–S2) and the reference sample (R) were freeze-dried using a LABCONCO FreeZone FreezeDry System (Fort Scott, Kansas City, MO, USA), at −54 °C and 1.510 mbar, for 24 h.

2.4.2. FTIR Spectroscopy

The ATR–FTIR spectra of the hydrogels after lyophilisation (xerogels) were recorded on a FTIR Bruker Vertex 70 Spectrophotometer (Ettligen, Germany), with a ZnSe single reflection ATR accessory.

2.4.3. NMR Spectroscopy

The ¹H NMR spectra were recorded on a Bruker Avance NEO 400 MHz spectrometer (Ettlingen, Germany, Bruker Biospin) equipped with a 5 mm broadband inverse detection z-gradient probe, using NOESY water presaturation–pulse sequence. All the spectra were recorded through 64 scans and were manually processed with Bruker TopSpin 4.2.0 spectrometer control and processing software.

2.4.4. Scanning Electron Microscopy

The morphology of the xerogels was investigated with a Scanning Electron Microscope SEM EDAX—Quanta 200 (Eindhoven, Germany) and the obtained images were processed with Image J and Origin Pro 8 Software.

2.4.5. Wide-Angle X-ray Diffraction

Wide-angle X-ray diffraction analysis was performed on a Rigaku Miniflex 600 diffractometer using CuKα emissions in the angular range of 2–80° (2θ) with a scanning step of 0.0025° and a recording rate of 1°/min.

2.4.6. Polarised Optical Microscopy

The obtained systems were evaluated through polarised optical microscopy (POM) on a Leica DM2500 microscope (Hamburg, Germany).

2.4.7. Atomic Force Spectroscopy (AFM)

The samples collected from surface of the duodenoscope and the samples exposed to the hydrogel and microbial strains were analysed using an NTEGRA Spectra (NT-MDT, Zelenograd, Russia) instrument with the 3.1–37.6 N/m force constant cantilever of a silicon nitride cantilever (NSC10; NT-MDT, Russia) in tapping mode.

2.5. Antioxidant Activity

The antioxidant activity of the samples was evaluated by analysing their ability to capture the 2,2-diphenyl-1-picrylhydrazyl (DPPH) radicals from a stock solution 0.025 mg DPPH/ mL in ethanol. The spectra of all samples, DPPH control solution, and the solutions of the samples, incubated with the DPPH solution, were recorded using an UV–Vis Spectrophotometer Cary 60. The DPPH maximum from 517 nm was used for the calculation of the samples’ radical scavenging ability. Known amounts of samples, all containing 10 mg of chitosan, were dissolved in 1.5 mL water and 10.5 μL acetic acid. To these solutions, 1.5 mL DPPH solution in ethanol was added and incubated for 1 h at room temperature and in the dark. After 1 h, the UV–VIS spectra were recorded and the absorbance at 517 nm was read.

The antioxidant activity of the samples (RSA%) was calculated using the following equation:

$$\mathrm{RSA}= \frac{{\mathrm{A}}_{\mathrm{ref}}-{\mathrm{A}}_{\mathrm{sample}}}{{\mathrm{A}}_{\mathrm{ref}}}\times 100,$$

where RSA is the radical scavenging ability, A_ref is the absorbance of the DPPH solution without sample, and A_sample is the absorbance of the DPPH solution, which was incubated with the samples for 1 h.

2.6. Antimicrobial Activity

External samples of 1 cm² each were taken from an international market reference duodenoscope (TJF-160F, Olympus Corporation, Tokyo, Japan), selected from a high-volume tertiary gastroenterology centre, previously used in approximately 500 ERCPs. This model, belonging to the penultimate generation of duodenoscopes, has established itself as one of the most used instruments globally, characterised by reliability over time and high-level technological quality.

The antimicrobial activity screening of the hydrogels samples (S1, S1.5, and S2) and controls (chitosan as Ch; ZnO,2-formylphenylboronic acid as BA; and chitosan + ZnO as sample R) was performed by using viable cell-counting method [40]. Three reference strains with high incidence in the nosocomial infections produced by ERCPs were used, i.e., Staphylococcus aureus ATCC25923 (S. aureus), Klebsiella pneumoniae ATCC10031 (K. pneumoniae), and Candida albicans ATCC90028 (C. albicans). The bacterial strains were refreshed on nutrient agar (NA) and the yeast strain was refreshed on Sabouraud dextrose agar (SDA) at 37 °C. Microbial suspensions of these cultures were prepared in sterile nutrient broth medium in order to obtain turbidity optically comparable to a 0.5 of the McFarland standards. A total of 500 μL of the hydrogels and controls was placed into the solution, which contained 0.5 mL bacterial suspension and 4.5 mL 1X PBS solution as well as the coating samples for the duodenoscopes, followed by incubation in a shaker at 37 °C up to 24 h. A control experiment was conducted. A total of 5 μL of the control samples and of the treated samples was removed at determined periods of incubation time (10 min, 20 min, 40 min, 1 h, 2 h, 3 h, 6 h, and 24 h) and spread on plate count agar (PCA) plates. The number of colonies was counted after 24 h of incubation at 37 °C.

All tests were carried out in triplicate to verify the results. After incubation, the plates were analysed using SCAN1200^®, version 8.6.10.0 (Interscience, Saint-Nom-la-Bretèche, France). The ratio of the inhibited the growth of bacteria (IRG) was calculated and is defined as:

IRG (%) = (Na − Nb)/Na × 100%

where Na and Nb are the average values of colonies of the control group and the experimental groups, respectively.

The duodenoscope samples were removed, washed with sterile water, and the surface morphology was examined by using a scanning electron microscope SEM EDAX—Quanta 200 (Eindhoven, Germany) and atomic force spectroscopy (AFM) NTEGRA Spectra (NT-MDT, Zelenograd, Russia).

2.7. Statistical Analysis

Data analysis was performed using GraphPad Prism software version 7.00 for Windows (GraphPad Software, La Jolla, San Diego, CA, USA, https://www.graphpad.com/, accessed on 1 May 2023). The obtained results represent the mean ± standard deviation (SD) of three different experiments. Two-way ANOVA was applied to determine the statistical significance between the mean of cells viability. A value of p < 0.05 was considered statistically significant, and the significance is noted as * p < 0.05; ** p < 0.01; and *** p < 0.001.

3. Results and Discussion

New hydrogels composites containing ZnO–NPs were obtained through the combination of chitosan with 2-formylphenylboronic acid in three molar ratios of their functionalities (Table 1, Scheme 1). The design of the systems took into consideration the antimicrobial properties of all components in order to obtain materials with strong antimicrobial activity against different strains, usually associated with nosocomial infection outbreaks brought on by MDRO after ERCP procedures. The systems are based on reversible imine linkages, which are kept together in a three-dimensional network by hydrophobic clusters formed through hydrophilic/hydrophobic segregation, forming a matrix in which ZnO–NPs can be obtained [37,41,42].

3.1. Structural Characterisation by ¹H-NMR and FTIR Spectroscopy

The structural characterisation of the hydrogels was first carried out using ¹H-NMR spectroscopy, through comparison with chitosan. The chitosan spectrum presented the characteristic chemical shifts according to its chemical structure: at 3.1 ppm, the H2 from the D-glucoamine unit appeared, and between 3.4 and 4.2 ppm, superposed signals corresponding to the H3–H6 protons from the D-glucosamine unit appeared. Due to the water suppression, the H1 signal was not present in the spectrum (Figure 1a). In the spectra of the S1.5 and S2 hydrogels, besides the signals from chitosan, a new peak appeared at 8.5 ppm, corresponding to the imine proton. Moreover, the chemical shift corresponding of the unreacted aldehyde was also present as an intense peak at 9.9 ppm. This can be explained by the fact that the imine formation is an equilibrium reaction which depends mainly on the aldehyde’s reactivity. Previous studies on iminoboronate chitosan hydrogels demonstrated the farily low reactivity of 2-FPBA, indicating its incomplete consumption in the imination reaction with chitosan. In this particular case, very probably because of the presence of ZnO–NPs, the imine formation was hampered, with the imine to aldehyde integrals ratio being lower than in the absence of ZnO–NPs [36,37].

FTIR spectroscopy was used as a complementary method for the structural characterisation of the obtained systems. The FTIR spectrum of the ZnO–NPs presented a similar pattern to those found in the literature, and the bands that appeared in the finger print region, between 1400 and 600 cm⁻¹, were characteristic to ZnO [43]. Moreover, the broad band with two maxima at 3545 and 3385 cm⁻¹ (Figure 1) represents the OH stretching vibrations due to the absorbed water molecules on the surface of ZnO–NPs. Regarding the FTIR spectrum of the R sample, it presents some differences in terms of bands positions and intensities, compared to those in ZnO, indicating strong interactions between chitosan and the nanoparticles. The structural characterisation of the xerogels by FTIR revealed a new band at 1625 cm⁻¹, indicating the formation of imine linkages between chitosan and boronic aldehyde. The band is broad due to being superposed with the amide band from chitosan [36]. The changes that appeared in the 3700–2700 cm⁻¹ spectral region are due to the fact that the signals are shifted to lower wavenumber as a consequence of chitosan’s presence, and the band in that region corresponds to the amino and hydroxyl stretching vibrations from its structure [44].

FTIR spectroscopy was used as a complementary method for the structural characterisation of the obtained systems. The FTIR spectrum of the ZnO–NPs presented a similar pattern to those found in the literature, and the bands that appeared in the finger print region, between 1400 and 600 cm⁻¹, were characteristic to ZnO [43]. Moreover, the broad band with two maxima at 3545 and 3385 cm⁻¹ (Figure 1) represents the OH stretching vibrations due to the absorbed water molecules on the surface of ZnO–NPs. Regarding the FTIR spectrum of the R sample, it presents some differences in terms of bands positions and intensities, compared to those in ZnO, indicating strong interactions between chitosan and the nanoparticles. The structural characterisation of the xerogels by FTIR revealed a new band at 1625 cm⁻¹, indicating the formation of imine linkages between chitosan and boronic aldehyde. The band is broad due to being superposed with the amide band from chitosan [36]. The changes that appeared in the 3700–2700 cm⁻¹ spectral region are due to the fact that the signals are shifted to lower wavenumber as a consequence of chitosan’s presence, and the band in that region corresponds to the amino and hydroxyl stretching vibrations from its structure [44].

3.2. Supramolecular Characterisation via Wide-Angle X-ray Diffraction

The supramolecular arrangement of the imino-chitosan xerogels containing ZnO was investigated via WXRD (see Figure 2), through comparison with the reference samples, ZnO, and R. The pattern of the ZnO is similar to that found in the literature, indicating the high purity of the obtained nanoparticles [45], indexed within the hexagonal ZnO wurtzite, according to Joint Committee on Powder Diffraction Standards (JCPDS) card no. 36–1451 [46]. The diffraction peak from 36.2 two theta degrees was used in order to evaluate the size of the crystallites by applying the Debye–Scherrer Equation [45]:

$$\mathrm{D}=\frac{\mathrm{K}\times \mathsf{\lambda }}{\mathsf{\beta }\times \cos \mathsf{\theta }} ,$$

where D is the NP crystalline size, K is the Scherrer constant (0.98), λ is the wavelength of Copper-K radiation (1.5406 Å), and β represents the full width at the half maximum (FWHM) of the diffraction peak. The average size of the ZnO–NPs was determined to be 78 nm.

The diffractogram of the R sample presented three broad diffraction peaks, at 10.7, 22, and 39 two theta degrees, corresponding to the distances of 8.2, 4.1, and 2.4 Å. The first two peaks are from chitosan, while the last peak is from ZnO, being more shifted than the signals of the pure ZnO, indicating significant changes in the oxide supramolecular architecture, due to their interactions with the chitosan backbone. The diffractograms of the xerogels containing ZnO presented a similar pattern with previously reported xerogels based on iminoboronate chitosan derivatives [41], with a difference that also appeared in R’s diffractogram: a sign of the presence of ZnO in the hydrogel matrix.

The high degree of ordering of the samples was also confirmed through POM (Figure 2). The ZnO–NPs, due to their small size, were not clearly observed with the optical microscope, but their agglomeration caused birefringence due to their crystalline nature (Figure 2). All the other samples presented strong birefringence under polarised light due to their highly ordered structure at a supramolecular level, as was also evidenced by XRD.

3.3. Morphological and Elementary Characterisation by SEM and EDAX

The morphology of the samples was investigated via scanning electron microscopy. The ZnO powder was also investigated, confirming that nanoentities with regular shape and a mean diameter of ~80 nm were obtained. All the samples presented a highly porous microstructure with pores of micrometric sizes. A porous morphology was also obtained for the reference sample (R) by adding ZnO to the chitosan solution without using 2-FPBA for hydrogelation. This sample had the largest pores, around 40 μm, and a lacunarity of 73%. In comparison with the reference sample, the hydrogels containing ZnO had much lower pore sizes, around 10 μm, and a lacunarity that was dependent on the aldehyde content from 53% for S1 to 67% for S2. Another important observation after SEM investigation was the fact that no ZnO aggregates were observed in the microphotographs confirming the very fine distribution of the ZnO into the hydrogels (Figure 3).

To attest the presence of the ZnO in the hydrogels, the samples were submitted to energy-dispersive X-ray analysis (EDAX). As can be observed, in the R sample, containing chitosan and ZnO as well as the chemical elements from chitosan (C, H and N), Zn also appeared, due to the presence of ZnO. The same also occurred in the case of the hydrogels, where, alongside the elements from chitosan and 2-FPBA (C, O, and B), zinc also appeared (Figure 4).

3.4. Antioxidant Activity

The antioxidant activity of the developed hydrogel composites was evaluated using the DPPH assay and compared with those of the reference samples, namely ZnO–NPs, chitosan, and a chitosan–ZnO mixture (Figure 5). The experiments were carried out on the reference samples at the same concentration as those that were used in the formulations. As can be observed from the experimental results, the ZnO–NPs presented a very weak inhibition ability of only ~8%, very probably due to the very low concentration, while chitosan presented a quite high radical scavenging capacity of ~56%. The combination of chitosan with ZnO–NPs (R) presented an intermediate value of 44%, indicating interactions between the components which prevented chitosan from exerting its full antioxidant potential. However, the formulations, based on iminoboronate hydrogels and ZnO–NPs, presented a high antioxidant activity, with a maximum of 66% for the sample S1. The colour of the DPPH solution changed in the presence of the analysed samples from deep purple to yellow/colourless/light purple, the colour shift being directly correlated with the antioxidant capacity of each sample (Figure 5).

All these data show that the hydrogels present a fairly high radical scavenging potential, which may play an important role in the antimicrobial bioapplications of the developed systems.

3.5. Antimicrobial Activity

The diagrams (Figure 6a–c) show the growth inhibition of the microorganism’s colonies when treated with the hydrogel samples and with the controls. Distinct activities against the reference tested strains were noticed (Figure 6d–f). In case of the Gram-positive bacterial strain E. faecalis, the cell viability decreased to 0% after 2 h of incubation for samples S1 and S1.5 and to 4% for sample S2 (Figure 6a). In case of the controls, a very reduced antibacterial activity of ZnO and BA was noticed, but an increased activity in the case of Ch and R leading to a synergism was observed in the case of the hydrogels. This type of synergism was noticed for all the tested microorganism strains.

In the case of the Gram-negative bacterial colonies of K. pneumoniae, it is very clear that the controls collaborate to obtain the antimicrobial activity of the three types of hydrogels. For sample S1 and S1.5, it took 6 h for all the bacterial cells to be destroyed, while after the same period of incubation, sample S2 reduced the bacterial populations to almost 1%. In this particular case, R also has a very good antibacterial activity, destroying up to 98% of the cells after 6 h of incubation. ZnO and Ch also display notable activity, reducing cell viability by 90% in 6 h.

In the third case, the yeast cells are completely destroyed by the hydrogels after 24 h of incubation. The same activity was also noticed in controls, i.e., BA and R, leading to the same synergism that was previously observed. ZnO had smaller antifungal activity, and Ch was proven to provide an important contribution to the results, destroying up to 75% of the yeast cells after 24 h of incubation.

Microbial cell viability decreased for all the samples in a time-dependent manner. Significantly different cell viabilities among the time points were found (detailed statistics can be seen in

Table 2. Detailed statistics of the antimicrobial activity of the samples against: (a) E. faecalis; (b) K. pneumoniae; (c) C. albicans.

(a) Incubation Time	Relative Cell Viability (Mean ± SD)
	Ch	BA	ZnO	R	S1	S1.5	S2
10 min	42.10 ± 0.22 ***	99.48 ± 0.68 ***	99.75 ± 0.28 ***	72.25 ± 2.01 ***	16.98 ± 1.37 ***	10.60 ± 0.50 ***	27.36 ± 0.99 ***
20 min	24.89 ± 1.01 ***	98.58 ± 0.94 ***	99.66 ± 0.58 ***	9.45 ± 0.86 ***	8.43 ± 0.70 ***	7.99 ± 0.40 ***	18.43 ± 0.70 ***
40 min	9.32 ± 0.71 ***	95.23 ± 1.06 ***	86.96 ± 2.76 ***	3.08 ± 0.95 ***	4.70 ± 0.55 ***	1.52 ± 0.26 ***	14.52 ± 0.86 ***
1 h	11.61 ± 0.91 ***	86.82 ± 1.52 ***	79.45 ± 0.87 ***	2.82 ± 0.34 ***	4.46 ± 0.60 ***	1.17 ± 0.26 ***	8.02 ± 0.27 ***
2 h	13.23 ± 0.22 ***	90.94 ± 3.29 ***	82.42 ± 1.04 ***	2.11 ± 0.10 ***	0.66 ± 0.20 ***	0.00 ± 0.00 ***	4.34 ± 0.67 ***
3 h	12.58 ± 0.64 ***	75.84 ± 1.58 ***	88.48 ± 1.12 ***	2.04 ± 0.22 ***	0.23 ± 0.24 ***	0.00 ± 0.00 ***	2.00 ± 0.13 ***
6 h	4.49 ± 0.60 ***	76.87 ± 2.83 ***	87.78 ± 0.75 ***	0.66 ± 0.21 ***	0.00 ± 0.00 ***	0.00 ± 0.00 ***	0.00 ± 0.00 ***
24 h	0.49 ± 0.29 ***	70.59 ± 1.12 ***	79.63 ± 0.74 ***	0.00 ± 0.00 ***	0.00 ± 0.00 ***	0.00 ± 0.00 ***	0.00 ± 0.00 ***
(b) Incubation Time	Relative Cell Viability (Mean ± SD)
	Ch	BA	ZnO	R	S1	S1.5	S2
10 min	86.66 ± 3.16 ***	99.75 ± 0.30 ***	94.95 ± 1.72 ***	67.70 ± 1.13 ***	86.71 ± 3.39 ***	71.28 ± 1.26 ***	87.30 ± 1.47 ***
20 min	84.19 ± 0.86 ***	97.63 ± 1.06 ***	97.96 ± 2.54 ***	64.89 ± 1.56 ***	84.79 ± 0.58 ***	60.36 ± 0.97 ***	79.27 ± 0.88 ***
40 min	79.51 ± 0.97 ***	87.73 ± 0.90 ***	98.09 ± 0.35 ***	39.62 ± 1.01 ***	13.62 ± 0.85 ***	22.62 ± 1.26 ***	42.56 ± 2.06 ***
1 h	57.73 ± 1.99 ***	84.13 ± 1.35 ***	95.93 ± 0.70 ***	26.42 ± 1.18 ***	9.66 ± 0.41 ***	11.43 ± 0.84 ***	42.23 ± 2.86 ***
2 h	28.08 ± 1.77 ***	40.48 ± 0.80 ***	93.23 ± 2.89 ***	13.09 ± 0.65 ***	8.12 ± 0.79 ***	9.40 ± 0.67 ***	24.64 ± 1.01 ***
3 h	25.10 ± 0.97 ***	25.26 ± 0.46 ***	69.44 ± 1.19 ***	11.30 ± 1.21 ***	5.23 ± 1.21 ***	7.73 ± 1.02 ***	19.11 ± 0.87 ***
6 h	9.49 ± 0.88 ***	18.28 ± 1.70 ***	8.96 ± 0.22 ***	1.18 ± 0.29 ***	0.00 ± 0.00 ***	0.07 ± 0.12 ***	1.50 ± 0.45 ***
24 h	0.21 ± 0.11 ***	4.96 ± 0.48 ***	1.84 ± 0.65 ***	0.00 ± 0.00 ***	0.00 ± 0.00 ***	0.00 ± 0.00 ***	0.00 ± 0.00 ***
(c) Incubation Time	Relative Cell Viability (Mean ± SD)
	Ch	BA	ZnO	R	S1	S1.5	S2
10 min	86.92 ± 1.60 ***	79.82 ± 2.49 ***	93.34 ± 0.95 ***	99.52 ± 0.70 ***	72.57 ± 1.34 ***	63.34 ± 2.95 ***	85.31 ± 4.89 ***
20 min	48.53 ± 1.52 ***	77.98 ± 2.54 ***	92.54 ± 1.58 ***	93.54 ± 1.70 ***	44.56 ± 0.97 ***	60.20 ± 1.13 ***	93.13 ± 3.36 ***
40 min	44.51 ± 3.67 ***	79.57 ± 1.48 ***	92.13 ± 2.63 ***	74.13 ± 0.31 ***	39.27 ± 0.83 ***	59.54 ± 1.25 ***	91.20 ± 1.24 ***
1 h	42.12 ± 4.29 ***	82.80 ± 2.51 ***	76.14 ± 2.30 ***	71.24 ± 1.39 ***	45.93 ± 2.45 ***	58.33 ± 2.17 ***	88.38 ± 1.41 ***
2 h	45.54 ± 5.62 ***	70.99 ± 0.74 ***	70.36 ± 0.68 ***	55.44 ± 1.14 ***	40.30 ± 0.91 ***	56.38 ± 3.48 ***	78.29 ± 2.55 ***
3 h	48.35 ± 2.79 ***	65.36 ± 4.83 ***	72.54 ± 2.50 ***	48.34 ± 1.18 ***	38.67 ± 1.82 ***	41.35 ± 1.06 ***	69.90 ± 0.20 ***
6 h	28.53 ± 2.25 ***	34.21 ± 0.93 ***	70.57 ± 0.76 ***	35.52 ± 2.42 ***	38.90 ± 3.65 ***	34.66 ± 0.65 ***	34.47 ± 3.95 ***
24 h	23.36 ± 2.93 ***	0.00 ± 0.00 ***	68.22 ± 2.00 ***	0.00 ± 0.00 ***	0.00 ± 0.00 ***	0.00 ± 0.00 ***	0.00 ± 0.00 ***

*** p < 0.0001.

a–c). These results suggested that the microbial cells were damaged by hydrogels and that their toxicity was time-dependent. It was determined that the optimal period of time for the present study is 6 h. According to the two-way ANOVA, a statistically significant (p < 0.0001) difference was found between the species when cell viability was considered. Furthermore, the process included the evaluation of the influence of the type of sample and incubation time on the viability or cells. The most significant effect on the cell viability was produced by the incubation time, whereas the material type had less effect.

From the SEM images (Figure 6g–i) and AFM images (Figure 6j–l), it is obvious that the coating material of the duodenoscope exhibits degradation and erosion marks brought on by heavy use and frequent reprocessing. However, the material preserves its anti-adherence properties, but a few very damaged bacterial cells of E. faecalis (Figure 6f), K. pneumoniae (Figure 6g), and C. albicans (Figure 6i) were observed on the tested surfaces after the incubation with the hydrogels, especially with S1, which had the strongest antimicrobial activity of all the tested microbial strains. The same results are also presented by the AFM images, showing that after the incubation of hydrogel S1, there were only a few cells left on the surface of the material, with the hydrogels preventing cell adhesion and biofilm formation.

Figure 6j–l shows the AFM topographic images of the duodenoscope samples before and after incubation with the microbial strains and hydrogels. The samples are characterised using an inhomogeneous morphology, as has been previously observed in an optical study carried out on similar duodenoscopes [47]. AFM was also used to investigate whether biofilms were formed on the duodenoscope samples treated with hydrogel after incubation with K. pneumoniae (Figure 6k) and E. faecalis (Figure 6l). For both bacterial strains, no colonies or aggregates were found on the tested surfaces. In the case of K. pneumoniae, the attachment of rare rod-shaped cells was noticed. For E. faecalis, small round-shaped cells (with diameter of 0.7 μm) individually distributed on the solid surface were observed. Although the presence of few bacterial cells was confirmed, the bacteria left behind following hydrogel treatment failed to grow after incubation, suggesting that they had been destroyed by the hydrogels.

Additionally, from the SEM images, it can be noted that the hydrogels not only drastically reduced cell density and viability but also did not allow biofilm formation. It is known that, once formed, the presence of a biofilm on microorganisms protects them from outside influences, such as disinfectants and antibiotics [48]. A biofilm is an accumulation of microorganisms on a surface enclosed in a matrix of exopolysaccharides [49,50]. Additionally, it must be taken into account that there is a notable difference between “contamination”, meaning the presence of microorganisms on a duodenoscope, and “infection”, meaning microorganisms infiltrating human tissue or producing clinical disease; these are not synonymous. Duodenoscopes infected with specific microorganisms may result in infection. Not all organisms are harmful, and it is not always apparent how frequently they cause infections or what kind of damage they do. That being said, our results proved that generally, after 6 h of the incubation of the hydrogels, there was no chance of infection or contamination on the surface of the external polymers of the duodenoscope.

Our previous studies proved that iminoboronate–chitosan hydrogels based on 2-FPBA have fungicidal activity against various yeast strains such as C. albicans, C. glabrata, and C. parapsilosis [36,37]. Reports on formyl phenyl boronic acid (FPBA) showed its antimicrobial effects against S. aureus, P. aeruginosa, Aspergillus sp., Fusarium sp., Penicillium sp., and Candida sp. due to the presence of two hydroxyl groups [49,50,51]. In this particular case, BA was proven to destroy up to 85% of the K. pneumoniae cells and 75% of C. albicans cells after 6 h of incubation. BA was not very efficient against E. faecalis, destroying only 30% of the bacterial cells after 24 h of incubation.

Positive charges from the amino groups of chitosan interact electrostatically with negatively charged components on the microbial membrane, resulting in the disruption of cell membrane [52] or the blocking of the exchange of nutrients by covering porins [53], penetrating the cell wall to affect DNA/RNA and protein synthesis [54]. The antimicrobial effect of Ch depends on its molecular weight and varies among different microorganisms. This study proved that Ch destroys up to 90% of K. pneumoniae and up to 70% of C. albicans after 6 h of incubation. As in the case of BA, chitosan was not very efficient by itself against E. faecalis.

Usually, ZnO–NPs display antimicrobial activities via multiple mechanisms: interactions with the phospholipid bilayer, binding to cytosolic proteins, or the formation of reactive oxygen species (ROS), which produce damages to the phospholipids in the membranes, lipoproteins, and nucleic acids, causing oxidative stress and eventually destroying the microorganisms [55]. Additionally, the mechanism of ZnO–NPs implies the release of Zn²⁺ into the media, which causes bacterial damage through active transport inhibition, amino acid metabolism, and the disruption of an enzyme system [38]. The literature presents examples of ZnO–NPs with a higher antibacterial activity against Gram-negative strains than against Gram-positive strains [56,57,58]. Generally, it is thought that Gram-negative bacteria are more susceptible than Gram-positive to attack from external factors due to their cell wall composition [59]—in the case of Gram-negative bacteria, bacterial cells are covered by a layer of lipopolysaccharides (1–3 μm thick) and thin peptidoglycans (~8 nm thick), whereas Gram-positive bacteria have a peptidoglycan layer (~80 nm) thick) with covalently attached teichoic and teichuronic acids. Our results are similar to those presented in the literature, with ZnO–NPs being more efficient against K. pneumoniae (killing up to 90% of the bacterial cells after 24 h of incubation) than against E. faecalis (destroying up 20% of the bacterial cells after 24 h of incubation). ZnO–NPs also displayed limited antifungal activity, with 70% of cells being viable after the same period of incubation.

Recently, chitosan hybrid materials with metal NPs and oxide agents have been developed with the excellent properties of individual components and outstanding simultaneous synergistic effects [60]. Due to the presence of amine and hydroxyl groups on chitosan, the ZnO–chitosan complex is currently attracting great interest regarding its potential use as an antimicrobial agent [61]. The presence of ZnO–NPs in combination with chitosan as a natural compound that strongly complexes with metal oxide NPs has led to the increase in chitosan efficiency against Campylobacter sp., M. luteus, and S. aureus [61,62]. In this study, it was also noticed that ZnO–chitosan complex was very efficient against the tested reference strains after 6 h of incubation.

The antimicrobial activity determined for the ZnO–NPs is lower than that determined for the ZnO–chitosan complex, perhaps because chitosan acts as a barrier to the migration of ZnO to the medium and allows ZnO–NPs to be in direct contact with the bacterial cell surface, generating ROS and leading to cell death. The antimicrobial activity of the hydrogels is higher than that of the controls, and accumulates the activity of the precursors, their activity being significantly correlated with the incubation time.

4. Conclusions

In conclusion, both the structural and morphological features of the hydrogels contribute to their antimicrobial activity. The hydrogels presented very good antimicrobial activity against the different classes of microorganisms represented by a Gram-positive bacterial strain (E. faecalis), a Gram-negative bacterial strain (K. pneumonia), and a yeast strain (C. albicans). At the same time, the hydrogels limited the microbial adherence to the polymer coating of the duodenoscope. With these remarkable properties, the present hydrogels may be used in the prevention of bacterial biofilm development on duodenoscopes, as well as for the design of new antiseptics and disinfectants with efficient protective actions against the microorganism colonisation of inert surfaces.