Silver Nanoparticle-Infused Pullulan Films for the Inhibition of Foodborne Bacteria

Abstract

The aim of this research was to examine the antibacterial activity of commercially available silver nanoparticles against foodborne bacteria and to evaluate the properties of pullulan films incorporating these nanoparticles, including their antibacterial activity and selected physical properties. First, the antibacterial activity of silver nanoparticles against foodborne bacteria was investigated. The following parameters were assessed to evaluate the antibacterial activity of silver nanoparticles: minimum inhibitory concentration (MIC), minimum bactericidal concentration (MBC), percentage antibacterial activity, bacterial survival based on time–kill curves, leakage of DNA and intracellular proteins using spectrophotometric measurements, and changes in bacterial cell morphology using scanning and transmission electron microscopy (SEM and TEM). Pullulan films with silver nanoparticle content ranging from 2 to 32 µg/cm² were obtained. The films were evaluated for antibacterial activity and physical properties, including macroscopic and microstructural (SEM) observations, thickness, light barrier, and color. Silver nanoparticles at a concentration of 25 µg/mL achieved 100% inhibition of the test bacteria, with destruction of bacterial cells after 3 or 6 h of incubation, depending on the silver nanoparticle concentration. Incorporation of silver nanoparticles into pullulan films, even in lower amounts, resulted in an antibacterial effect. All films had a compact and uniform microstructure and were shiny and flexible. Analysis of variance showed a significant (p < 0.05) effect of the addition of silver nanoparticles on the thickness, transparency, and color of the films. The obtained pullulan films containing silver nanoparticles were characterized by strong inhibitory activity against foodborne bacteria, had a brown color of varying intensity, a uniform microstructure, a smooth surface, and were barriers to UV radiation and visible light.

1. Introduction

Silver and its compounds have been a subject of interest for thousands of years due to their antiseptic properties [1]. Nowadays, silver nanoparticles (AgNPs) have received particular attention because of their unique physicochemical characteristics, including optical, electrical, thermal, and biological properties [2,3]. Silver nanoparticles possess strong antibacterial activity; therefore, they are the subject of many studies. Silver nanoparticles are highly toxic to many Gram-positive and Gram-negative bacteria, including drug-resistant pathogens [4,5,6]. The bactericidal effect of silver nanoparticles depends on many factors, including the size of the nanoparticles [7], shape [8], and concentration [5]. Because of antimicrobial activity properties, the nanoparticles were used in biomedical and industrial products. For example, AgNPs are employed in medical devices, catheters, wound dressings, dental materials, and implants to prevent microbial contamination [3,9]. Silver nanoparticles can potentially be used as a preservative in cosmetics, thus reducing the use of parabens [10]. Moreover, silver-containing materials are applied to textiles or food packaging materials [11,12].

Recently, one important direction has involved the use of biopolymers as stabilizing and dispersing agents for AgNPs to achieve a nanocomposite material with antibacterial properties. Biopolymers not only enhance nanoparticle homogeneity and structural stability but also preserve strong antimicrobial activity within nanocomposite films [13,14]. Such an approach provides new opportunities in the design of multifunctional materials that combine antimicrobial protection with biocompatibility, enabling their applications in biomedicine, healthcare, and food packaging materials [15,16]. Examples of bio-nanocomposites include gelatin/Ag nanocomposite film [17], chitosan-based nanocomposite films incorporated with AgNPs [18,19], starch-based nanocomposite films with silver [20], and cellulose film and its derivatives loaded with AgNPs [21,22,23]. Nanocomposites are characterized by improved stability, increased resistance to nanoparticle agglomeration, and high antibacterial effectiveness, which makes them promising materials for a wide range of applications.

Pullulan is a polysaccharide of microbial origin, produced by fermentation from a food-grade hydrolyzed starch by the fungus Aureobasidium pullulans. This compound is composed of repeating maltotrioses linked by α-(1,6)-glycosidic bonds. Pullulan is a non-hygroscopic, biodegradable, tasteless, and odorless biopolymer approved by the European Food Safety Authority as a food additive (E1204) and has GRAS status [24,25,26,27,28,29]. Pullulan dissolves well in water, and after drying, thin, flexible, transparent, colorless, edible and biodegradable films are obtained from aqueous solutions [30]. This feature makes pullulan a versatile material for various packaging applications, such as films, coatings, capsules, and microcapsules [24]. The use of pullulan film and capsules to deliver nutraceuticals and pharmaceuticals has been proposed [31]. Pullulan films and coatings are applied directly to food to extend the shelf life of food products. Pullulan films have favorable mechanical, optical, and barrier properties for gases, including oxygen [32,33]. In order to obtain pullulan films with antimicrobial activity, extracts, plant essential oils, and bacteriocins are included [34,35,36,37,38]. Growing interest in this biopolymer is also due to its ability to fabricate metal nanoparticles. It has been demonstrated that pullulan can serve as a reducing and stabilizing agent for the synthesis of AgNPs. Kanmani and Lim [39] demonstrated that pullulan effectively reduces silver ions and stabilizes the formed nanoparticles, preventing their aggregation. Using FTIR analysis, the authors confirmed the involvement of hydroxyl functional groups in pullulan in the reduction and stabilization processes. However, some differences were observed in the reported nanoparticle sizes. While TEM analysis showed rod, spherical, and oval-shaped AgNPs with diameters ranging from 2 to 40 nm, dynamic light scattering (DLS) measurements indicated a broader, multimodal distribution between 5.6 and 97.6 nm, suggesting a heterogeneous population of nanoparticles. Similarly, Burduniuc et al. [40] also confirmed the ability of pullulan to act as a reducing and stabilization agent. Authors showed that the hydroxyl group of pullulan reduces Ag+ to Ag0 and stabilizes particles, providing steric stabilization. Obtained AgNPs presented irregular shapes of different sizes. Moreover, pullulan-capped AgNPs maintain stability and prevent aggregation. It has been demonstrated that the hydroxyl groups of pullulan form a coating around the nanoparticles, providing steric protection against aggregation and ensuring the stability of pullulan-stabilized AgNPs for up to six months [41]. Pullulan is not only used as an agent to fabricate AgNPs, but also, in the form of pullulan film, can be used as a carrier for this compound. Lee et al. [42] showed that silver nanoparticles obtained using pullulan were then successfully incorporated into pullulan-based nanocomposite films. It was reported that the addition of AgNPs had an influence on the physical and mechanical properties of the prepared films. While color intensity, transparency, moisture content, water vapor barrier properties, hydrophobicity, thickness, and elongation at break of the films were significantly increased, tensile strength and elastic modulus of the films were decreased. Additionally, film characterization using FTIR confirmed that the hydroxyl groups of pullulan exhibit a strong affinity for Ag ions, resulting in good compatibility between AgNPs and the biopolymer. Pullulan film with silver nanoparticles also presents good antimicrobial properties. For example, a pullulan-based film with nanosilver destroyed spores and reduced the viability of the fungus Aspergillus niger [43], and also presents activity against bacteria (E. coli, Salmonella sp., Klebsiella pneumonia, P. aeruginosa, S. aureus, L. monocytogenes) [44]. In turn, Khalaf et al. [45] and Morsy et al. [46] demonstrated that incorporating AgNPs into pullulan film effectively inhibits the growth of S. aureus and L. monocytogenes, suggesting that this nanocomposite material has the potential to improve the microbial quality of meat and poultry products. Additionally, Qi et al. [47] investigated the antimicrobial properties of chitosan/pullulan composite films reinforced with AgNPs. It was demonstrated that the tested nanomaterials effectively reduce the infection of exogenous microorganisms, prevent nutrient loss, and prolong the preservation period and shelf life of litchi. Based on this data, nanocomposite pullulan film shows promising potential as a food packaging material.

Previous studies have focused on nanoparticles synthesized at the laboratory scale, which may vary in their physicochemical characteristics depending on preparation methods and conditions. Our study focuses attention on the use of commercially available silver nanoparticles. Commercial silver nanoparticles are manufactured under controlled and standardized conditions, ensuring consistency from batch to batch. In this context, this approach provides improved reproducibility in the industrial-scale production of nanomaterials. Furthermore, the results obtained for commercial nanoparticles may serve as a useful reference for validating laboratory-synthesized nanoparticles. Harun-Ur-Rashid et al. [48] reported that integrating AgNPs into polymer matrices has significant potential for use in commercial biomedical products. However, as the authors noted, optimizing the fabrication of AgNPs provides precise control over nanoparticle size, shape, dispersion, and stability, thereby ensuring scalability for industrial production. Dos Santos et al. [49] demonstrated that commercial AgNPs were more effectively absorbed and adhered to textile fibers, indicating greater stability within the material during washing cycles compared to nanoparticles synthesized specifically for their study. Shan et al. [50] also reported that the use of commercial nanoparticles allowed the formulation of a paste characterized by a low sintering temperature, high bond strength, and excellent electrical and thermal conductivity, ensuring reliable performance and batch-to-batch reproducibility. Additionally, silver pastes used as a bonding adhesive designed for devices requiring high thermal and electrical efficiency, enabled effective heat transfer from electronic components.

The aim of this research was to examine the antibacterial activity of commercially available silver nanoparticles against foodborne bacteria and to evaluate the properties of pullulan films incorporating these nanoparticles, including their antibacterial activity and selected physical properties. The MIC and MBC of silver nanoparticles, time–kill curves of the test bacteria, cellular morphological changes, and cell content leakage under the influence of AgNPs were determined. Pullulan films with silver nanoparticles at various concentrations were then prepared. The antibacterial activity of the films was determined using the disk diffusion method, and their selected physical properties were determined. The macroscopic appearance, microstructure visualized in SEM and TEM images, thickness, transparency under visible light, and color were described.

2. Materials and Methods

2.1. Materials

A commercial colloidal solution of silver nanoparticles (AgNPs), HydroSilver 1000, was supplied by Amepox (Łódź, Poland). The formulation contains AgNPs of 50–60 nm with a protective shell of polyvinyl alcohol (PVA) and silver concentration in the liquid of 1000 ppm. HydroSilver 1000 is characterized by high purity of ingredients and the following technical parameters: a translucent, opalescent liquid, viscosity 19 ± 2 mPa·s, specific gravity 1.0 ± 0.2 g/mL. Pullulan was provided by Hayashibara Biochemical Laboratories Inc. (Okayama, Japan).

Tryptic soy broth (TSB), Tryptic soy agar (TSA), Mueller-Hinton Broth (MHB), and Mueller-Hinton Agar (MHA) were purchased from BTL (Łódź, Poland). NaCl and glycerol were from Chempur (Piekary Śląskie, Poland), phosphate-buffered saline (PBS) was obtained from VWR International (Solon, OH, USA), and resazurin was from PoL-Aura (Olsztyn, Poland). A commercial disk with antibiotic azlocillin (Azlocillin 75, Mast Diagnostics, Bootle, UK) was used to determine the susceptibility of bacteria.

Nine foodborne bacteria were investigated: Gram-negative: Escherichia coli ATCC 13067, Klebsiella pneumoniae ATCC 13883, Pseudomonas aeruginosa ATCC 27853, Salmonella enterica subsp. enterica ser. Enteritidis ATCC 13076, Shigella sonnei NIPH–NIH “s”, and Gram-positive: Staphylococcus aureus ATCC 25923, Enterococcus faecalis ATCC 29212, Bacillus cereus ATCC 11778, and Listeria innocua ATCC 33090. The strain S. sonnei NIPH–NIH “s” is a clinical isolate from the National Institute of Public Health–National Institute of Hygiene (Warsaw, Poland). L. innocua is a surrogate of the pathogenic species L. monocytogenes [51].

2.2. Bacterial Culture and Inoculum Preparation

The microorganisms were stored frozen at −80 °C. They were incubated on TSB medium at 37 °C to activate the bacteria for 24 h. Afterward, the bacteria were transferred and spread onto Petri dishes containing TSA medium and cultured at 37 °C for 24 h. Then, bacteria were harvested and suspended in saline solution (0.85% NaCl). The cell suspension was standardized using a Densimat (bioMérieux, Marcy-l’Étoile, France) to 0.5 McFarland, corresponding to approximately 1 × 10⁸ cells/mL. The bacterial inoculum was diluted appropriately and used for further studies.

2.3. Determination of MIC and MBC of AgNPs

The minimum inhibitory concentration (MIC) was determined using a microdilution assay according to the CLSI (Clinical and Laboratory Standards Institute) protocol [52]. Initially, a 10% stock solution of silver nanoparticles was prepared in Mueller-Hinton Broth (MHB) medium. In the next step, 100 µL of the silver nanoparticle stock solution was added to each well in the first column of a 96-well plate, while the wells in adjacent columns were filled with 50 µL of MHB. To establish a series of double dilutions, 50 µL of the nanoparticle solution was transferred from the first column to the second column, and this dilution process was continued through the subsequent columns. Then, 50 µL of bacterial inoculum was added to each well, resulting in an inoculum concentration in each well of approximately 5 × 10⁵ cells/mL. Positive controls (bacteria inoculum in MHB medium without silver nanoparticles) and negative controls (MHB medium with silver nanoparticles) were also prepared. The plate was incubated at 37 °C for 24 h. Then, 10 µL of 0.015% (w/v) resazurin solution (color indicator to assess cell viability) was added to each well, followed by incubation at 37 °C for 2 h. Viable bacterial cells cause resazurin reduction and change color from blue to pink. The MIC was defined as the lowest concentration of silver nanoparticles at which no observable color change occurred in the indicator.

To assess the MBC (minimum bactericidal concentration), 10 μL of the content of wells showing no indicator color change was transferred onto MHA plates. These plates were then incubated at 37 °C for 24 h. The MBC was defined as the lowest concentration of AgNPs that reduced the initial inoculum by 99.9%.

Additionally, the following classification based on the MBC/MIC ratio was used to determine whether an antibacterial agent is bactericidal or bacteriostatic. An antibacterial agent is considered bactericidal if the MBC/MIC ratio is ≤4 and bacteriostatic if the MBC/MIC ratio value is >4 [53].

The percentage of inhibition was calculated using Equation (1):

$$Percentage of inhibition \left(\%\right)={\displaystyle \frac{Number of inhibited strains at concentration corresponding to MIC}{Total number of strains tested}}\times 100$$

(1)

2.4. Time–Kill Assay

The time–kill curve assay was performed using the agar plate viable count method to evaluate the inhibitory effect of silver nanoparticles (AgNPs) on bacterial growth. Bacterial suspensions were standardized to approximately 1 × 10⁵ CFU/mL in MHB, to which AgNPs were added at concentrations corresponding to ½ × MIC, 1 × MIC, and 2 × MIC. Control samples containing only MHB without AgNPs were also prepared to serve as negative controls. All samples were incubated at 37 °C for 24 h. Bacterial viability was assessed by taking samples at time intervals of 0, 1, 3, 6, and 24 h, followed by serial dilutions and plating on MHA cultivation media. The plates were incubated at 37 °C for 24 h, followed by enumeration of bacterial colonies using an automated colony counter (ProtoCol3, Synbiosis, Cambridge, UK). Bacterial counts were expressed as log CFU/mL. The time–kill curve was generated by plotting log (CFU/mL) against tested time intervals. All experiments were performed in triplicate.

The bactericidal effect of silver nanoparticles was determined as a reduction by ≥3 log CFU/mL in the viable bacterial count compared to the initial inoculum [54].

2.5. Determination of Nucleic Acid and Protein Leakage

Leakage of nucleic acid and proteins from bacterial cells exposed to AgNPs was determined as described previously with slight modification [55].

Selected bacterial strains (E. coli ATCC 25922, S. Enteritidis ATCC 13076, S. sonnei NIPH–NIH “s”, S. aureus ATCC 25923, E. faecalis ATCC 29212, L. innocua ATCC 33090) were cultivated in MHB at 37 °C for 18 h. Then, the bacteria were centrifuged at 10,000 rpm (Centrifuge 5804R, Eppendorf, Hamburg, Germany) for 10 min and washed three times with PBS solution. Next, bacteria were resuspended in PBS solution and treated with AgNPs at concentrations corresponding to ½ × MIC, 1 × MIC, and 2 × MIC. The sample without AgNPs served as a control. Cultures were incubated at 37 °C for 6 h on an orbital shaker at 200 rpm. Afterwards, samples were collected and centrifuged at 10,000 rpm for 10 min. The supernatant was examined spectrophotometrically (Metertech UV–VIS SP-8001, Metertech Inc., Taipei, Taiwan) at 260 nm and 280 nm to estimate the amounts of nucleic acids and proteins, respectively, released from the bacterial cells.

2.6. Bacterial Morphology Exposed to AgNPs Observed by SEM and TEM

Morphological changes in the selected bacterial strains (E. coli ATCC 25922, S. Enteritidis ATCC 13076, and S. aureus ATCC 25923) were examined using scanning and transmission electron microscopy (SEM and TEM). Each bacterial culture was individually suspended in PBS containing AgNPs at concentrations equivalent to 1 × MIC. Bacterial suspensions in PBS without AgNPs served as controls. After 24 h of incubation at 37 °C, the samples were centrifuged at 10,000 rpm for 10 min and subsequently prepared for SEM and TEM analysis according to the following procedures.

Sample preparation for SEM: Bacterial cells were fixed with 2.5% glutaraldehyde (Scharlau, Hamburg, Germany) at 4 °C for 24 h, and then dehydrated with a gradient ethanol concentration (25%, 50%, 70%, 90%, and 100%) and finally freeze-dried. Then, samples were coated by sputtering with gold and subjected to observation using SEM under a high vacuum with an accelerating voltage of 20 kV.

Sample preparation for TEM: Bacterial cells were treated with 2.5% glutaraldehyde (Scharlau, Hamburg, Germany) at 4 °C for 2 h, and then fixed with 1% osmium tetroxide (Merck, Darmstadt, Germany) at 4 °C for 1 h, followed by dehydration with a gradient ethanol concentration. Finally, the cells were embedded in Epon 812 resin (Spi-Chem, West Chester, PA, USA) and cut with a diamond knife on an ultramicrotome (LKB Pharmacia, Uppsala, Sweden) to obtain ultrathin sections (70 nm), which were transferred to copper grids. Microscopy was performed using a JEM 1220 TEM microscope (JEOL, Tokyo, Japan) at an electron energy of 120 keV.

2.7. Preparation of Pullulan Film with Colloidal Solution of AgNPs (HydroSilver 1000)

Solvent casting methods were used to prepare pullulan film with AgNPs. Pullulan film-forming solutions were prepared in distilled water by dissolving pullulan (10% w/v) and glycerol (2% w/v). The solution was stirred magnetically at 1000 rpm for 20 min (IKA, Warsaw, Poland) and then sterilized at 121 °C for 15 min. After cooling to room temperature, colloidal solution (HydroSilver 1000, Amepox, Łódź, Poland) was aseptically added at concentrations of 1%, 3%, 5%, 10%, 15%, and 20% (v/v) (

Table 1. Concentration of HydroSilver 1000 colloid in film-forming solution and AgNPs content in 1 mL of film-forming solution and 1 cm² of dried pullulan film.

Films	HydroSilver 1000 [%]	AgNPs [µg/mL]	AgNPs [µg/cm2]
PF	0	-	-
PFHS1	1	10	2
PFHS3	3	30	5
PFHS5	5	50	8
PFHS10	10	100	16
PFHS15	15	150	24
PFHS20	20	200	32

), and the formulation was mixed using a magnetic stirrer. To maintain a constant total volume across all samples, the amount of distilled water used in the pullulan–glycerol solution was adjusted according to the volume of the colloidal AgNP suspension added. The film-forming solution of 10 mL was poured into sterile 90 mm diameter Petri dishes and dried in a laminar cabinet at ambient temperature for 24 h. Dried films were peeled off and conditioned at 22 ± 3 °C and relative humidity (RH) of 50 ± 2% for 48 h before analysis. A total of six pullulan films containing AgNPs were obtained and labeled as PFHS1, PFHS3, PFHS5, PFHS10, PFHS15, and PFHS20, corresponding to HydroSilver 1000 concentrations of 1%, 3%, 5%, 10%, 15%, and 20% (v/v), respectively. A film without silver nanoparticles (PF) was prepared and served as a control. Table 1 presents the percentage concentration of HydroSilver 1000 solution in film-forming solution, followed by calculating the AgNPs content in 1 mL of film-forming solution and 1 cm² of dried pullulan film.

2.7.1. Antibacterial Activity of Pullulan Film with AgNPs Determined Using Disk Diffusion Methods

A bacterial inoculum prepared according to Section 2.2 was uniformly spread by swab onto the surface of MHA plates and left to dry for 15 min. From each pullulan film sample, 6 mm diameter disks were cut with a circular knife and placed on previously prepared inoculated MHA plates. Pullulan disks without AgNPs served as negative controls, and a disk with an antibiotic (azlocillin) served as a positive control. The plates were incubated at 37 °C for 24 h. After incubation, the inhibition zones around the disk were measured in millimeters using a digital caliper (Standard Digital Caliper IP54, Shahe, Wenzhou, China). All tests were performed in nine replicates, and the results were expressed as mean values ± standard deviation (SD).

The sensitivity of the bacterial strains to the tested films was classified based on the diameter of the inhibition zone (DIZ) according to the following criteria: not sensitive (DIZ < 8 mm), sensitive (DIZ = 9–14 mm), very sensitive (DIZ = 15–19 mm), and extremely sensitive (DIZ > 20 mm) [56].

2.7.2. Scanning Electron Microscopy

The surface morphology and microstructure of pullulan films were visualized using a scanning electron microscope (type FEI, Quanta 200, JEOL, Tokyo, Japan). All samples were examined under low vacuum using an LFD detector and an accelerating voltage of 30 kV.

2.7.3. Thickness

The thickness of pullulan films was determined using a thickness gauge (BYKO-Test 4500, BYK-Gardner, Geretsried, Germany). To obtain representative data, six measurements were taken per film sample at randomly chosen locations, including each film’s central and external regions.

2.7.4. Opacity and Light Transmittance

The opacity and light transmittance of pullulan films were measured using a Metertech UV–VIS SP-8001 spectrophotometer (Metertech Inc., Taipei, Taiwan). Pullulan film strips of dimensions 1 × 4 cm were placed directly into the spectrophotometer’s test chamber, and the absorbance was measured at λ = 600 nm.

The opacity of the films was calculated using Equation (2) [57]:

$$\mathrm{Opacity} = {\displaystyle \frac{{Abs}_{600}}{Th}}$$

(2)

where Abs₆₀₀ is the absorbance value at 600 nm and Th is the film thickness (mm).

Additionally, using a UV-Vis spectrophotometer, the ultraviolet (UV) barrier and visible light transmission properties of the pullulan films were evaluated by measuring transmittance at wavelengths of 280 nm and 600 nm, respectively. Transmittance values were expressed as the percentage of light transmitted through the film at each wavelength. The measurement of each film sample was performed in triplicate.

2.7.5. Color Properties of Pullulan Film

The surface color characteristics of pullulan films were assessed using a CR-400 colorimeter (Minolta, Tokyo, Japan), based on the CIE Lab* color space. Three measurements were taken at four randomly selected locations across each film sample. The color parameters L* (lightness), a* (red-green color), and b* (yellow-blue color) were recorded. The total color difference (ΔE) between the modified and unmodified films was subsequently calculated using Equation (3).

$$\mathsf{\Delta }E=\sqrt{{\left(L_{PF}-L_{PFHS}\right)}^2+{\left(a_{PF}-a_{PFHS}\right)}^2+{\left(b_{PF}-b_{PFHS}\right)}^2}$$

(3)

where L_PF, a_PF, and b_PF are values for control pullulan films without AgNPs, and L_PFHS, a_PFHS and b_PFHS are values for pullulan films with AgNPs.

2.8. Statistical Analyses

Statistical analyses were performed using Statistica 13.3 PL (TIBCO Software Inc., Palo Alto, CA, USA). A one-way ANOVA followed by Tukey’s post hoc test was used to analyze the antimicrobial and physical properties of the films. A two-way ANOVA followed by Tukey’s post hoc test was applied to evaluate the time–kill curve results with concentration and time as the two sources of variation. For all analyses, p-values < 0.05 were considered statistically significant. The heatmap was generated using Microsoft Excel, and cluster analysis was performed using Statistica 13.3 PL.

3. Results

3.1. MIC and MBC of AgNPs Against Foodborne Bacteria

The antibacterial activity of AgNPs was observed against all tested foodborne bacteria, and the results are presented in

Table 2. Minimum Inhibitory Concentration (MIC) and Minimum Bactericidal Concentration (MBC) of AgNPs against tested strains of bacteria.

Bacteria Strains	AgNPs
	MIC	MBC	MBC/MIC *
	[µg/mL]
Bacteria Gram-negative
E. coli ATCC 25922	12.5	25	2
P. aeruginosa ATCC 27853	12.5	25	2
K. pneumoniae ATCC 13883	12.5	25	2
S. Enteritidis ATCC 13076	12.5	25	2
S. sonnei NIPH-NIH “s”	12.5	25	2
Bacteria Gram-positive
S. aureus ATCC 25923	12.5	25	2
E. faecalis ATCC 29212	12.5	25	2
L. innocua ATCC 33090	25	50	2
B. cereus ATCC 11778	6.25	12.5	2

* MBC/MIC ratio is ≤4, bactericidal effect; MBC/MIC ratio value is >4, bacteriostatic effect.

. In the case of Gram-negative bacteria, a narrow range of MIC and MBC values for AgNPs was observed, 12.5 μg/mL and 25 μg/mL, respectively. However, in the group of Gram-positive bacteria, the range of these values was wider. The highest sensitivity was demonstrated by the B. cereus strain (MIC 6.25 μg/mL, MBC 12.5 μg/mL), followed by S. aureus and E. faecalis (MIC 12.5 μg/mL, MBC 25 μg/mL). Higher resistance was found in L. innocua (MIC 25 μg/mL; MBC 50 μg/mL). The MBC/MIC values in all cases were 2, which, in relation to the classification [53] confirms that the AgNPs present in the HydroSilver 1000 solution have a bactericidal effect against all tested foodborne bacteria.

The percentage antibacterial activity of AgNPs in the tested concentration range against all test bacteria is illustrated in Figure 1. No bacteriostatic effect was observed for AgNPs at the lowest concentrations, ranging from 1.56 to 3.13 μg/mL. Partial inhibition (approximately 78%) occurred at 12.5 μg/mL. Complete inhibition of the growth of all strains (100%) occurred at a concentration of 25 μg/mL. This concentration can therefore be considered as the minimum effective concentration of AgNPs against all tested foodborne bacteria. Similar results were obtained by Singh et al. [58] in the control of bacterial biofilms. Silver nanoparticles at a concentration of 25 μg/mL showed a reduction in P. aeruginosa biofilm, and at a concentration of 50 μg/mL it was effective against E. coli biofilm formation. In a slightly lower MIC range, AgNPs inhibit the growth of multidrug-resistant enteroaggregative E. coli (MIC 8–16 μg/mL) and MBC (16–32 μg/mL) [59]. Silver nanoparticles with smaller sizes in the range of 5.0–6.0 nm stabilized with PVA showed stronger activity against clinical strains of S. aureus, E. coli, and P. aeruginosa (MIC 11.6 μg/mL), which is attributable to their larger surface available for interaction with bacteria [60]. Stabilization of silver nanoparticles using polymers prevents unwanted oxidation and agglomeration of silver nanoparticles in solution. Polyvinylpyrrolidone (PVP), polyvinyl alcohol (PVA), and polyethylene glycols (PEGs) are commonly used for this purpose. Other polymers have also been tested, e.g., polyacrylonitrile and polyacrylamide, poly(vinyl acetate-co-butyl acrylate-co-neodecanoate), and chitosan with thyme extract, with good results [60,61,62]. For example, PVA-modified silver nanoparticles are characterized by excellent dispersibility in water and physicochemical stability, which translates into better biological activity [63].

3.2. Effect of AgNPs on Bacterial Survival

The kinetics of bacterial inactivation over a 24 h incubation period were assessed using killing curves for the tested foodborne pathogens. These curves show the rate at which AgNPs exert their antibacterial effects. Silver nanoparticles were used in the study at concentrations corresponding to three values: ½ × MIC (sublethal), 1 × MIC (minimum inhibitory concentration), and 2 × MIC (double the MIC). The results are presented in Figure 2. Additionally, the logarithmic reduction in bacterial cell counts over time at each concentration was calculated and is presented in

Table 3. Log reduction in bacterial cell counts over time by AgNPs at different MIC levels.

Bacteria Strains	∆ log CFU/mL
	½ × MIC				1 × MIC				2 × MIC
	1 h	3 h	6 h	24 h	1 h	3 h	6 h	24 h	1 h	3 h	6 h	24 h
Bacteria Gram-negative
E. coli ATCC 25922	0.06	1.56	2.56	1.11	0.52	3.12	8.01	8.22	0.66	8.03	8.03	8.22
P. aeruginosa ATCC 27853	0.61	2.69	4.56	5.78	0.82	5.99	7.74	8.98	1.01	5.99	7.74	8.98
K. pneumoniae ATCC 13883	0.34	2.24	4.46	7.88	0.26	3.33	6.20	7.88	0.76	5.99	7.71	7.88
S. Enteritidis ATCC 13076	0.16	1.55	4.21	3.86	0.45	2.77	8.03	9.08	0.66	5.37	8.03	9.08
S. sonnei NIPH–NIH “s”	0.20	2.20	7.91	9.99	0.37	2.85	7.91	9.99	0.43	5.53	7.91	9.99
Bacteria Gram-positive
S. aureus ATCC 25923	0.58	1.35	2.83	2.60	0.89	2.70	6.84	7.93	1.33	5.38	6.84	7.93
E. faecalis ATCC 29212	0.79	1.71	2.98	1.09	1.42	3.53	7.73	7.99	1.61	5.53	7.73	7.99
L. innocua ATCC 33090	0.48	2.90	3.14	4.75	0.98	3.79	6.99	8.14	1.65	6.02	6.99	8.14
B. cereus ATCC 11778	0.03	1.37	1.29	1.29	0.75	2.94	7.39	8.36	1.83	5.37	7.36	8.36

. The number of cells of all test strains increased over 24 h only in the control samples. The presence of AgNPs in the growth medium, regardless of the concentration, statistically significantly (p < 0.05) limited the growth of strains. A reduction in cell count by 3 log CFU/mL or more differentiates the activity of compounds into bactericidal and bacteriostatic [52]. The use of AgNPs at a sublethal concentration (½ × MIC) revealed bacteriostatic activity against E. coli, S. aureus, E. faecalis, and B. cereus after 6 h of incubation and bactericidal activity against other bacterial strains. After 3 h of incubation, AgNPs at a concentration of 1 × MIC had a bactericidal effect against E. coli (3.12 log CFU/mL), P. aeruginosa (5.99 log CFU/mL), K. pneumoniae (3.33 log CFU/mL), E. faecalis (3.53 log CFU/mL), and L. innocua (3.79 log CFU/mL), and a bacteriostatic effect against the remaining strains. After 6 h of incubation, the cell count of all test strains was below the detection limit (<10 CFU/mL). Silver nanoparticles at a concentration of 2 × MIC after 3 h of incubation reduced the cell count of all strains by 3 or more logarithmic cycles. In general, it can be seen that the higher the concentration of AgNPs is, the shorter is the killing time of the test strain cells. Complete cell destruction of the tested strains occurred after 6 h of incubation at 1 × MIC and after 3 h of incubation at 2 × MIC.

3.3. Leakage of DNA and Intracellular Proteins from Cells Assessed by Spectrophotometric Measurements

Confirmation of cell destruction was carried out by assessing leakage of intracellular content during incubation of strains with AgNPs. Nucleic acid and protein leakage in the supernatant of tested bacteria were measured at 260 nm and 280 nm, respectively (

Table 4. Leakage of nucleic acid and protein from bacterial cells after treatment with AgNPs at different MIC levels.

Bacteria Strains	Concentration of AgNPs	Leakage of Bacterial Cellular Material at 260 nm and 280 nm
		260 nm	280 nm
		6 h	6 h
E. coli ATCC 25922	control	0.137 ± 0.025 a **	0.107 ± 0.009 a
	½ × MIC	0.182 ± 0.008 b	0.124 ± 0.006 b
	1 × MIC	0.210 ± 0.022 c	0.157 ± 0.020 c
	2 × MIC	0.229 ± 0.012 d	0.179 ± 0.012 d
S. Enteritidis ATCC 13076	control	0.181 ± 0.037 a	0.122 ± 0.039 a
	½ × MIC	0.173 ± 0.004 a	0.120 ± 0.005 a
	1 × MIC	0.233 ± 0.006 b	0.165 ± 0.005 b
	2 × MIC	0.326 ± 0.003 c	0.260 ± 0.003 c
S. sonnei NIPH–NIH “s”	control	0.196 ± 0.048 a	0.155 ± 0.043 a
	½ × MIC	0.354 ± 0.026 b	0.236 ± 0.023 b
	1 × MIC	0.339 ± 0.015 b	0.238 ± 0.014 b
	2 × MIC	0.416 ± 0.014 c	0.330 ± 0.012 c
S. aureus ATCC 25923	control	0.348 ± 0.011 a	0.203 ± 0.010 a
	½ × MIC	0.332 ± 0.021 a	0.211 ± 0.020 a
	1 × MIC	0.395 ± 0.002 b	0.258 ± 0.002 b
	2 × MIC	0.513 ± 0.001 c	0.369 ± 0.001 c
E. faecalis ATCC 29212	control	0.165 ± 0.004 a	0.131 ± 0.002 a
	½ × MIC	0.320 ± 0.002 b	0.229 ± 0.003 b
	1 × MIC	0.347 ± 0.007 c	0.238 ± 0.006 b
	2 × MIC	0.469 ± 0.005 d	0.391 ± 0.005 c
L. innocua ATCC 33090	control	0.071 ± 0.004 a	0.055 ± 0.004 a
	½ × MIC	0.149 ± 0.008 b	0.106 ± 0.006 b
	1 × MIC	0.236 ± 0.013 c	0.157 ± 0.007 c
	2 × MIC	0.454 ± 0.012 d	0.274 ± 0.007 d

** a–d—different letters in the column indicate statistical differences at p < 0.05 determined by a one-way ANOVA followed by Tukey’s test.

). An increase in absorbance values was observed in all samples, indicating leakage of nucleic acids and proteins from the cells. Increasing the AgNPs concentration to 2 × MIC contributed to greater leakage of components from the cells, indicating greater degradation of the cell wall and cytoplasmic membrane in the cells. Our results are consistent with Lu et al. [64], who found that PVA-stabilized AgNPs copolymer disrupted the integrity of E. coli and S. aureus bacterial cell membranes, which caused leakage of cell contents, leading to disruption of the homeostasis of the intracellular environment and preventing the normal functioning of cell vital activity.

3.4. Morphological Assessment of Bacterial Cells After AgNPs Exposure

Morphological changes in the bacteria cell were examined using SEM and TEM, and the micrographs are shown in Figure 3. The micrographs illustrate the possible effects of AgNPs on the cell wall and cytoplasmic membrane. The effects of cell damage are shown in Figure 3. Formation of pores in the cell wall was observed, causing cytosol leakage outside the cell. Furthermore, AgNPs were observed to significantly deform cells, altering their shape, causing them to become folded, and shrinkage was observed. The above-mentioned features led to loss of cell integrity.

Both spectrophotometric determination and microscopic observations confirmed that AgNPs have a destructive effect on bacterial cells. Sondi and Salopek-Sondi [65] observed damage to the E. coli cell wall and the accumulation of AgNPs in the bacterial membrane, which causes a significant increase in permeability. Cell materials leak through pores in the cell membrane. The external cell structures act as a protective barrier, protecting the cell from environmental factors. These structures are the first to come into contact with AgNPs and are the first to undergo perforation and penetration [66,67]. In the cytoplasmic membrane, silver ions can bind to phosphorus and sulfur compounds present in cytoplasmic membrane proteins, disrupting their metabolic functions [68]. If damaged, AgNPs induce other effects related to changes in cell shape, resulting in multifaceted deformations. Silver accelerates the production of reactive oxygen species (ROS) inside the cell, causing oxidative stress, which is lethal to bacteria [69]. Furthermore, AgNPs can bind to DNA, which inhibits cell proliferation and leads to cell death [68,70,71].

3.5. Antibacterial Activity of Pullulan Films with AgNPs

Controlled release of AgNPs from the polymer film is crucial because it determines its effectiveness over time and determines the long-term antibacterial properties of the film [67]. The antibacterial activity of pullulan films containing AgNPs was assessed by the disk diffusion method against foodborne test bacteria. The average diameters of the growth inhibition zones of the test strains treated with pullulan films containing AgNPs are shown in

Table 5. Inhibition zone diameters of bacterial pathogens exposed to pullulan film with silver nanoparticles.

Bacteria Strains	Pullulan Films
	PF	PFHS1	PFHS3	PFHS5	PFHS10	PFHS15	PFHS20	AZL75
	Concentration of AgNPs in film [µg/cm2]
	0	2	5	8	16	24	32	x
Bacteria Gram-negative
Growth inhibition zones [mm ± SD]
E. coli ATCC 25922	0.0 a	8.4 ± 0.2 bB **	11.6 ± 0.3 cC	12.5 ± 0.4 dCD	13.1 ± 0.3 eA	14.5 ± 0.3 fA	14.4 ± 0.3 fAB	16.5 ± 0.4 gB
K. pneumoniae ATCC 13883	0.0 a	6.8 ± 0.6 bA	7.9 ± 0.6 cA	9.9 ± 0.6 dA	14.6 ± 0.5 fD	15.8 ± 0.4 gBC	15.9 ± 0.4 gBC	13.1 ± 0.8 eA
P. aeruginosa ATCC 27853	0.0 a	12.6 ± 0.9 bE	13.0 ± 0.4 bE	13.7 ± 0.4 cF	15.2 ± 0.3 dE	15.7 ± 0.4 dBC	15.8 ± 0.2 dB	26.9 ± 0.6 eF
S. Enteritidis ATCC 13076	0.0 a	10.6 ± 0.7 bCD	10.9 ± 0.3 bB	12.9 ± 0.4 cD	16.2 ± 0.3 dF	15.6 ± 0.4 dB	15.4 ± 0.5 dB	23.5 ± 1.2 eE
S. sonnei NIPH–NIH “s”	0.0 a	11.4 ± 1.0 bD	12.5 ± 0.5 cDE	13.9 ± 0.3 dF	14.1 ± 0.3 dC	16.0 ± 0.5 eC	15.8 ± 0.8 eB	18.7 ± 1.0 fC
Bacteria Gram-positive
B. cereus ATCC 11778	0.0 a	9.6 ± 0.6 bC	10.8 ± 0.7 cB	11.7 ± 0.4 dB	13.6 ± 0.4 eBC	17.0 ± 0.2 fE	17.3 ± 0.2 gD	13.3 ± 0.8 eA
E. faecalis ATCC 29212	0.0 a	10.2 ± 0.4 bC	10.7 ± 0.3 bB	12.0 ± 0.5 cBC	13.5 ± 0.4 dAB	17.5 ± 0.3 eF	17.8 ± 0.1 eDE	29.3 ± 0.6 fG
S. aureus ATCC 25923	0.0 a	10.4 ± 0.5 bCD	12.0 ± 0.5 cCD	14.0 ± 0.6 dF	15.2 ± 1.1 eE	17.0 ± 0.6 fE	17.0 ± 0.4 fCD	20.0 ± 0.8 gD
L. innocua ATCC 33090	0.0 a	9.9 ± 0.7 bC	10.7 ± 1.1 bB	12.2 ± 0.8 cBC	16.3 ± 0.5 dF	19.2 ± 0,4 eG	18.9 ± 0.2 eE	33.4 ± 0.3 fH

** a–g—different letters in the row indicate statistical differences at p < 0.05; A–H—different letters in the column indicate statistical differences at p < 0.05. PF—pullulan film, PFHS—pullulan film with HydroSilver1000 in a concentration range from 1 to 20%.

. In this study, the antibiotic azlocillin was used as a positive control, and a pullulan film without AgNPs as a negative control to assess the antibacterial properties of the films. Azlocillin is an antibiotic with a broad spectrum of activity against both Gram-positive and Gram-negative bacteria.

The incorporation of AgNPs, even in the lower amounts (PFHS1 and PFHS3), resulted in an antibacterial effect, except for K. pneumoniae (zones < 8 mm). Higher antibacterial potency expressed by the size of the inhibition zones of the strains’ growth [56] was correlated with a higher content of AgNPs in the pullulan film (Table 5). The heatmap revealed the division of the films into two groups in terms of antibacterial potency (Figure 4): PFHS1, PFHS3, and PFHS5 films were weaker, while PFHS10, PFHS15, and PFHS20 were stronger. Additionally, bacterial strains were divided into two groups according to their sensitivity to the films: E. coli, K. pneumoniae, B. cereus, E. faecalis, and L. innocua had weaker and more diverse sensitivity, while P. aeruginosa, S. sonnei, S. Enteritidis, and S. aureus were highly sensitive. The inhibitory effect of the films on azlocillin was varied. The PFHS20 film had an inhibitory potency against K. pneumoniae and B. cereus even greater than that of azlocillin (121% and 130%, respectively). There was no statistically significant difference (p > 0.05) between the growth inhibition zones of strains in PFHS20 and PFHS15 films. This means that the effectiveness of these films was the same. Therefore, it is more economical to use the pullulan film containing AgNPs at a lower concentration of 24 µg/cm² (PFHS15).

The PFHS15 and PFHS20 films showed good efficacy against foodborne bacteria. The antibacterial activity is related to the release of silver ions from the pullulan film into the environment and/or contact of microorganisms with metallic silver nanoparticles [43,61]. Polymer matrices improve the stability of AgNPs and antibacterial properties [61,72]. Results presented by Khalaf et al. [45] and Morsy et al. [46] also confirmed that pullulan-based films impregnated with silver nanoparticles exhibit good efficacy against S. aureus and L. monocytogenes. The composite coating based on bacterial cellulose with montmorillonite and nanosilver shows good antibacterial activity against S. enterica, E. coli, S. aureus, and B. subtilis and is characterized by high mechanical strength, making it a promising material for food preservation applications [73]. A chitosan-gum tragacanth biomaterial containing nanosilver effectively inhibited the growth of P. aeruginosa and S. aureus [74]. Wang et al. [75] composed a film of chitosan, gelatin, and nanosilver with improved conductivity and antibacterial activity against E. coli and S. aureus, suggesting potential use in biomedicine. Pinto et al. [43] demonstrated the antifungal activity of a pullulan film with AgNPs. Increasing the AgNPs content in the pullulan film resulted in a greater reduction in fungal concentration, spore destruction, and growth inhibition of A. niger. The composite pullulan film with AgNPs has significant potential for applications in food packaging [72].

3.6. Visual Appearance and Microscopic Characterization of Pullulan Films with AgNPs

The visual appearance, along with the microscopic characterization of all tested films, is shown in Figure 5. The films were smooth to the touch, flexible, and shiny. Incorporating AgNPs into the transparent and colorless control film changed the color, with the intensity varying gradually from light yellow to dark brown, depending on the AgNPs content in the film. Very similar films were obtained by Wypij et al. [44].

In both the cross-section and on the surface, the control pullulan film (PFC) was characterized by a uniform structure and a smooth surface (Figure 5). Our observations are consistent with the pullulan film microstructure described by Mulla et al. [76]. Incorporating a 1–5% solution of silver nanoparticles (PFHS1, PFHS3, PFHS5) into the pullulan film did not change the film’s microstructure. The films exhibited a compact and uniform structure with visible individual bulges, as shown in Figure 5. This indicates good mixing of AgNPs with the polysaccharide matrix. After increasing the AgNPs addition, the films remained uniform in the cross-section in the pullulan films PFHS10, PFHS15, and PFHS20, but numerous bulges were observed on the surface (Figure 5). This proves very good dispersion of AgNPs with a diameter of 50–60 nm in the polysaccharide matrix. Similar observations of the structure of pullulan films with the addition of silver nanoparticles with a diameter of 100 nm and zinc with an average size of 110 nm at a concentration of 0.02%, v/v, were observed by Morsy et al. [46]. Islam and Yeum [77] confirmed the good embedding of Ag nanoparticles in a pullulan/PVA blend matrix from which electrospun nanofibers were prepared and found that silver nanoparticles were evenly distributed in them. Vimala et al. [78] stated that the polymers provide excellent stability of AgNPs, slow their release into the environment, and ensure prolonged antibacterial activity. These results confirm that pullulan films are a suitable material for delivering antimicrobial substances, including metal nanoparticles, to food surfaces. Good dispersion of AgNPs and deposition of small AgNPs on the composite film surface with a rough film structure, with almost uniform nanoparticle distribution, were observed in moxifloxacin-loaded chitosan-pullulan-silver-nanocomposite films [79]. Overall all tested pullulan film with AgNPs exhibited homogenous cross-section microstructure, which according to available literature it can be believed that pullulan helps in stabilization and prevent the agglomeration of AgNPs thus, maintain their good dispersion and promotes the uniform distribution of silver nanoparticles throughout the pullulan film [42].

3.7. Physical Properties of Pullulan Films with AgNPs

The physical properties of pullulan films containing AgNPs are presented in

Table 6. Physical properties (thickness, opacity, transmittance and color) of pullulan film with silver nanoparticles.

Film Sample	Concentration of AgNPs in Film [µg/cm2]	Thickness [mm]	Opacity [a.u./mm]	Transmittance (%)		Color
				280 nm	600 nm	L*	a*	b*	ΔE
PF	0	0.071 ± 0.005 a **	0.53 ± 0.04 a	79.05 ± 1.38 a	91.76 ± 0.40 a	91.25 ± 0.15 a	2.08 ± 0.04 a	−5.21 ± 0.03 a	-
PFHS1	2	0.080 ± 0.004 ab	0.59 ± 0.04 a	52.10 ± 0.79 b	89.63 ± 1.16 a	84.22 ± 0.20 b	4.71 ± 0.30 b	2.86 ± 0.12 g	7.9
PFHS3	5	0.095 ± 0.007 b	0.61 ± 0.07 a	51.25 ± 0.72 b	87.57 ± 0.76 a	79.14 ± 0.21 c	4.38 ± 0.13 c	25.78 ± 0.62 f	24.0
PFHS5	8	0.098 ± 0.007 b	1.64 ± 0.06 b	51.17 ± 1.74 b	69.16 ± 1.20 b	71.11 ± 0.24 d	10.16 ± 0.34 d	20.89 ± 0.64 e	26.7
PFHS10	16	0.115 ± 0.011 c	2.37 ± 0.31 c	33.59 ± 3.20 c	53.73 ± 3.86 c	63.67 ± 1.13 e	16.47 ± 0.70 f	18.59 ± 0.26 d	33.9
PFHS15	24	0.125 ± 0.002 c	2.54 ± 0.23 c	20.77 ± 1.50 d	48.30 ± 3.19 c	41.82 ± 2.54 f	20.20 ± 0.61 g	11.05 ± 0.56 c	53.0
PFHS20	32	0.144 ± 0.005 d	3.83 ± 0.26 d	15.53 ± 1.32 e	28.24 ± 2.12 d	41.45 ± 1.57 f	13.50 ± 0.96 e	0.01 ± 0.0 b	51.4

** a–g—different letters in the column indicate statistical differences at p < 0.05 determined by a one-way ANOVA followed by Tukey’s test. PF—pullulan film, PFHS—pullulan film with HydroSilver1000 in a concentration range from 1 to 20%.

. Thickness is one of the most important parameters characterizing films. The thickness of transparent polymeric food packaging materials is typically in the range of 0.050–0.1 mm [80]. The thickness of the films increased statistically significantly (p < 0.05) from 0.080 (PFHS1) to 0.144 mm (PFHS20). The PFHS20 film was twice as thick as the PF film. Our results are consistent with those reported by Wypij et al. [44], but differ from the observations of Khalaf et al. [45], who found no association of the thickness of pullulan films with AgNPs. The authors attributed their results to the small—even negligible—molecular mass of AgNPs in relation to the mass of the polysaccharide used.

Optical transparency is an important packaging feature for consumers because it allows them to visually assess the appearance and freshness of food and make a purchasing decision [81]. On the other hand, adverse effects of light energy on food have been observed due to the degradation of nutrients and, consequently, changes in food quality, shelf life, and safety [82]. In the study of film transparency, two main parameters are considered: opacity and transmittance [80]. Analysis of variance showed a statistically significant (p < 0.05) effect of AgNPs addition on both film parameters. The opacity values of the films varied from 0.53 (PF) to 3.83 (PFHS20), and since they are less than 5, they are still transparent [80]. In turn, transmittance in the visible range did not differ significantly (p > 0.05) only in PFHS1 and PFHS3 compared to PF, and these films can be considered highly transparent (91.76–87.57% transmittance). In PFHS5–PFHS20 films, transmittance values decreased from 69.16% to 41.82%, placing these films in the category of translucent materials [80]. Similarly, transmittance in the UV range (280 nm) was significantly reduced by the AgNPs films (from 52.1 to 15.53%) compared to the PF film (79.05% transmittance), which is beneficial from the point of view of food protection against UV radiation, which damages food by causing photodegradation of nutrients [83]. A possible explanation for this phenomenon is the absorption of both UV radiation and visible light by the silver nanoparticles. UV absorption reduces food damage by delaying the formation of free radicals [84].

The color parameters of the films are expressed using the CIE L*a*b* scale. Color differences compared to the control pullulan film were also calculated. Analysis of variance showed a statistically significant (p < 0.05) effect of AgNPs on the color of the films. The increase in AgNPs content in the film caused a statistically significant (p < 0.05) darkening of the color from the light control film (L* 91.25 for PF) to a much darker one (L* 63.47 for PFHS20). The color of the films with AgNPs had a greater proportion of red (positive a* values) and yellow (positive b* values) compared to the color of the film without AgNPs. The absolute color difference (ΔE) increased with increasing AgNPs content in the film, indicating large, visible color changes for the consumer.

4. Conclusions

This study presented the results of antibacterial activity of commercially available silver nanoparticles against select 9 strains of foodborne bacteria as well as the properties of pullulan films infused with AgNPs, including their antibacterial activity and selected physical properties. The results of the first part of the study showed that silver nanoparticles exhibited strong antibacterial activity against all tested bacterial strains. Silver nanoparticles exhibited relatively uniform antibacterial activity against all Gram-negative bacteria and two Gram-positive strains (S. aureus and E. faecalis), with MIC and MBC values of 12.5 µg/mL and 25 µg/mL, respectively. In case of other tested Gram-positive bacteria, L. innocua expressed the greatest sensitivity to AgNPs, while B. cereus was the most resistant. Treatment of bacterial cells with AgNPs results in the leakage of nucleic acids and proteins from the cells. In this context, our results indicate that one of the mechanisms of antibacterial activity is connected with membrane damage, which was also confirmed by examining morphological changes in bacterial cells exposed to AgNPs using SEM and TEM. Silver nanoparticles exhibited severe structural damage on the bacteria, causing damage in the structural integrity of the cell wall and membrane, as well as broken, destroyed cells and leakage of cell contents. In the second part, our research has shown that pullulan films containing silver nanoparticles combine the beneficial properties of a biopolymer with the antibacterial activity of silver. All pullulan films with AgNPs possess antibacterial activity. Moreover, an increase in the AgNPs concentration in pullulan film resulted in a stronger inhibitory effect against the tested bacterial strains. Furthermore, among all the tested bacterial strains, L. innocua was the most sensitive. Pullulan films with the addition of AgNPs contributed to a gradual increase in thickness and opacity. In terms of optical properties, increasing concentrations of AgNPs resulted in decreased UV and visible light transmittance, along with a noticeable color change from colorless (control) to dark brown.

The promising results obtained in this study open a perspective for further research on pullulan films containing commercially available silver nanoparticles, particularly in exploring their mechanical and barrier properties. To confirm the formation and compatibility of pullulan-based films with silver nanoparticles, comprehensive analyses should be conducted, including the determination of chemical composition (FTIR), crystallinity and structural integrity (XRD), and thermal characteristics (DSC). Moreover, to evaluate their potential use as food packaging, future studies should include in vitro tests to assess the antibacterial activity of these materials directly on food products.