One-step synthesis of cellulose/silver nanobiocomposites using a solution plasma process and characterization of their broad spectrum antimicrobial efficacy

Abstract

Solution plasma process (SPP) is a one-step synthesis technique which expeditiously produces ultra-pure, stable, and uniform nanoparticles in polymer solutions with plasma discharge. Silver nanoparticles (AgNPs) were synthesized in a cellulose matrix as biocomposites by discharging plasma for 180 s at 800 V with a frequency of 30 kHz using a pulsed unipolar power supply into solutions containing cellulose (1–3%) and AgNO₃ (1–5 mM). 3D scaffolds of the resulting cellulose/AgNP biocomposites were prepared by lyophilization and cross-linked with UV irradiation. UV-Vis spectroscopy showed a characteristic absorbance maximum in the range of 350–440 nm for the AgNP biocomposites with increase in the intensity of the peaks as the concentration of AgNO₃ increased. The peaks exhibited a red shift transition due to the AgNP formation. The nanobiocomposites were pure when examined by FTIR spectroscopy. The 3D scaffolds had a micro-porous structure with pores of (68–74) ± 2 μm in diameter when observed using a FE-SEM instrument equipped with an EDS function. TEM analysis showed that spherical AgNPs in the size range of 5–30 nm were well distributed in the biocomposites of C3Ag3 and C3Ag5. The nanobiocomposites had a broad spectrum of antimicrobial activity against various pathogens with a minimal inhibition concentration of 5.1–20.4 μg ml⁻¹ for bacteria and 81.6–255.0 μg ml⁻¹ for fungi. They killed gram negative bacteria most effectively, but did not affect fungal growth very well, implying their potential as topical antimicrobial agents for the topical treatment of wounds. SPP seems to be the most effective and safest method to synthesize various biocompatible polymer–metal nanoparticle biocomposites.

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1. Introduction

The perpetual copious research, which continues to provide new illuminating scientific data, validates the desideratum for a continuation of the research, development, and analysis of the synthesis of nanomaterials by physical, chemical and biological routes for their application in nanotechnology in general and for nanomedicine in particular. The generation of nanoparticles, typically by either chemical or physical methods, often requires the addition of toxic chemicals and expensive equipment, and mostly involves multiple steps.¹ During the biological route, two consecutive steps of synthesis and purification of nanoparticles resulted in the production of nanoparticles with biomolecules on the surface, which could then trigger harmful signals during utilization.² Insufficient synthesis methodologies to prevent the incorporation of additives, detergents, or chemicals that might not completely be removed can also contribute to the toxicity of the nanoparticles prepared.³

Solution plasma process (SPP) is an ideal method for the synthesis of nanoparticles without adding toxic chemical reagents and with no need of expensive instruments. However, SPP has not been employed widely for that purpose, despite the extensive scientific evidence and its potentiality in applications of materials science and medicine.^4,5 SPP involves a sequence of physical and chemical reactions, in which water molecules split into free radicals (H˙, OH˙, electrons, UV) and solutes (precursors) into ions to form nanoparticles during the discharge of plasma in the solution.⁶ Faster chemical reactions at lower temperature with greater variability are possible by plasma generated in solutions of various solutes and solvents, since the density of molecules in the liquid phase is much higher than that of the gas phase.⁷ SPP can be widely used for the degradation of polymers, surface coating, and the fabrication of materials at the nanoscale level with various dimensions and structures.^8–10 Especially, nanoparticles synthesized can be simultaneously fabricated into macromolecular polymers by SPP such that the particles are evenly scattered in the matrix without agglomeration by forming a 3D scaffold. Thereby, plasma plays an active role in the synthesis and stabilization of nanoparticles derived from the solutes in the solution without adding hazardous chemicals.^11–13

The meticulous evidence-based research, analyses, and reviews on the plasma mediated synthesis of nanomaterials, identifies the growing prospect of validating its applicability in material science, biomedicine, and particularly in nanomedicine.⁴ The generation of nanomaterials (metals, oxides, composites, polymers) using plasma in liquids by adding hazardous chemicals for reduction and stabilization was reviewed extensively and is summarized with regard to size and applications in Table 1.^14–26 There is strong evidence that interactions between the nanoparticles and polymers are responsible for the alteration in electrical and thermo-mechanical properties, increasing specific surface area with decreasing particle size, and that the amount of interfacial polymer layer strongly depends on the size and concentration.²⁶

Table 1 Summary of the synthesis of nanomaterials (metals, oxides, composites, polymers) and their size and application using plasma in liquids

Nanomaterials synthesized	Size of the materials	Applications	References
Metal nanoparticles
C60	7 nm	Conductivity	14
Au	15 nm	Catalytic property	15
Cu	33.7 ± 5.8 nm	Material engineering	16
Ni	10–200 nm	Semi-heterogeneous catalysis	17
Ag	61.8 ± 21.8 nm	Sensors	18
Metal–metal nanocomposites
Ag/Pt	5 nm	Heterogeneous catalytic activity	19
Ag/Si	20 nm	Catalytic property	20
Pt/C	38.14 nm	Membrane fuel cells	21
Metal oxide nanoparticles
WO3	5 nm	Solar energy	22
ZrO2	5 nm	Solar energy	23
Fe3O4	19 nm	Magnetic and optics	24
Polymer–polymer nanocomposites
Cellulose/aniline	100–200 nm	Electrical conductivity	25
Polymer–metal nanocomposites
Gelatin/Ag	10–15 nm	Antimicrobial	4, 26
Cellulose/Ag	5–20 nm	Broad spectrum of antimicrobial property	This study

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Among diverse nanomaterials, significant focus has been made on silver (Ag) due to its unique properties, such as conductivity, stability, catalytic, and antimicrobial properties.² Silver and silver based composites have been reported to exhibit antimicrobial activity against a wide range of microorganisms such as bacteria, fungi, protozoa, and recently viruses.²⁶ It is of paramount significance to develop, identify, validate, and employ an enhanced synthesis of polymer–metal composites that are effective against pathogens, since various multi-drug resistant pathogens which are not controlled by commercial antibiotics, have emerged and are a serious threat to human society.²⁷ Silver nanoparticles (AgNPs) well distributed in non-toxic and stable biological polymers such as cellulose, one of the most abundant biomasses on earth, would maximize its potential as an antimicrobial agent against pathogens without causing their resistance to it.

Therefore, in this study, the synthesis of cellulose/AgNP biocomposites was attempted using a one-step SPP without the addition of any hazardous chemicals as reducing and/or stabilizing agents. Physical and chemical parameters were optimized for the synthesis of the biocomposites and the resulting biocomposites were characterized using TEM, FE-SEM, EDS, FTIR, and UV-Vis spectroscopy. Additionally, the cellulose/AgNP biocomposites were assessed for their antimicrobial activity against several human pathogens such as bacteria (Escherichia coli, Pseudomonas aeruginosa, Vibrio parahaemolyticus, Staphylococcus aureus and Bacillus cereus), yeast (Candida albicans), and mold (Aspergillus parasiticus). To our knowledge, this may be the first work on the generation of cellulose/AgNP biocomposites by SPP for a medicinal application, which might be applied to topical wound healing materials, thereby significantly reducing microbial infection and promoting the recovery of dermal disease.

2. Materials and methods

2.1. Solution plasma process set up

Briefly, 150 ml of solution containing 1, 3, or 5 mM silver nitrate (AgNO₃, Junsei Co., Japan) and 1, 2, or 3% hydroxypropyl methylcellulose (HPMC, AN4, Samsung Fine Chemicals, Ltd., Korea) was mixed in a 200 ml Pyrex beaker specially designed for the process in ambient conditions as described previously.²⁸ Plasma was generated using a pulsed field unipolar power supply (IAP-1010, EN Technology, Korea) in the solution with a voltage, frequency, pulse width, electrode distance, and discharge time of 800 V, 30 kHz, 2 μs, 1 mm, and 3 min, respectively. A magnetic stirrer was used for the constant mixing and complete dispersion of the solutes in the solution. In order to understand the effects of plasma discharged in the solution, pH, temperature, and color change were observed and recorded periodically at every 1 min throughout the experimental process. The products of the process were notated as CxAgy, where x and y represent the concentration of cellulose and silver nitrate, respectively.

2.2. Fabrication of the cellulose/AgNP

Cellulose/AgNP biocomposites were successfully fabricated using the freeze drying (lyophilization) method. Briefly, 5 ml of the synthesized nanoparticle biocomposites was transferred into a sterile Petri dish and frozen at −80 °C overnight in a deep freezer (Forma Scientific, USA). The frozen samples were lyophilized at −40 °C with the pressure of 6.38 × 10⁻⁴ MPa using a freeze dryer (FDV-7024, OPERON, Korea). The dried samples were cross linked by UV irradiation (254 nm) for 30 min to reduce the water solubility.

2.3. Characterization of the cellulose/AgNP biocomposites

The generation of AgNPs in the cellulose matrix using solution plasma was initially confirmed by an UV-Visible spectrophotometer (UV-3600, UV-Vis NIR spectrophotometer, Shimadzu, Japan) in the range of 200–1200 nm. The viscosity of the synthesized biocomposites was also assessed using a viscometer (Vibro SV-100, A&D Co., Japan) at room temperature. An apparent distribution of AgNPs was observed using transmission electron microscopy (TEM; JEOL-JSM, JOEL Ltd., Japan). The structure, texture, and porosity of the 3D scaffold were examined by a field emission scanning electron microscope (JEOL-JSM-7001F, JEOL Ltd., Japan) equipped with an energy dispersive spectroscopy facility in order to assess the elemental percentage within the biocomposite. Both the purity and functional association of the biocomposite were studied using Fourier transform infrared spectroscopy (FTIR; Vertex 8V, Bruker, Germany) in the range of 400 cm⁻¹ to 4000 cm⁻¹.

2.4. Cultivation of microorganisms

Seven microbial pathogens including E. coli, P. aeruginosa, V. parahaemolyticus, S. aureus, and B. cereus (bacteria), A. parasiticus (fungus), and C. albicans (yeast) were used to test the antimicrobial activity of the cellulose/AgNP biocomposites. Bacteria were cultured in Luria-Bertani medium (LB; yeast extract 0.5%, NaCl 0.5%, tryptone 1%) at 37 °C; A. parasiticus in potato dextrose medium (PD; potato dextrose 2.4%) at 25 °C; and C. albicans in yeast peptone medium (YPD; yeast extract 1%, peptone 2%, dextrose 2%) at 30 °C. Agar (1.5%) was added to each medium when it was used as solid plates. The amount of cells was determined using the McFarland standard.

2.5. Evaluation of the antimicrobial properties of the cellulose/AgNP biocomposites

2.5.1. Agar diffusion assay. The antimicrobial properties of the biocomposite scaffolds were assessed by inhibiting microbial reproduction on agar plates, which resulted in the formation of a clear zone around the discs according to the Kirby–Bauer agar diffusion method.²⁹ About 10⁴ colony forming units (CFUs) of freshly cultured microbial cells or fungal spores were spread on Műller-Hinton agar plates (beef extract 0.2%, acid digest of casein 1.75%, starch 0.15%, agar 1.7%; Difco Co., USA) and discs of the C3/Ag1, Ag3, or Ag5 biocomposite scaffolds (6 mm in diameter) were placed on them aseptically. The resulting zone of growth inhibition was measured to assess the antimicrobial effect in 24 h (bacteria), 48 h (yeast), or 72 h (fungus) of incubation at an appropriate temperature as described above.

2.5.2. Minimal inhibitory concentration (MIC) of the cellulose/AgNPs. The MIC of the biocomposites for each microorganism was determined by adding 0.0–326 μg ml⁻¹ of the AgNPs using the biocomposite of C3Ag5 to each microbial liquid culture (1–2 × 10⁵ CFU; 5 ml). The tubes were incubated at 37 °C for 18–24 h with gentle shaking. The MIC was determined as the lowest concentration of the nanobiocomposites that produced no visible microbial growth (no turbidity) in comparison with that of the control tube to which no nanobiocomposites were added.

2.5.3. Reduction of CFU assay. The bactericidal effect of the cellulose/AgNP biocomposites was assessed quantitatively by monitoring the reduction of CFU upon treatment with the nanobiocomposites during incubation. The amount of the cellulose/AgNP biocomposite to add was determined based on the MIC results. A precise amount of each cellulose/AgNP biocomposite was added to LB broth inoculated with either E. coli or S. aureus (∼10⁶ CFU ml⁻¹) and incubated at 37 °C for 1–16 h. Cellulose free of AgNP was used as a control in all of the experimental sections. During incubation, an aliquot of culture was taken periodically and inoculated on LB agar plates after appropriate dilution to determine the CFU remaining by incubating at 37 °C overnight.

2.5.4. Inhibition assay of fungal growth. Effect of the nanobiocomposites on mycelia growth of A. parasiticus was examined by placing a block of fungal hyphae (6 mm) on the centre of a PDA plate that had been smeared with 200 μl of the cellulose/AgNP biocomposites. A PDA plate with no nanobiocomposites smeared was used as the control. All of the plates were incubated for 72–120 h at 25 °C before observation on the morphology, hyphal growth, and sporulation pattern via pigmentation of sporangium.

3. Results and discussion

3.1. Synthesis of the cellulose/AgNP biocomposites

Analysis of the credible scientific data produced in this current study proved that pure, stable, and hazardous chemical free cellulose/AgNP biocomposites were successfully synthesized by the one-pot synthesis method of SPP. A total of 9 different biocomposites were prepared with various combinations of cellulose (1, 2, 3%) and AgNO₃ (1, 3, 5 mM) concentrations. The initial pH, temperature, and color of the solution were noted as 7.0 ± 0.2, 28 °C, and a pale white color, respectively, before the plasma discharge. Once the plasma was discharged, the solutions turned brown. This could be due to the reduction of silver from ionic (Ag⁺) into metallic (Ag⁰) nanoparticles in the cellulose matrix by the active species of free radicals generated by the solution plasma. The intensity of the color developed has been reported to be dependent on the concentration of AgNO₃ and the polymer in use as well as the discharge time.¹⁶ Previously, a similar color change was observed during the synthesis of AgNPs using gelatin as the polymer matrix.²⁶ During the process, the pH of the solution was slightly decreased to an acidic pH (6.0 ± 0.2) and the temperature was raised to 90 °C. The viscosities of the solutions increased due to the varying concentrations of cellulose used such that 1%, 2% and 3% of cellulose solution had viscosities of 3.0, 7.6 and 7.8 cps, respectively after plasma discharge for 180 s. It is of paramount significance that the quality and efficacy of nanobiocomposites synthesized for biomedical applications are meticulously monitored, reviewed, and improved to ensure that all healthcare providers and patients have access to the most effective applications for treatment. Currently, a significant portion of the synthesized nanobiocomposites used in biomedical applications require a series of reactions which use hazardous chemicals as reducing and stabilizing agents, which may be toxic to normal cells. SPP, an eco-friendly process, which synthesizes pure and stable nanobiocomposites without employing a reducing or stabilizing agent, may provide an ideal method for producing superior medical grade nanobiocomposites.

The formation of AgNPs in the solution was further confirmed by spectrophotometry and FTIR spectroscopy. UV-Vis spectra were acquired for the 9 cellulose/AgNP biocomposites of various concentrations of cellulose and AgNO₃. Among them, the spectra of the nanobiocomposites based on 3% cellulose displayed the most significant surface plasmon resonance (SPR) band (Fig. 1). An absorbance maximum was obtained in the range of 350–440 nm, a characteristic peak of AgNPs due to SPR in the solution. A characteristic SPR band for AgNPs of C3Ag1, C3Ag3, and C3Ag5 was centered at 358 nm, 438 nm, and 440 nm, respectively, with increasing intensity of the SPR peaks as the concentration of the precursor (AgNO₃) increased. The peaks exhibited a red shift transition due to the AgNPs formed at various shapes and sizes, and the concentration of silver used. It has been reported that a blue shift of the peak could be generated by the formation of smaller particles when high concentrations of polymer were used.²⁸ In the previous report, the formation of AgNPs in the gelatin matrix was confirmed by spectrophotometry, showing a red shift of the SPR when more silver was used and a blue shift when more gelatin was used as the matrix.²⁶

Fig. 1 UV-Vis spectra of the cellulose/AgNP biocomposites: the nanobiocomposites, C3Ag1, C3Ag3, C3Ag5 had surface plasmon resonance at 350–450 nm. Inset picture shows the biocomposites turned a brown color based on the AgNP concentration in the solution by plasma discharge.

The purity of the nanobiocomposites of C3Ag1, C3Ag3, and C3Ag5 was examined by FTIR spectroscopy (Fig. 2). The characteristic peaks of cellulose such as O–H stretching, C–H stretching, C–H wagging, C–H bending, and C–O stretching were observed.³⁰ The spectra of the biocomposites of cellulose/AgNPs showed broadening, presence, and absence of peaks probably due to the cellulose molecules interacting with silver ions.³¹ The vibrations at 1409.6 cm⁻¹ and 1053.3 cm⁻¹ likely represent C–H bending and C–OH bending, respectively, which might play a functional role for the capping of AgNPs in the cellulose matrix. Additionally, vibrations at 1383 cm⁻¹ and 728 cm⁻¹ correspond to the C–H and –CH₃ groups present in the HPMC. These functional groups confer surplus stability to the matrix.

Fig. 2 FTIR spectra of the cellulose/AgNP biocomposites: characteristic vibrations (1409 cm⁻¹ and 1053 cm⁻¹) of cellulose were used as the control (C3Ag0). C–H bending and C–OH bending were due to the interaction of the functional groups with AgNPs in the nanobiocomposites of C3Ag1, C3Ag3, and C3Ag5.

3.2. Physical properties of the cellulose/AgNP biocomposites

Micro-porous 3D scaffolds of the cellulose/AgNP biocomposites synthesized using various concentrations of cellulose and AgNO₃ were prepared by lyophilization and they were cross-linked with UV irradiation to increase the water insolubility (Fig. 3A and B). Through this process, the nanobiocomposites could also be sterilized. The scaffolds containing a low percentage of cellulose (1% or 2%) showed an unstable and fragile texture, whereas that containing 3% cellulose was quite stable and firm. Moreover, it has been suggested that a high viscosity of biocomposites is suitable for scaffold formation.²⁶

Fig. 3 Structure of the 3D scaffold type biocomposites: 3D scaffolds of the control (A) and the C3Ag5 biocomposite (B) are compared. FESEM analysis showed that the micro-porous structure of C3Ag1 (C) had inter-connected cross walls with blend ends; C3Ag3 (D) and C3Ag5 (E) had inter-connected cross walls with micro-fibril edges. Spherical AgNPs were observed encrusted on the matrix of C3Ag5 (F). The size of the micro-pore was dependent on the concentration of silver.

The 3D scaffolds with micro-porous structures of the cellulose/AgNP biocomposites were observed using FESEM equipped with EDS. The microstructures of the biocomposites C3Ag1, C3Ag3, and C3Ag5 were slightly different from each other, having inter-linked and multi-walled structures (Fig. 3). Significant fibril-like structures were formed on the edges of C3Ag3 and C3Ag5, the so called micro-fibrils. Micro-fibril structures are an important feature of cellulose polymers; they could be useful for the scaffold for cell proliferation in tissue engineering applications.³² Grande et al. reported that cellulose-based nanocomposites showed good biocompatibility and had a high potential for the development of artificial skin and other types of tissues.³³ For this purpose, the pore size, pore orientation, fibre structure and fibre diameter of the 3D scaffolds were the important factors.

The mean diameters of the pores of C3Ag3 and C3Ag5 were calculated to be 68 ± 2 and 74 ± 2 μm, respectively (Fig. 3D and E). The pore size was dependent on the amount of cellulose, which affected the viscosity of the biocomposites.²⁶ The cellulose biocomposites synthesized in this study had much higher viscosities (3.0–7.8 cps) and larger pore sizes than gelatin biocomposites (2.4–4.8 cps; 17.67 ± 7.2 to 26.52 ± 12.8 μm). When gelatin biocomposites were synthesized by longer plasma discharge (780 s), they showed lower viscosity and smaller pore size than those synthesized by shorter discharge (180 s). This could be due to the degradation of the gelatin molecules into smaller molecules, which then resulted in the deformation of connected micropores.^26,34 The topographic evaluation identified the location of spherical shaped AgNPs, ranging from 15–20 nm in size, that were encrusted on the cellulose matrix (Fig. 3F).

EDS analysis showed that elements of C, O, and Ag in the constitution of the C3Ag5 biocomposites and the percentage of the elements varied according to the concentrations of cellulose and silver (Fig. 4). The Ag element in the C3Ag5 biocomposite was estimated as 14.19% (Fig. 4B), while no signal of Ag was detected in the control (Fig. 4A). The elements, percentage, atomic weight, and series of the biocomposites are listed in the inset table of Fig. 4.

Fig. 4 EDS analysis of the C3Ag5 biocomposites: elemental signal and percentage of cellulose without AgNPs (A) and cellulose/AgNP biocomposite (B).

The morphology and size of the AgNPs in the biocomposites were examined by TEM (Fig. 5). In both of the C3Ag3 and C3Ag5 biocomposites, spherical AgNPs with sizes of 5–30 nm in diameter were observed to be well distributed throughout the cellulose matrix without agglomeration, but not in C3Ag1 (Fig. 5A–C). Mostly, the size of the AgNPs was <15 nm and the mean particle size of C3Ag1, C3Ag3, and C3Ag5 was approximately 14.17, 11.35, and 11.08 nm, respectively. It has been reported that polymers may act as capping agents for the stabilization of AgNPs.³¹ Rai et al. discussed that the size and shape of the nanoparticles are important factors for the surface chemistry and antimicrobial property.²⁷ AgNPs smaller than 10 nm in diameter were reported to exhibit electronic effects when they interact with bacteria, thereby enhancing the reactivity. Truncated triangular nanoparticles exhibited the most effective antibacterial activity, followed by spherical, and then rod shaped ones. The size distribution of the biocomposites is shown in Fig. 5D. A more uniform size of AgNPs could be obtained when a higher amount of AgNO₃ was used in the SPP.

Fig. 5 TEM analysis of the AgNPs in the biocomposites: the size and shape of the AgNPs in the biocomposites were observed after the removal of cellulose. The C3Ag1 biocomposite had agglomerated particles with an average size of 14.17 nm (A); C3Ag3 had anisotropic spherical particles of 11.35 nm (B); and C3Ag5 had stable spherical particles of 11.08 nm (C). The graph represents the size distribution of the particles in the 3 nanobiocomposites (D).

3.3. Antibacterial activity of the cellulose/AgNP biocomposites

Based on the physical and chemical characterization of the biocomposites (Table 2), antimicrobial assays were carried out using C3Ag1, C3Ag3, and C3Ag5. The cellulose/AgNP biocomposites were analysed for their antibacterial properties against gram negative (E. coli, P. aeruginosa, V. parahaemolyticus) and gram positive (S. aureus and B. cereus) pathogens, most of which can cause infectious disease in humans and animals via food or water poisoning. All the scaffolds of C3Ag1, C3Ag3 and C3Ag5 were subjected to agar diffusion analysis for their antibacterial activities, monitoring the inhibition of bacterial growth by the formation of clear zones around the discs of the scaffolds (6 mm in diameter) placed on Műller-Hinton agar plates (Fig. 6A and B). C3Ag0 containing no AgNPs was used as a control. The zone of growth inhibition caused by C3Ag5 was 15 mm for E. coli, 14 mm for P. aeruginosa, 13 mm for B. cereus, and 12 mm for V. parahaemolyticus and S. aureus. Generally, gram negative bacteria were inhibited more for their growth than gram positive bacteria. However, in this test, the AgNPs embedded in the biocomposite discs probably were not in direct contact with the cells on the plates, indicating that the size of the clear zone might not be proportional to the antibacterial efficacy of the biocomposites. The antibacterial effect of the AgNPs was likely to be exhibited through the particle itself by turning into Ag⁺ ions and generating reactive oxygen species. They also would disturb the growth signaling pathway inside the bacterial cell by modulating the tyrosine phosphorylation of proteins that are important for cell viability.³⁵

Table 2 Comparison of the properties of the nanobiocomposites synthesized using various concentrations of cellulose and silver

Nanobiocomposites	Scaffold formation	Antimicrobial property	Nanoparticles formed
Cellulose (1%)
1 mM, 3 mM, 5 mM	Fragile	Poor	Agglomeration
Cellulose (2%)
1 mM, 3 mM, 5 mM	Fragile	Intermediate	No agglomeration
Cellulose (3%)
1 mM, 3 mM, 5 mM	Stable and firm	Excellent	No agglomeration

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Fig. 6 Zones of growth inhibition formed by the cellulose/AgNP discs: various degrees of anti-biogram pattern were observed by the C3Ag1, C3Ag3 and C3Ag5 discs against various pathogenic microorganisms, E. coli (A), S. aureus (B), C. albicans (C), and A. parasiticus (D). The table compares the sizes of the zones of inhibition formed by the discs against various pathogens.

The MIC of the cellulose/AgNP biocomposites was also determined using C3Ag5. LB broth (5 ml) was inoculated with 1/50 volume of overnight culture of each pathogen and various concentrations of AgNPs (0–81.6 μg ml⁻¹) were added to each tube. Then, the tubes were incubated at 37 °C for 24 h with gentle shaking (150 rpm) and the turbidity of each tube was visually observed. The biocomposites of C3Ag5 showed a MIC of 5.1 μg ml⁻¹ for gram negative bacteria (E. coli, P. aeruginosa, and V. parahaemolyticus), and MICs of 15.4 and 20.4 μg ml⁻¹ for the gram positive bacteria B. cereus and S. aureus, respectively (Table 3). Generally, gram positive bacteria are considered to be more resistant to antibacterial agents than gram negative bacteria due to the thicker cell walls they have. In a previous study, a gelatin/AgNP biocomposite (G3Ag5) showed a MIC of 20 μg ml⁻¹ for E. coli and 40 μg ml⁻¹ for S. aureus,²⁶ suggesting that the cellulose/AgNP biocomposite was more effective as an antibacterial agent.

Table 3 MICs of the cellulose/AgNP biocomposites

Microorganisms	MIC: C3Ag5 AgNPs (μg ml^−1)
E. coli	5.1
P. aeruginosa	5.1
V. parahaemolyticus	5.1
S. aureus	20.4
B. cereus	15.3
C. albicans	81.6
A. parasiticus	255.0

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The kinetics of CFU reduction by the biocomposites was examined using two bacteria, E. coli and S. aureus, by a time course assay of the bactericidal effect for 16 h (Fig. 7). The E. coli cells were affected more drastically than those of S. aureus. Almost all the E. coli cells (99.9%) were killed in 2 h incubation by all types of the biocomposites tested (Fig. 7A), while the CFU of S. aureus was reduced gradually during a period of 16 h (Fig. 7B). Complete reduction of the E. coli and S. aureus cells was observed for 4 and 16 h of incubation, respectively, under the conditions of the experiments (Fig. 7). For S. aureus, the more AgNPs present, the faster the reduction rate of CFU was. Silver probably played an active role in inhibiting the bacterial growth by binding covalently to the cell surface and eventually disrupting the cell membrane.³⁶ An attached agent disrupts the cell membrane of the bacterial cells by physical and ionic phenomena.³⁶ Silver ions were reported to interact with the thiol groups of enzymes and proteins in the membrane and cytoplasm that are important for bacterial respiration and the transportation of various substances across the membrane. Moreover, silver ions have been known to be effective in preventing infection of wounds^37,38

Fig. 7 Bactericidal effect of the cellulose/AgNP biocomposites: reduction of CFU by the C3Ag5 biocomposite was examined against E. coli (A) and S. aureus (B). Both bacteria were cultured at 37 °C by adding 5.1 μg ml⁻¹ (E. coli) and 20.4 μg ml⁻¹ (S. aureus) of AgNPs.

3.4. Antifungal activity of cellulose/AgNP biocomposites

The cellulose/AgNP biocomposites were analysed for their antifungal property against two pathogens; a yeast, C. albicans and a mold, A. parasiticus. No obvious clear zone of growth inhibition was observed against both C. albicans and A. parasiticus in agar diffusion analysis (Fig. 6C and D). Especially, A. parasiticus showed immature spore formation around the biocomposite discs, suggesting that the AgNPs did not inhibit the growth, but affect differentiation of the fungus. When the samples from the yellow or green part of the plate were examined under a microscope, much less spores were present in the yellow part than in the green part. Therefore, the biocomposites were not likely to be effective on the retardation of eukaryotic cell growth as much as they were on bacterial growth. The MIC of the biocomposites against C. albicans and A. parasiticus was determined as 81.6 μg ml⁻¹ and 255.0 μg ml⁻¹, respectively (Table 3), which were much higher than those for the bacteria. The results also suggested that the biocomposites did not affect fungal growth as effectively as bacterial growth. The results implied that the biocomposites might be safe to human cells at the concentrations effective for killing bacterial pathogens.

The effect of the biocomposites on the hyphal growth of A. parasiticus was tested by placing a block of the fungal hyphae (6 mm) on a PDA plate that had been smeared with C3Ag5 biocomposites (Fig. 8). In 3 days of incubation at room temperature, hyphal growth began to be observed around the fungal block. In 7 days, the fungal hypha was extended to ∼39 mm in diameter with green pigmentation, indicating that sporulation was in progress on the control plate with no biocomposite smeared (Fig. 8A). However, the growth was retarded by the biocomposites, showing growth to a diameter of <11 mm and taking a yellow color during the same period of culture (Fig. 8B–D). Previously, the damage of spores by nanocomposites was reported to be associated with the disruption of the cell wall, leading to a leakage of the cytoplasmic content and, subsequently, to cell death.^39,40 This effect was more pronounced by increasing the AgNP content in the films.³⁹

Fig. 8 Effect of the cellulose/AgNP biocomposites on the hyphal growth of A. parasiticus: on the center of a PDA plate smeared with no biocomposite (A), C3Ag1 (B), C3Ag3 (C), or C3Ag5 (D), a small block of the fungal hyphae (6 mm in diameter) was placed and incubated for 3 days at 25 °C.

4. Conclusion

Cellulose/AgNP biocomposites were successfully synthesized using various concentrations of cellulose and AgNO₃ by an eco-friendly one-step process of solution plasma. AgNPs were synthesized in the cellulose matrix by discharging plasma for a very short time of 180 s in the solution. The advantage of SPP for the generation of nanoparticles is not only its rapidity but also there is no need for additional hazardous chemicals such as reducing or stabilizing agents. 3D scaffolds of the biocomposites were formed by a simple lyophilization process. Analyses on the micro-porous structure, elemental percentage, intensity, and purity of the nanobiocomposites confirmed their physico-chemical properties that were suitable for stability and functionality as reactive agents. Spherical nanoparticles that were well distributed without agglomeration were observed by TEM and they had a size range of 5–30 nm in diameter. The sizes of the pores and nanoparticles in the biocomposites were likely to be modulated by the concentrations of cellulose (the matrix) and AgNO₃ (the precursor) in the solution. The cellulose/AgNP biocomposites exhibited a broad spectrum of antimicrobial activity against various pathogens, being most effective against gram negative bacteria, and then gram positive bacteria in the order C. albicans and A. parasiticus. The two fungi were quite resistant to the nanobiocomposites, implying that they might not be toxic to human cells. The results suggested that the cellulose/AgNP biocomposites have potential for application as a topical antibacterial agent or wound dressing material with antibiotic activity that does not induce drug resistance among pathogenic bacteria. From all the results obtained in this study, SPP seems to be the most effective and safest way to synthesize polymer based metal nanoparticle composites that can be applied to biomedicine.

Acknowledgements

This study was supported by Incheon National University, Republic of Korea.

References

1. D. MubarakAli, J. Arunkumar, K. HarishNag, K. A. SheikSyedIshack, E. Baldev, D. Pandiaraj and N. Thajuddin, Colloids Surf., B, 2013, 103, 166.
2. M. Jeyaraj, G. Sathishkumar, G. Sivanandhan, D. MubarakAli, M. Rajesh, R. Arun, G. Kapildev, M. Manickavasagam, N. Thajuddin, K. Premkumar and A. Ganapathi, Colloids Surf., B, 2013, 106, 86.
3. T. B. Henry, F. M. Menn, J. T. Fleming, J. Wilgus, R. N. Compton and G. S. Sayler, Environ. Health Perspect., 2007, 115, 1059.
4. S. C. Kim, J. W. Kim, G. J. Yoon and S. W. Nam, Curr. Appl. Phys., 2013, 13, 548.
5. Y. Wei, X. Zuo, X. Li, S. Song, L. Chen, J. Shen, Y. Meng, Y. Zhao and S. Fanf, Mater. Res. Bull., 2014, 53, 145.
6. P. Pootawang, S. C. Kim, J. W. Kim and S. Y. Lee, J. Nanosci. Nanotechnol., 2013, 13, 589.
7. O. Takai, Pure Appl. Chem., 2008, 80, 2003.
8. P. Prasertsung, S. Damrongsakkul and N. Saito, Polym. Degrad. Stab., 2013, 98, 2089.
9. X. Ma, F. Wu, J. Roth, M. Gell and E. H. Jordon, Surf. Coat. Technol., 2006, 201, 4447.
10. P. Pootawang, N. Saito, O. Takai and S. Y. Lee, Nanotechnology, 2012, 23, 395602.
11. I. A. Hamarneh, P. Pedrow, A. Eskhan and N. A. Lail, Surf. Coat. Technol., 2013, 234, 33.
12. T. Demina, D. Z. Zotova, M. Yablokov, A. Gilman, T. Akopova, E. Markvicheva and A. Zelenetskii, Surf. Coat. Technol., 2012, 207, 508.
13. J. Julale and V. Scholtz, Clinical Plasma Medicine, 2013, 1, 31.
14. E. Omurzak, J. Jasnakunov, N. Mairykova, A. Abdykerimova, A. Maatkasymova, S. Sulaimankulova, M. Matsuda, M. Nishida, H. Ihara and T. Mashimo, J. Nanosci. Nanotechnol., 2007, 7, 3157.
15. S. M. Kim, G. S. Kim and S. Y. Lee, Mater. Lett., 2008, 62, 4354.
16. P. Pootawang, N. Saito and S. Y. Lee, Nanotechnology, 2013, 24, 055604.
17. J. H. Kim, S. Yun, H. U. Ko and J. Kim, Curr. Appl. Phys., 2013, 13, 897.
18. S. H. Jin, S. M. Kim, S. Y. Lee and J. W. Kim, J. Nanosci. Nanotechnol., 2014, 14, 8094.
19. P. Pootawang and Y. S. Lee, Mater. Lett., 2012, 80, 1.
20. P. Pootawang, N. Saito, O. Takai and S. Y. Lee, Mater. Res. Bull., 2012, 47, 2726.
21. P. Pootawang, N. Saito, O. Takai and Y. S. Lee, Nanotechnology, 2012, 23, 395602.
22. L. Chen, T. Mashimo, H. Okudera, C. Iwamoto and E. Omurzak, RSC Adv., 2014, 4, 28679.
23. L. Chen, T. Mashimo, E. Omurzak, H. Okudera, C. Iwamoto and A. Yoshiasa, J. Phys. Chem. C, 2011, 115, 9370.
24. Z. Kelgenbaeva, E. Omurzak, S. Takebe, Z. Abdullaeva, S. Sulaimankulova, C. Iwamoto and T. Mashimo, Jpn. J. Appl. Phys., 2013, 52, 11NJ02,  DOI:10.7567/jjap.52.11nj02.
25. B. H. Lee, H. J. Kim and H. S. Yang, Curr. Appl. Phys., 2012, 12, 75.
26. S. C. Kim, S. M. Kim, G. J. Yoon, S. W. Nam, S. Y. Lee and J. W. Kim, Curr. Appl. Phys., 2014, 14, S172.
27. M. Rai, A. Yadav and A. Gade, Biotechnol. Adv., 2009, 27, 76.
28. P. Pootawang, N. Saito and O. Takai, Mater. Lett., 2011, 65, 1037.
29. J. McFarlands, JAMA, J. Am. Med. Assoc., 1907, 14, 1176.
30. C. Chinkap, L. Myunghee and K. C. Eun, Carbohydr. Polym., 2004, 58, 417.
31. D. MubrakAli, J. Arunkumar, M. Rahuman Sheriff, D. Pandiaraj, K. A. ShiekSyedIshack and N. Thajuddin, Pharm. Nanotechnol., 2013, 1, 78.
32. S. M. Oliveira, R. L. Reis and J. F. Mano, ACS Biomater. Sci. Eng., 2015, 1, 2.
33. C. J. Grande, F. G. Torres, C. M. Gomez and M. Carmen Bano, Acta Biomater., 2009, 5, 1605.
34. K. Nakagawa, M. Iwamoto, A. Nakajima, K. Shono and K. J. Satoh, Colloid Interface Sci., 2004, 278, 198.
35. S. Shrivastava, T. Bera, A. Roy, G. Singh, P. Ramachandrarao and D. Dash, Nanotechnology, 2007, 18, 225103.
36. S. Sarkar, A. D. Jana, S. K. Samnata and G. Mostafa, Polyhedron, 2007, 26, 4419.
37. S. L. Percival, P. G. Bowler and D. Russel, J. Hosp. Infect., 2005, 60, 1.
38. J. B. Wright, K. Lam, D. Habsen and R. E. Burrell, Am. J. Infect. Control, 1999, 24, 344.
39. V. Gopinath and P. Velusamy, Spectrochim. Acta, Part A, 2013, 106, 170.
40. D. MubarakAli, N. Thajuddin, K. Jeganathan and M. Gunasekaran, Colloids Surf., B, 2011, 85, 360.

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