Quaternized N-chloramine coated magnetic nanoparticles: a trifecta of superior antibacterial activity, minimal residual toxicity and rapid site removal

Abstract

We propose a new and highly effective tool to add to the ever shrinking toolbox for combating infections caused by antibiotic resistant bacteria; N-chloramine and quaternized N-chloramine were coated onto iron-oxide magnetic nanoparticles to generate antibacterial MNPs. Two differently-sized primary iron-oxide nanoparticles (3 nm and 10 nm) were synthesized and coated with silica and (3-chloropropyl)triethoxysilane, allowing subsequent introduction of N-chloramine precursors – dimethyl hydantoin (DMH) and quaternized dimethyl hydantoin (QDMH). The functionalized MNPs (MNP@DMH and MNP@QDMH) have a clear core–shell structure as evidenced by TEM images. Fe₃O₄ was identified (by combining X-ray diffraction with Mössbauer spectroscopy) to be the iron oxide in the 10 nm MNPs, while γ-Fe₂O₃ and Fe₃O₄ were the 3 nm MNP's oxide phases. Both MNPs (3 nm and 10 nm) have good magnetic responses, with saturation magnetizations of 40 ± 4 emu g⁻¹ and 65 ± 2 emu g⁻¹, respectively. Chlorination activated the antibacterial function and yielded two antibacterial MNPs: MNP@DMCl and MNP@QASCl. At the equivalent [Cl⁺] of 50 ppm, both coatings demonstrated fast inactivation of the model bacteria methicilin-resistant Staphylococcus aureus (MRSA) and multi-drug resistant (MDR) Pseudomonas aeruginosa. For either size of primary MNPs, MNP@QDMCl is more effective than MNP@DMCl. A hand-held magnet could quickly remove >99% of the functionalized MNPs from a wound simulant within 2 minutes.

---

1. Introduction

At the beginning of the 20th century, wound infections were the leading cause of death worldwide. The development of antibiotics contributed significantly to the decreased rate of wound infections in 1940s.¹ However, now the ever growing resistance to antibiotics has reached a critical tipping point, weakening and even incapacitating the major antibiotic drugs that are currently used. Infections, especially those caused by antibiotic-resistant Gram-positive and Gram-negative bacteria such as methicilin-resistant Staphylococcus aureus (MRSA) and multidrug-resistant (MDR) Pseudomonas aeruginosa, are of significant concern as detailed in the National Nosocomial Infections Surveillance.² A recent study published in Lancet Infectious Diseases revealed that bacterial resistance against the last-line antibiotic polymyxins can be spread between organisms through horizontal gene transfer.³ The gene responsible for bacterial resistance to polymyxin (MCR-1) exists in a variety of bacteria retrieved from pigs and people in South China.³ Clearly, there is a driving need to develop new strategies to cope with this rising antibiotic resistance. The use of topical biocides for the treatment of infected wounds could help reduce the usage of antibiotics. Biocides are chemical substances or microorganism that destroy or inhibit the growth of microorganisms. Biocides often have multiple microbial targets and a broader antimicrobial spectrum in comparison to antibiotics. Among the broad-spectrum biocides, N-chloramines are attracting considerable interest because of their unique properties that include powerful antibacterial activity, long-term stability, regenerability, and an innate lack of bacterial resistance.⁴ N-Chloramines attack multiple targets in bacterial cells causing irreversible damage to vital sites of the cells from the +1 oxidation state of chlorine's N–Cl bond. For this reason, N-chloramines are less likely to induce bacterial resistance. The use of these broad-spectrum biocides in the wound, however, has some disadvantages. While N-chloramines are cytotoxic to bacteria, they are also cytotoxic to skin cells responsible for wound healing (e.g. leukocytes, keratinocytes and fibroblasts) due to a lack of target specificity. But, the toxicity of the biocides towards both bacteria and skin cells is time and concentration dependent, so that timely removal of biocides from the wound might be an option to minimize their cytotoxicity and promote wound healing. Advances in the size and shape-selective synthesis of biocompatible iron-oxide nanoparticles point the way to a novel magnetic separation system if one can coat the nanoparticles with biocides.

Iron-oxide nanoparticles have very good biocompatibility and low cytotoxicity. Synthesis can be very low cost and “green” (both in synthesis and disposal).⁵ Functionalization of iron-oxide nanoparticles with biocides is an area of significant research interest. Due to the challenge of coating individual 5–10 nm primary iron-oxide nanoparticles, the previous efforts focused on trapping 5–10 nm primary iron-oxide nanoparticles in the matrix of silica or poly(styrene-acrylic acid) to yield 134–228 nm antibacterial coated magnetic particles.^6–8 As the biocide is grafted onto the surface of nanoparticles, it allows for a prolonged, localized and targeted effect on an infected wound. In this way, higher antimicrobial concentrations can be applied to target cells. Keeping in mind that the antimicrobial performance of N-chloramine based materials is strongly dependent on materials' surface area and contact time of microorganism with the material, the smaller proposed antimicrobial nanoparticles have the larger overall surface area leading to a more potent antimicrobial activity. Also, the bioactivity of antimicrobial nanoparticles is reportedly associated with surface charge. For example, positively charged silver nanoparticles were found to present superior antibacterial activity compared to neutral and negatively charged silver nanoparticles.^9,10

In this paper, instead of synthesizing antibacterial magnetic nanoparticles with large sizes (>100 nm), we present the synthesis of core–shell magnetic nanoparticles (MNPs) of biocompatible iron-oxides 3 to 10 nm in diameter, coated with silica, and functionalized with the N-chloramine moiety that have an overall size of 20 to 40 nm, depending on parent particle and coatings. To help boost the antimicrobial effect of the MNPs, a positive charge was introduced onto the MNPs silica coating. The positively charged 10 nm MNPs demonstrated superior antibacterial activity, and could be easily recovered from a simulated wound exudate with a simple magnet.

2. Experimental

2.1. Chemicals and reagents

All solvents and chemicals, such as FeCl₃·6H₂O, FeCl₂·4H₂O, tetraethylorthosilicate (TEOS), (3-aminopropyl) triethoxysilane (APTES), (3-chloropropyl)triethoxysilane (CPTES), and 5,5-dimethylhydantoin were purchased from either Aldrich or Fisher and used without further purification unless otherwise noted.

2.2. Small (3 nm diameter) Fe-oxide particles

8.67 g of FeCl₃·6H₂O and 3.14 g FeCl₂·4H₂O were dissolved in 25 mL 0.4 M HCl. The Fe solution was added dropwise to 250 mL 1.5 M NaOH over 30 min at room temperature. The nanoparticles were isolated using a strong magnet and rinsed three times with DI water, and once with 500 mL 0.01 M HCl. Nanoparticle stock solution was brought to a final total volume of 40 mL for the SiO₂ coating.

2.3. Large (10 nm diameter) Fe-oxide particles

2.70 g FeCl₃·6H₂O and 1.00 g FeCl₂·4H₂O were dissolved in 50 mL DI H₂O. 25 mL of 1.5 M NaOH was added dropwise until a pH of 12 was reached. 100 μL of oleic acid was added, and the solution was heated to 80 °C for one hour. The nanoparticles were isolated using a strong magnet and rinsed three times with DI water. Nanoparticle stock solution was brought to a final total volume of 190 mL for the SiO₂ coating step.

2.4. SiO₂ coating

160 mL ethanol and 5 mL (30 wt%) ammonia solution was added to nanoparticle stock solution. The entire mixture was sonicated for 5 min, and 1 mL TEOS was added immediately after sonication. The mixture was allowed to stir at room temperature for 16 h. The nanoparticles were isolated using magnetic separation, and rinsed with DI H₂O three times.

2.5. CPTES coating

1 g of MNP was placed in a vial. 10 mL of ethanol, 10 mL of water and 2 mL of ammonia were added into the vial. With a magnetic stirrer and stirring bar, the suspension was stirred thoroughly. Then, 2 g of CPTES was added drop-by-drop. The suspension was stirred for 3 h at room temperature. Then, the modified MNPs were washed 3 times with ethanol and 3 times with water. The resultant modified MNP is referred to as MNP@CPTES.

2.6. Synthesis of DMH–K⁺

8 g (62.4 mmol) of 5,5-dimethyl hydantoin, 4.12 g (62.4 mmol) of potassium hydroxide and 40 mL of ethanol were added to a round bottom flask. The solution was stirred and heated at reflux for 2 hours, followed by evaporating and drying the solvent under vacuum for 16 hours.

2.7. Synthesis of DMH–CH₂–N(CH₃)₂

6.6 g (0.05 mol) of 5,5-dimethyl hydantoin, 4.1 g of dimethylamine hydrochloride, 4 mL of formaldehyde (37%), and 20 mL of methanol were added to a round bottom flask; 2 g of sodium hydroxide were added to the mixture and the solution stirred at room temperature for 2 hours. The organic solvent was evaporated under vacuum afterwards. The residual product was diluted with water, and extracted with ethyl acetate. The organic phase was then evaporated and dried under vacuum to produce 4.7 g of pure product (25.4 mmol, yield = 51%).

¹H NMR (CDCl₃, 300 MHz, δ) 7.08 (m, 1H), 4.32 (s, 2H), 2.29 (s, 6H), 1.41 (s, 6H); ¹³C NMR (CDCl₃, 75 MHz, δ) 178.6, 157.4, 60.7, 58.7, 42.7, 25.2.

2.8. Surface modification of MNP@CPTES (3 nm and 10 nm MNPs)

The resulting MNP@CPTES was washed with DMF beforehand. Then, the volume was adjusted to 20 mL (DMF). 10 mL from the suspension were mixed with 1.5 g of DMH–K⁺, and 10 mL with 1.5 g KBr and 1.5 g of DMHCH₂N(CH₃)₂ affording respectively MNP@DMH and MNP@QAS1CDMH. The mixtures are stirred overnight at 120 °C. Then the MNPs were washed 3 times with ethanol and 3 times with water. The resultant modified MNPs are referred to as 3/10 nm MNP@DMH and MNP@QAS, respectively.

2.9. Chlorination and titration

The MNPs were immersed in 5 mL of bleach solution (2000 ppm pH adjusted to 8). The suspensions were mixed at room temperature for 1 h. Then, the particles were washed 4 times with water in order to remove the residual chlorine. The resulting MNPs are referred to as 3/10 nm MNP@DMCl and MNP@QASCl.

To quantitatively assess the loaded active chlorine [Cl⁺] on MNPs, a redox titration was adopted. The MNPs were mixed with 10 mL sodium thiosulfate solution (0.001 N) and 30 mL of water. The suspension was stirred for 30 min. Then, 2 mL of acetic acid at 5% was added and the remaining sodium thiosulfate was titrated using iodine solution at 0.001 N.

2.10. Antibacterial test

Muller Hinton broth and tryptic soy agar (TSA) were used to culture MDR P. aeruginosa, and tryptic soy broth (TSB) and TSA were used to culture MRSA. After being cultured on agar plates, the logarithmic-phase cultures were prepared by initially suspending several colonies in PBS (pH 7.4, 0.05 M) at a density equivalent to a 0.5 McFarland standard. The bacteria suspension was diluted by 100 times. 20 μL of the diluted P. aeruginosa and MRSA suspensions were further diluted into 60 mL of Muller Hinton broth and TSB broth, respectively. After 18 hours culturing at 37 °C, the concentration of the bacteria went up to 10⁸ CFU mL⁻¹. The suspensions were again diluted by 100 times in PBS, yielding a starting inoculum of 10⁶ CFU mL⁻¹ (total volume 20 mL). To maintain identical active chlorine content ([Cl⁺] = 50 ppm) in all the MNP samples, the amount of particles used in the antibacterial test was calculated based on the titration results. The MNP samples were placed in sterile tubes; 1 mL of the bacterial suspension was then added to the samples. The mixtures were sonicated for 10 seconds, vortex and then shaken (Twist shaker) for the duration of the experiment. After the contact had reached a predetermined time interval, 100 μL cell suspension were withdrawn; 0.9 mL PBS and 50 μL 1 N sodium thiosulfate were then added to quench the bactericidal effect. A quenched suspension was then serially diluted in PBS (10 times less concentrated than the previous one) and 100 μL of each dilution was placed onto the agar plates. The same procedures were applied to the blanks as the controls, with same matrices but without actual samples. The bacterial colonies on the agar plates were enumerated after being incubated at 37 °C for 24 hours. Percentage reduction of bacteria (%) = (A − B)/A × 100; log(reduction) = log(A/B) where A was the number of bacterial colonies in the control (CFU mL⁻¹), and B the number of bacteria colonies under the effect of the synthesized compounds.

2.11. Removal of MNPs from wound exudate simulant

A wound exudate simulant was made with the following recipe: 6.8 g L⁻¹ NaCl, 2.2 g L⁻¹ KCl, 25 g L⁻¹ NaHCO₃, 3.5 g L⁻¹ KH₂PO₄ and 20 g L⁻¹ BSA. Appropriated coated MNPs (described below) were then suspended in the simulant in a 3 mL cuvette. A stack of five 1-inch diameter rare-earth magnets was then used to pull MNPs out of the solution. The amount of MNPs remaining in the simulant at each time (15, 30, 60, 90, and 120 seconds) was determined based on the absorbance at wavelength 400 nm using the corresponding concentration/absorbance calibration curves.

2.12. Instrumentation

For every measurement made to test the biocide coated MNPs, a fresh sample was prepared from the same original lot used for study, after vortex mixing to ensure homogeneity and avoid issues such as change in concentration due to chaining or interaction with container, contamination, magnetic field history changing domain structure. FT-IR spectra were recorded on a Thermo Nicolet iS10 FT-IR Spectrometer. Zeta potential (0.1 mg mL⁻¹ of fragment suspension; Smoluchowski mode) was determined by dynamic light scattering using a Brookhaven ZetaPALS potential analyzer. NMR spectra were acquired at room temperature in 5 mm NMR tubes on a Bruker Avance 300 MHz NMR spectrometer. Transmission electron microscopy (TEM) images were collected using a JEOL-ARM200F at 200 kV. Nanoparticle samples were diluted with methanol and a droplet was placed and dried on copper-coated TEM grids. Dynamic light scattering (DLS) measurements to ascertain the hydrodynamic particle size distributions were measured at room temperature using a Photocor ​Complex system with the nanoparticles suspended in water. Powder X-ray diffraction (XRD) measurements were carried out on a Bruker D8 Discover with DaVinci in Bragg–Brentano geometry using Cu Kα radiation with a rotating stage and a zero-background sample holder. Magnetometry was performed on powder (dried) samples mounted in polycarbonate gel caps at 300 K with fields up to 5 T using a SQUID-based Quantum Design MPMS XL-5. Mössbauer spectra at room temperature were collected using a WissEl spectrometer with a 10 MBq ⁵⁷CoRh source in constant acceleration mode. The spectrometer was velocity calibrated using a 6 μm thick α-Fe foil at room temperature.

3. Results and discussion

Scheme 1 depicts the synthesis route of the N-chlorohydantoin coated Fe-oxide@SiO₂ core/shell nanoparticles. As the first step, we synthesized primary Fe-oxide nanoparticles with two sizes (nominally 3 nm and 10 nm diameter, determined from X-ray diffraction and transmission electron microcopy, as discussed below) by co-precipitation of Fe II and Fe III salts following Massart's method.¹¹ The Fe-oxide nanoparticles were then coated with a thin SiO₂ shell by hydrolysis condensation of TEOS (Stöber process) to give Fe-oxide@SiO₂ nanoparticles. The SiO₂ shell prevents Fe-oxide nanoparticles from agglomerating in aqueous solution, and provides abundant silanol groups on the surface that enable functionalization.

Scheme 1 Synthesis of nanoparticles with N-chlorohydantoin shell and Fe-oxide MNPs core.

Fig. 1 presents typical X-ray diffraction patterns of Fe-oxide nanoparticles with and without SiO₂ coating. Diffraction patterns showed the reflections characteristic of the Fe-oxide spinel structure. All reflections are Scherrer broadened by significantly different amounts between systems, indicating that the crystallite size of the two Fe-oxide nanoparticle systems differed substantially. A Rietveld refinement of the patterns incorporating Scherrer broadening effects (to ascertain the volume averaged nanocrystallite diameters) confirmed the spinel structure (Fd3̄m) of the Fe-oxides Fe₃O₄ or γ-Fe₂O₃ as expected from Massart-based synthesis. Refined lattice parameters of a = 8.296(1) Å for the small (more broadened pattern lineshape) 3 nm diameter MNPs and a = 8.374(1) Å for the large 10 nm diameter MNPs are also consistent with a spinel Fe-oxide (a ∼ 8.33 Å for bulk γ-Fe₂O₃ and a ∼ 8.39 Å for bulk Fe₃O₄). Keeping in mind that deviation from bulk parameters is common for nanoparticles, this difference in lattice parameters is large enough to indicate that the 3 nm MNPs are a mixture of the two oxide phases while the 10 nm MNPs are Fe₃O₄. This has implications on the magnetic response of the biocide coated MNPs (e.g. the overall magnetization of bulk γ-Fe₂O₃ is less than that of Fe₃O₄) for removal from, e.g. wounds, with an external field from a hand-held magnet. Therefore, we turned to a very composition and structure sensitive probe of Fe-compounds, Mössbauer spectroscopy (ESI†) that identify clearly that the 3 nm MNPs are a mixture of γ-Fe₂O₃ and Fe₃O₄, while the 10 nm MNPs are Fe₃O₄. The presence of a broad, amorphous “hump” between 20 and 28 degrees 2θ in Fig. 1c confirms the successful coating of Fe-oxide NPs with SiO₂, giving Fe-oxide@SiO₂ MNPs.

Fig. 1 (a) X-ray diffraction pattern of the small, 3 nm diameter Fe-oxide MNPs, and (b) of the larger, 10 nm diameter Fe-oxide MNPs. (c) A representative pattern of the SiO₂-coated (SiO₂ presents the broad, amorphous “hump” in the pattern between 20 and 28 degrees 2θ) 3 nm Fe-oxide MNPs. Red circles are the data, solid line is the refinement. Bragg markers identify the reflections of the Fd3̄m Fe-oxide phase, and blue lines are the residuals of the refinement.

Transmission electron microscopy images for the 3 nm and 10 nm Fe-oxide@SiO₂ MNPs are shown in Fig. 2. The particle sizes (Fig. 2d) were measured using ImageJ,¹² and fitted with a lognormal distribution and provided Fe-oxide core sizes of diameter D_small = 4.4 ± 0.2 nm and ln(σ_small) = 0.08 ± 0.01, and D_large = 9.6 ± 0.2 nm and ln(σ_large) = 0.07 ± 0.01 for the small and large particles, respectively, in good agreement with the 3 nm and 10 nm volume averaged particle diameters from XRD pattern refinements. It should be noted that the larger particles show also a small population of large sizes (D ∼ 35–50 nm), that was not observed for the small particles. After the magnetic particles were coated with silica, a thin layer of SiO₂ was observed in TEM (e.g. Fig. 2b).

Fig. 2 TEM images in (a) and (b) are of the 10 nm (volume averaged diameter) SiO₂ coated Fe-oxide MNPs, while (c) presents a high resolution TEM images of the 3 nm (volume averaged diameter) Fe-oxide MNPs. Note the change in size-bar of each image. (d) A typical particle size distribution, for the 10 nm Fe-oxide MNPs.

The field dependent magnetization of the MNPs (hysteresis loop measurements) for the 3 nm Fe-oxide nanoparticles, presented in Fig. 3a, show a saturation magnetization (M_s) at 300 K of 40 ± 4 emu g⁻¹, which is quite close to the value for that of nanoscale spinel Fe-oxide,¹³ γ-Fe₂O₃ (74 emu g⁻¹ in the bulk where nanoscale systems typically show a significantly reduced M_s from the bulk due to finite-size effects). A further decrease in M_s for the 3 nm Fe-oxide/SiO₂ core/shell MNPs of 25 ± 3 emu g⁻¹ (Fig. 3b) is a result of the non-magnetic silica fraction that adds to the total sample mass (but has a diamagnetic contribution that “cancels” the ferromagnetic response of the cores) and provides an estimate of the volume fraction of the SiO₂ versus Fe-oxide amounts to be 56 ± 3% (by volume), consistent with TEM results. For the 10 nm MNPs (Fig. 3c), M_s = 65 ± 2 emu g⁻¹, consistent with a much better well crystallized structure of the nanoparticles, in addition to the particle size, and in keeping with a Fe₃O₄-phase nanoparticle system (84 emu g⁻¹ in the bulk) (i.e. as identified by TEM and XRD). With the addition of the SiO₂ shell, M_s = 60 ± 1 emu g⁻¹; an estimate of the volume fraction of the SiO₂ versus Fe-oxide amounts to be 17 ± 1% (by volume), in order to introduce the N-chloramine moiety to the MNPs, a functional silane (3-chloropropyltriethoxysilane: CPTES) was incorporated onto the structure formed by the Fe-oxide/SiO₂ core/shell system. The synthesis of MNP@DMH and MNP@QAS nanoparticles was carried out in dimethylformamide (DMF) by reacting MNP@CPTES with DMH–K⁺ and (CH₃)₂NCH₂DMH, respectively, through a nucleophilic substitution. The conversion of surface bound N–H to N–Cl was achieved with sodium hypochlorite to generate antibacterial MNPs.

Fig. 3 Hysteresis loops measured at 300 K for the 3 nm (a) Fe-oxide and (b) Fe-oxide@SiO₂, and for the 10 nm (c) Fe-oxide and (d) Fe-oxide@SiO₂ nanoparticles.

After the coating of CPTES and DMH, a thick additional layer was observed clearly in TEM images for both the 3 and 10 nm MNPs, shown in Fig. 4. The thickness of the final coating was estimated to be, 1.1 ± 0.7 nm for the 3 nm Fe-oxide@SiO₂ MNPs, and 4.1 ± 0.3 nm for the 10 nm Fe-oxide@SiO₂ MNPs, also determined using ImageJ. The DLS results (discussed below) show that the size distribution of the particles before and after the coating is narrow and are consistent with these results obtained by TEM. However “clumps” of Fe-oxide@SiO₂ nanoparticles do form larger assemblies from multiple MNPs encapsulated by SiO₂ during the Stöber reaction.

Fig. 4 TEM images of the CPTES and DMH coated MNPs; (a) 3 nm and (b) 10 nm Fe-oxide core systems. Note that the final coating process can encapsulate several MNP cores, and the different size bars for image (a) and (b).

FT-IR spectra (Fig. 5) were recorded to identify the functional groups at different steps of the synthesis route. The MNPs (3 nm and 10 nm) present a characteristic peak at 577 cm⁻¹ corresponding to the vibration bond of Fe–O. SiO₂ coated particles (MNP@SiO₂​) show a peak at 3389 cm⁻¹ due to the stretching vibration of SiO–H groups. The strong and broad peak at 1037 cm⁻¹ and a shoulder peak at 1125 cm⁻¹ are assigned to the asymmetric stretching vibration mode of Si–O–Si. The coating with CPTES does not present any new peaks in the infrared spectra because of the overlapping of the bands corresponding to CPTES with the bands of the particles' SiO₂ shell.

Fig. 5 FTIR spectra of all synthesized MNPs.

Two peaks characteristic of two carbonyl groups in hydantoin were observed at 1698 cm⁻¹ and 1755 cm⁻¹ in the FTIR spectra of 3 nm and 10 nm MNP@DMH, and at 1730 cm⁻¹ and 1785 cm⁻¹ in the FTIR spectra of 3 nm and 10 nm MNP@QAS. The noticeable shift of hydantoin peaks was possibly attributed to the proximity effect of the quaternary ammonium salt to the hydantoin in the structure.

To quantify the effects of functionalization of the MNP formulations with QAS and DMH, photon correlation spectroscopy (PCS/DLS) experiments were performed to determine the nanoparticle hydrodynamic sizes and size distributions. Suspensions of the Fe-oxide/SiO₂ MNPs were made by adding 10 mL methanol to 5 mg of dried nanoparticles followed by ultrasonic agitation for 30 minutes. The functionalized samples were made also into dilute suspensions by adding 100 μL of concentrated stock suspension to 10 mL methanol. Light scattering measurements at seven scattering angles (30–120° in 15° intervals) were performed. The collected autocorrelation functions were fit to a distribution in translational Brownian relaxation time using the software DynaLS. Distributions in relaxation time were transformed to hydrodynamic diameter using the known viscosity and refractive index of methanol. The results of this fitting procedure are shown in Fig. 6 for a particular scattering angle of 45°, showing clearly an increase in the hydrodynamic size with additional functionalization. To assess more completely the hydrodynamic diameters of the nanoparticle samples, the mean sizes determined at all scattering angles were averaged, weighted by the total intensity of the scattered light recorded during each measurement (that is inevitably biased by the larger particles scattering more light). The mean hydrodynamic diameters and uncertainties (taken as the intensity weighted average deviation in the distributions) are summarized in Table 1. The increase in hydrodynamic diameter with the different coating agents tracks consistently between the core size of the MNP samples, and speaks to the “corona” surrounding the MNPs. The SiO₂ and CPTES coating could encapsulate more than one single primary MNP (as observed in the TEM shown in Fig. 4) to yield nanoparticle with diameters between 20 and 40 nm. MNP@QAS nanoparticles have slightly bigger hydrodynamic diameters than their MNP@DMH counterparts that could be ascribed to the higher degree of hydration of quaternized DMH (QAS) than DMH.

Fig. 6 Distribution of hydrodynamic diameters determined at a fixed scattering angle (45°) for the Fe-oxide@SiO₂ and fully functionalized nanoparticle systems.

Table 1 Summary of the intensity weighted average hydrodynamic diameter and uncertainty determined by photon correlation spectroscopy experiments performed at several scattering angles for each nanoparticle sample

Samples	Hydrodynamic diameter, D, (nm)	log-normal distribution width, ln(σD)
3 nm Fe-oxide@SiO2	23.7 ± 0.1	0.06 ± 0.02
3 nm Fe-oxide@DMH	28.7 ± 0.1	0.03 ± 0.01
3 nm Fe-oxide@QAS	33.2 ± 0.2	0.07 ± 0.03
10 nm Fe-oxide@SiO2	22.7 ± 0.1	0.06 ± 0.01
10 nm Fe-oxide@DMH	31.4 ± 0.1	0.03 ± 0.01
10 nm Fe-oxide@QAS	33.7 ± 0.3	0.08 ± 0.03

---

Zeta-potential (ξ) is a critical parameter that is closely related to particle surface charge. At basic pH, all the particles presented ξ = −40 mV as shown in Fig. 7, indicating good colloidal stability and low aggregation. At acidic pH, a noticeable change was observed between MNP@DMH and MNP@QAS. MNP@QAS nanoparticles presented more positive profiles at acidic pHs compared to MNP@DMH nanoparticles. The isoelectric point (IEP) refers to the pH at which particles do not carry net electrical charge. The IEP of 3 nm MNP@QAS is ∼5, higher than that 3 nm MNP@DMH. The IEP of 10 nm MNP@DMH is lower than pH 3.9 whereas 10 nm MNP@QAS has an IEP of ∼5.5. These results indicate clearly that the necessary incorporation of positive charge on the surface of the particles has occurred.

Fig. 7 Zeta-potentials of functionalized MNPs as a function of pH of the suspension medium.

The activation of antibacterial functions (conversion of N–H to N–Cl) with sodium hypochlorite is a crucial step prior to an antimicrobial test. Activation permits the loading of oxidative chlorine onto the surface of the MNPs. The activation process was performed by mixing MNP@DMH and MNP@QAS with sodium hypochlorite, which leads to the transfer of active chlorine (Cl⁺) from ClO⁻ to the hydantoin function groups on the surface of the MNPs through electrophilic substitution. To qualitatively assess this conversion, a few drops of 5% potassium iodide solution was added to the chlorinated and dried MNP nanoparticles. The successful conversion of N–H to N–Cl was evident from the appearance of a yellow-brownish color (2I⁻ + Cl⁺ → I₂ + Cl¹⁻). Table 2 presents the quantitative results of active chlorine loading ([Cl⁺] in ppm) on different testing samples. The 10 nm MNPs had lower chlorine loadings than the 3 nm MNPs. Difference in chlorine concentration between MNP@DMH and MNP@QAS is mainly attributed to the reactivity of their starting N-chloramine precursors with the C–Cl bond on MNP@CPTES. Chemically (CH₃)₂NCH₂DMH possesses a bulky structure with low degree of freedom, which makes its nucleophilic reaction with chloro-functional groups on CPTES/SiO₂ less favourable than with DMH–K⁺.

Table 2 Active chlorine loadings on various N-chloramine coated MNP samples

3 nm MNP@DMCl (ppm)	3 nm MNP@QASCl (ppm)	10 nm MNP@DMH (ppm)	10 nm MNP@QASCl (ppm)
2276 ± 85	818 ± 64	1202 ± 121	485 ± 90

---

The antimicrobial performance of N-chloramine and N-chloramine modified materials is directly proportional to the active chlorine loading and the contact time. Therefore, the concentrations of N-chloramine MNPs were adjusted to get the same [Cl⁺] (50 ppm) in the bacterial suspension to allow us to examine the effects of the MNPs size and the positive charge included into the structures. Two bacterial strains which are mainly responsible for wound associated infections were chosen as representative bacteria for the antibacterial test. They are Gram-negative bacterium MDR P. aeruginosa, and MRSA. Table 3 presents the antimicrobial efficacy of various N-chloramine coated MNPs against MRSA (non-activated MNPs and MNP@SiO₂ served as controls). The control MNPs were not effective in killing MRSA, and the reduction was less than 40% even after 2 hours of contact. 3 nm MNP@DMCl showed a total kill of MRSA (>6 log reduction) after 2 h of contact time whereas 1 h was sufficient for 3 nm MNP@QASCl to result a total kill. 10 nm MNPs exhibited the same trends observed for their smaller analogues. 10 nm MNP@DMCl presented 6-log reduction of bacteria after 2 h, and 10 nm MNP@QASCl could achieve a complete wipe-out of MRSA after only 30 min of contact. These comparative results indicate clearly the positive effect of the QAS incorporated into the MNP coating structure for the antimicrobial efficacy. The effect of the size between the 3 nm particles and the 10 nm was not obvious, which speaks to a population of several MNPs encapsulated during the SiO₂ and CPTES coating processes so that the conformational sizes were equivalent for some of the 3 and 10 nm MNP coated systems. In the previously reported effort of introducing hydantoin based N-chloramine onto 228 nm Fe-oxide@SiO₂ MNPs, 80% reduction of S. aureus was obtained after 1 hour contact, despite the lower bacterial concentration (10^5–6 cfu mL⁻¹) and the higher MNP concentration (200 mg mL⁻¹) used. 6 log reduction of MRSA achieved by 3 nm MNP@QASCl in 1 hour and 10 nm MNP@QASCl in 30 min clearly indicates the overall improvement in DMH coating and bacterial contact due to both the surface positive charge and the larger surface area of the smaller sized MNPs.

Table 3 Antimicrobial efficacy of MNPs against MRSA^a

Samples	Cl (mg mL^−1)	Particles (mg mL^−1)	Bacteria reduction at various contact times (min)
			5 min	15 min	30 min	1 h	2 h
a Inoculum concentration: 8.75–9.5 × 10^6 CFU mL^−1.
3 nm MNP@DMCl	0.05	22	9 ± 11	14 ± 16	25 ± 5	26 ± 5	100 ± 0
3 nm MNP@QASCl	0.05	61	16 ± 3	26 ± 16	21 ± 2	100 ± 0	100 ± 0
10 nm MNP@DMCl	0.05	41	1 ± 4	4 ± 6	22 ± 2	22 ± 20	100 ± 0
10 nm MNP@QASCl	0.05	103	44 ± 9	87 ± 2	100 ± 0	100 ± 0	100 ± 0
3 nm MNP@SiO2	0	61	34 ± 29	34 ± 7	9 ± 11	18 ± 9	24 ± 6
10 nm MNP@SiO2	0	103	42 ± 2	27 ± 12	41 ± 6	74 ± 1	26 ± 2
3 nm MNP@DMH	0	22	15 ± 6	10 ± 31	24 ± 12	14 ± 7	4 ± 5
3 nm MNP@QAS	0	61	26 ± 16	26 ± 12	25 ± 17	11 ± 4	12 ± 18
10 nm MNP@DMH	0	41	38 ± 4	31 ± 9	35 ± 7	40 ± 4	30 ± 16
10 nm MNP@QAS	0	103	46 ± 8	43 ± 2	43 ± 6	44 ± 5	37 ± 6

---

The antimicrobial performance of the MNPs against MDR P. aeruginosa is presented in Table 4. The non-chlorinated particles and SiO₂ coated MNPs showed a relatively high toxicity against MDR P. aeruginosa. This toxicity is proportional to the amount of the particles added in the bacterial solution. The non-chlorinated QAS showed a higher toxicity against MDR P. aeruginosa compared to the non-chlorinated DMH. These results indicate a higher affinity between MDR P. aeruginosa and MNP@QAS because of the charge attraction (consistent with the ξ-potential measurements described above). The activated MNPs had a higher antimicrobial effect than their non-activated analogues. 3 nm MNP@DMCl affected a total reduction after 2 h of contact time, whereas 3 nm MNP@QASCl resulted in a total reduction after 15 min. The 10 nm MNP coated systems exhibited the same trends where 10 nm MNP@DMCl resulted in only 3 log reduction of MRSA after 2 h and 6 log reduction was obtained by 10 nm MNP@QAS after 15 min of contact time. These comparative results also indicate an improvement the antimicrobial efficacy with the incorporation of positive charge at the surface, like the 3 nm MNP coated systems.

Table 4 Antimicrobial efficacy of MNPs against MDR P. aeruginosa^a

Samples	Cl (mg mL^−1)	Particles (mg mL^−1)	Bacteria reduction at various contact times (min)
			5 min	15 min	30 min	1 h	2 h
a Inoculum concentration: 1.56–2.63 × 10^6 CFU mL^−1.
3 nm MNP@DMCl	0.05	22	62 ± 4	64 ± 5	73 ± 3	76 ± 6	100 ± 0
3 nm MNP@QASCl	0.05	61	76 ± 3	100 ± 0	100 ± 0	100 ± 0	100 ± 0
10 nm MNP@DMCl	0.05	41	50 ± 3	59 ± 1	60 ± 4	80 ± 3	99.95 ± 0.01
10 nm MNP@QASCl	0.05	103	74 ± 7	100 ± 0	100 ± 0	100 ± 0	100 ± 0
3 nm MNP@SiO2	0	61		—		57 ± 10	29 ± 2
10 nm MNP@SiO2	0	103				70 ± 2	54 ± 10
3 nm MNP@DMH	0	22		—		37 ± 11	13.9 ± 5.8
3 nm MNP@QAS	0	61		—		85 ± 2	92 ± 1
10 nm MNP@DMH	0	41		—		54 ± 3	33 ± 1
10 nm MNP@QAS	0	103		—		70 ± 5	71 ± 3

---

Finally, the enhancement of the antibacterial activity of the MNPs is absolute with the introduction of the cationic N-chloramine onto the surfaces. A mechanism can be proposed to explain the origin of this boost. Since both Gram-positive and Gram-negative bacteria are negatively charged owing to the presence of the teichoic acid and phosphate on their membrane and cell walls, adding a positive charge on a support facilitates the adsorption of MNPs onto bacteria for a faster oxidative chlorine transfer to the target sites, leading to the most effective bacterial deaths.

Since the biocide N-chloramine is non-selective against bacteria and human skin cells, it is desirable to remove the N-chloramine based antibacterial MNPs from the wound bed after they inactivate bacteria in the infected wound to minimize residual toxicity. We conducted an experiment of recovering small and large MNP@DMH and MNP@QAS from a wound simulant with the following ingredients: 6.8 g L⁻¹ NaCl, 2.2 g L⁻¹ KCl, 25 g L⁻¹ NaHCO₃, 3.5 g L⁻¹ KH₂PO₄ and 20 g L⁻¹ BSA. As shown in Fig. 8, the tested MNPs reached >99% recovery after 60–120 seconds of applying an external magnetic field, and 10 nm MNPs showed a faster recovery (>80% recovery after 15 s) than the 3 nm MNPs (55% recovery after 15 s). The faster recovery of 10 nm MNPs might be due to the higher intrinsic magnetization of the Fe₃O₄ phase and the larger primary core size. In addition, the more rapid recovery of the 10 nm MNP@QAS compared to the 10 nm MNP@DMH is indicative of surface charge (e.g. “corona” effects) where the DMH is more strongly bound to the wound simulant – stronger static magnetic fields, or a combination of static and dynamic applied magnetic fields, should further improve the speed of recovery to offset this electrostatic charge effect.

Fig. 8 Recovery of MNPs from a wound simulant.

4. Conclusions

27–36 nm diameter N-chloramine based antibacterial MNPs were successfully synthesized with two differently sized Fe-oxide cores: 3 nm and 10 nm. The 10 nm Fe-oxide core was found to be composed of purely Fe₃O₄ whereas 3 nm Fe-oxide core is composed of the two oxide phases according to their X-ray diffraction patterns and Mössbauer spectra. As compared with N-chloramine MNPs (MNP@DMCl), quaternized N-chloramine MNPs (MNP@QAS) demonstrated more effective inactivation of both MRSA and MDR P. aeruginosa. 3 nm and 10 nm MNP@DMH and MNP@QAS could be quickly removed from the wound simulant within 2 minutes by an externally applied magnetic field. These antibacterial MNPs can be an effective tool for combating wound infections caused by antibiotic resistant bacteria.

Conflict of interest

The authors declare no competing financial interest.

Acknowledgements

The authors would like to acknowledge the funding from the Natural Sciences and Engineering Research Council of Canada (NSERC), the Collaborative Health Research Project (CHRP) program, the Canada Foundation for Innovation (CFI), and the University of Manitoba's Faculty of Science through their Initiation Grants program.

References

1. C. K. Murray, M. K. Hinkle and H. C. Yun, J. Trauma, 2008, 64(3), S221–S231.
2. The NNIS System, Am. J. Infect. Control, 2004, 32, 470.
3. Y. Liu, Y. Wang and T. R. Walsh, et al., Lancet Infect. Dis., 2015, 16, 161.
4. H. Rahma, S. Asghari, S. Logsetty, X. Gu and S. Liu, ACS Appl. Mater. Interfaces, 2015, 7, 11536.
5. V. Yathindranath, L. Rebbouh, D. F. Moore, D. W. Miller, J. van Lierop and T. Hegmann, Adv. Funct. Mater., 2011, 21, 1457.
6. A. Dong, S. Lan, J. Huang, T. Wang, T. Zhao, L. Xiao, W. Wang, X. Zheng, F. Liu, G. Gao and Y. Chen, ACS Appl. Mater. Interfaces, 2011, 3, 4228.
7. D. Alideertu, S. Yue, L. Shi, Q. Wang, Q. Cai, X. Qi, Y. Zhang, G. Gao, F. Liu and C. Harnoode, ACS Appl. Mater. Interfaces, 2013, 5, 8125.
8. L. Bromberg, E. P. Chang, T. A. Hatton, A. Concheiro, B. Magariños and C. A. Lorenzo, Langmuir, 2011, 27, 420.
9. A. Abbas, Y. Ghahramani, A. Gholami, B. Hemmateenejad, S. Dorostkar, M. Nabavizadeh and H. Sharghi, J. Nanomater., 2015, 2015, 1–8.
10. T. Silva, L. R. Pokhrel, B. Dubey, T. M. Tolaymat, K. J. Maier and X. Liu, Sci. Total Environ., 2014, 468–469, 968.
11. R. Massart, IEEE Trans. Magn., 1981, 17, 1247.
12. C. A. Schneider, W. S. Rasband and K. W. Eliceiri, Nat. Methods, 2012, 9, 671.
13. T. N. Shendruk, R. D. Desautels, B. W. Southern and J. van Lierop, Nanotechnology, 2007, 18, 455704.

---

Footnote

† Electronic supplementary information (ESI) available. See DOI: 10.1039/c6ra13389d

---