**Bi-Functional Alginate Oligosaccharide–Polymyxin Conjugates for Improved Treatment of Multidrug-Resistant Gram-Negative Bacterial Infections**

#### **Joana Stokniene 1,\*, Lydia C. Powell 1, Olav A. Aarstad 2, Finn L. Aachmann 2, Philip D. Rye 3, Katja E. Hill 1, David W. Thomas <sup>1</sup> and Elaine L. Ferguson <sup>1</sup>**


Received: 19 October 2020; Accepted: 9 November 2020; Published: 11 November 2020

**Abstract:** The recent emergence of resistance to colistin, an antibiotic of last resort with dose-limiting toxicity, has highlighted the need for alternative approaches to combat infection. This study aimed to generate and characterise alginate oligosaccharide ("OligoG")–polymyxin (polymyxin B and E (colistin)) conjugates to improve the effectiveness of these antibiotics. OligoG–polymyxin conjugates (amide- or ester-linked), with molecular weights of 5200–12,800 g/mol and antibiotic loading of 6.1–12.9% *w*/*w*, were reproducibly synthesised. In vitro inflammatory cytokine production (tumour necrosis factor alpha (TNFα) ELISA) and cytotoxicity (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) of colistin (2.2–9.3-fold) and polymyxin B (2.9–27.2-fold) were significantly decreased by OligoG conjugation. Antimicrobial susceptibility tests (minimum inhibitory concentration (MIC), growth curves) demonstrated similar antimicrobial efficacy of esterand amide-linked conjugates to that of the parent antibiotic but with more sustained inhibition of bacterial growth. OligoG–polymyxin conjugates exhibited improved selectivity for Gram-negative bacteria in comparison to mammalian cells (approximately 2–4-fold). Both OligoG–colistin conjugates caused significant disruption of *Pseudomonas aeruginosa* biofilm formation and induced bacterial death (confocal laser scanning microscopy). When conjugates were tested in an in vitro "time-to-kill" (TTK) model using *Acinetobacter baumannii*, only ester-linked conjugates reduced viable bacterial counts (~2-fold) after 4 h. Bi-functional OligoG–polymyxin conjugates have potential therapeutic benefits in the treatment of multidrug-resistant (MDR) Gram-negative bacterial infections, directly reducing toxicity whilst retaining antimicrobial and antibiofilm activities.

**Keywords:** gram-negative bacteria; multidrug resistance; polymer therapeutics; colistin; polymyxin B

#### **1. Introduction**

Antimicrobial resistance (AMR) is a significantly growing global challenge that is associated with elevated morbidity and mortality rates, high healthcare costs and >700,000 deaths annually [1,2]. Excessive use of antibiotics in animal husbandry, agriculture, and human and veterinary medicine has contributed to a dramatic increase in life-threatening multi- and pan-drug resistant bacterial infections [3]. This environmental exposure has been compounded by decreases in the development

of novel antimicrobials and it has been predicted that AMR could result in 10 million annual deaths by 2050 [4]. According to the World Health Organisation (WHO), Gram-negative bacteria such as carbapenem-resistant *Pseudomonas aeruginosa* and *Acinetobacter baumannii*, extended spectrum β-lactamase-producing and carbapenem-resistant *Klebsiella pneumoniae* and *Escherichia coli* represent a major clinical threat and burden to public health [5]. It has been estimated that Gram-negative bacterial resistance resulted in 960,000 hospital admission days in Europe in 2017 [6].

Polymyxins (Scheme 1), such as polymyxin B and colistin (polymyxin E), are a potent class of polypeptide antibiotics. Despite the clinical efficacy of colistin against Gram-negative bacteria, it is recommended for employment as an antibiotic of last resort, both to avoid resistance and, importantly, due to dose-limiting nephro- and neurotoxicity [7]. To reduce this toxicity and optimise antimicrobial activity, drug absorption and target specificity, several novel derivatives of polymyxin antibiotics are being developed [8–11]. Structural modifications have involved the N-terminal fatty acyl moiety or Dab side chains and demonstrated the importance of Dab at residue five in antimicrobial activity [12]. Progression to clinical trials of these polymyxin derivatives has, however, been limited due to their narrow spectrum of antimicrobial activity, and their cytotoxicity/poor tolerability in animal studies [13].

**Scheme 1.** Graphic structure of (**a**) colistin and (**b**) polymyxin B. The hydrophilic heptapeptide ring is linked to a hydrophobic acyl tail through a tripeptide fragment. The only structural difference between both molecules is an amino acid residue at position 6: d-leucine in colistin is replaced by d-phenylalanine in polymyxin B. Composition of the fatty acyl tail: 6-methyloctanoic acid for polymyxin B1/E1 and 6-methylheptanoic acid for polymyxin B2/E2.

Polymer therapeutics have emerged as a promising strategy to combat antimicrobial resistance, particularly when used to reinstate "old" antibiotics [14]. Conjugation of an antibiotic to a water-soluble polymer offers many advantages compared to small molecule drugs, including reduced toxicity/immunogenicity, prolonged plasma half-life and improved pharmacodynamic targeting through the enhanced permeability and retention (EPR) effect [15]. Colistin has previously been conjugated to both, dextrin [16] and poly(ethylene glycol) (PEG) [17], however, complete restoration of antibiotic activity was not achieved after amylase-unmasking of dextrin–colistin conjugates, presumably due to the presence of oligosaccharides attached to the colistin amine groups [18]. The use of alternative conjugation chemistry may offer the opportunity to optimise reinstatement of antibiotic

activity at sites of infection/inflammation [19]. Moreover, conjugation of the antibiotic to bioactive polysaccharides affords the opportunity to deliver anti-infective bi-functional polymer therapeutics.

Although alginates, like dextrin, are recognised as non-toxic by the Food and Drug Administration (FDA), their large molecular weight and lack of mammalian, alginate-degrading enzymes has limited their use in protein/peptide conjugation. More recently, a low molecular weight alginate oligosaccharide (OligoG, Mn 3200 g/mol), was extracted as a sodium salt from marine algae (*Laminaria hyperborea*) with >85% of residues being composed of α-L-guluronic acid. Although OligoG has no (MIC) value, it inhibits bacterial growth, adherence and biofilm development, and potentiates the activity of antibiotics against Gram-negative MDR pathogens [20–24]. This low molecular weight alginate also possesses hydroxyl and carboxyl functional groups that can be used for drug conjugation.

We hypothesised that conjugation of guluronic-rich, low molecular weight alginates to antibiotics, such as polymyxins, could create a bi-functional antibiotic polymer therapeutic [25]; combining the antimicrobial properties of both the antibiotic and the alginate, while simultaneously reducing systemic toxicity of the antibiotic, and facilitating size-dependent targeting by the EPR effect at the site of infection. Polymyxins were chosen as a model drug because previous studies have demonstrated that OligoG can enhance the antimicrobial efficacy of colistin against MDR, Gram-negative *P. aeruginosa* both in vitro and in vivo [26].

The aim of the study was to generate and characterise a bi-functional polymyxin conjugate using OligoG to optimise the antimicrobial function of these last resort antibiotics. A range of OligoG-polymyxin conjugates were synthesised and their physicochemical properties, in vitro cytotoxicity and biological activity characterised. Antimicrobial activity was assessed using MIC assays, growth curves, confocal laser scanning microscopy (CLSM) imaging and "time-to-kill" (TTK) studies.

#### **2. Materials and Methods**

#### *2.1. Materials*

OligoG CF-5/20 and the high molecular weight alginate PRONOVA UP MVG (>60% guluronic acid and Mw of 200,000 g/mol) were provided by AlgiPharma AS (Sandvika, Norway). The LIVE/DEAD® BaclightTM Bacterial Viability kit was from Invitrogen Molecular Probes (Paisley, UK). Pullulan gel filtration standards were from Polymer Laboratories (Church Stretton, UK). All chemicals were obtained from either Fisher Scientific (Loughborough, UK) or Sigma-Aldrich (Poole, UK) unless otherwise stated and were of analytical grade.

#### *2.2. Cell Lines and Cell Culture*

Human kidney proximal tubule cells (HK-2) were obtained from the American Type Culture Collection (ATCC) (Manassas, VA, USA) and screened to be free of mycoplasma contamination before use. Keratinocyte serum-free (K-SFM) medium (with l-glutamine), bovine pituitary extract (BPE, 0.05 mg/mL), human recombinant epidermal growth factor (EGF, 5 ng/mL), 0.05% *w*/*v* trypsin-0.53 mM ethylenediaminetetraacetic acid (EDTA) were from Invitrogen Life Technologies (Paisley, UK).

#### *2.3. Bacterial Isolates and Growth Media*

The bacterial strains (Table S1) used have been previously described [20,27]. Bacteria were grown on either tryptone soy agar (TSA) or blood agar (BA) plates supplemented with 5% *v*/*v* defibrinated horse blood. Bacterial overnight cultures were grown in tryptone soy (TS) broth and Mueller–Hinton (MH) broth was used for susceptibility testing. All media were from LabM (Bury, UK). Artificial sputum (AS) medium was prepared as previously described by Pritchard et al. [24].

#### *2.4. Synthesis of OligoG–Polymyxin Conjugates*

To synthesise amide (A)-linked conjugates (Scheme 2a), OligoG (1000 mg, 0.3 mmol), 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC; 96.8 mg, 0.5 mmol) and

N-hydroxysulfosuccinimide (sulfo-NHS; 109.6 mg, 0.5 mmol) were dissolved under stirring (15 min at 21 ◦C) in distilled water (dH2O; 10 mL). To this, colistin sulphate (146.7 mg, 0.1 mmol) or polymyxin B (144.4 mg, 0.1 mmol) was added followed by drop-wise addition of NaOH (0.5 M) until pH 8 was reached. The reaction mixture was stirred for 2 h at 21 ◦C, then stored at −20 ◦C prior to purification.

**Scheme 2.** Schematic showing steps in the synthesis of the OligoG–colistin conjugate. (**a**) Using an amide linker (OligoG–A–colistin conjugate). (**b**) Using an ester linker (OligoG–E–colistin conjugate).

To synthesise ester (E)-linked conjugates (Scheme 2b), OligoG (1000 mg, 0.3 mmol), *N*,*N* -dicyclohexyl carbodiimide (DCC; 64.5 mg, 0.3 mmol), 4-dimethylaminopyridine (DMAP; 6.4 mg, 0.05 mmol) and colistin sulphate (146.7 mg, 0.1 mmol) or polymyxin B (144.4 mg, 0.1 mmol) were dissolved while stirring overnight at 21 ◦C in anhydrous DMSO (10 mL). The reaction was stopped by pouring the mixture into excess chloroform (~100 mL). Formed precipitates were collected by filtration and dissolved in dH2O (10 mL), then stored at −20 ◦C prior to purification.

#### *2.5. Purification of OligoG–Polymyxin Conjugates*

OligoG–polymyxin conjugates were purified from the reaction mixture by fast protein liquid chromatography (FPLC) using an AKTA Purifier system (GE Healthcare; Amersham, UK) connected to a prepacked Superdex 75 16/600 GL column with a UV detector and a fraction collector (Frac-950). Data analysis was performed using Unicorn 5.31 software (2011; GE Healthcare; Amersham, UK). Samples (2 mL) were injected into a 2 mL loop using phosphate buffered saline (PBS) buffer (pH 7.4) as a mobile phase at 1 mL/min. Fractions were collected, pooled and lyophilised. Then, conjugates were re-dissolved in a minimal volume of dH2O and dialysed (1000 g/mol cut-off) against 5 × 1 L dH2O to remove PBS salts. The final conjugates were lyophilised and stored at −20 ◦C.

#### *2.6. Characterisation of OligoG–Polymyxin Conjugates*

Size exclusion chromatography with multi-angle light scattering detection (SEC-MALS) or refractive index detection (SEC-RI), were used to measure the approximate molecular weight and polydispersity of the conjugates. SEC-MALS was performed at ambient temperature on an HPLC system consisting of a solvent reservoir, on-line degasser, automatic sample injector, HPLC isocratic pump, pre-column and serially connected columns (TSKgel 4000 and 2500 PWXL). The column outlet was connected to a Dawn HELEOS-II multi-angle laser light scattering photometer (Wyatt, MO, USA) (λ0 = 663.8 nm) followed by a Shodex RI-501 RI detector. The eluent was 0.15 M NaNO3 with 0.01 M EDTA, pH 6.0 and the flow rate was 0.5 mL/min. Samples (10 mg/mL) were filtered (pore size 0.45 μm) before injection and analysed twice with injection volumes of 25 and 50 μL. A weighted specific refractive index increment (dn/dc) value was calculated from the % *w*/*w* colistin using dn/dc = 0.150 and 0.185 for alginate and colistin, respectively. Data were collected and processed using the Astra software (version 7.3.0; Wyatt, USA).

The SEC-RI system consisted of two TSK gel columns (G5000PWXL and G3000PWXL) (Tosoh, Germany) in series connected to a Gilson 133 differential refractometer (Middleton, WI, USA). Samples were prepared in PBS (3 mg/mL) and eluted using PBS (pH 7.4) as the mobile phase at a flow rate of 1 mL/min. Cirrus GPC software (version 3.2, 2006) from Polymer Laboratories (Church Stretton, UK,) was used for data analysis. Molecular weight was determined relative to pullulan molecular weight standards.

The FPLC system described above, connected to a Superdex 75 (10/300 GL) column, was also used to assess conjugate purity. Samples (3 mg/mL in PBS) were injected into a 100 μL loop at 0.5 mL/min. The area under the curve was used to estimate the percentage of free and conjugated antibiotic. The total polymyxin content of conjugates was determined by bicinchoninic acid (BCA) assay using colistin sulphate or polymyxin B standards.

Before and after OligoG conjugation, the number of available primary amine groups on colistin and polymyxin B was determined using the ninhydrin assay. First, a lithium acetate buffer (4 M) was prepared by dissolving lithium acetate dihydrate 40.81% *w*/*v* in dH2O. Acetic acid (glacial) was added to reach pH 5.2 before adjusting the final volume. Next, ninhydrin reagent was prepared by dissolving ninhydrin 2% *w*/*v* and hydrindantin 0.3% *w*/*v* in 7.5 mL of DMSO and 2.5 mL of lithium acetate buffer. Test compounds (86 μL) were diluted with ninhydrin reagent (1:1) and heated in a water bath (100 ◦C) for 15 min. Samples were subsequently cooled to room temperature and mixed with 50% *v*/*v* ethanol solution (130 μL). Then, aliquots (100 μL) were transferred into the wells of a 96-well microtitre plate and analysed spectrophotometrically at 570 nm. Calibration of the assay was achieved using ethanolamine (0–0.1158 mM).

NMR spectroscopy was used to confirm OligoG–polymyxin conjugation (Supplementary Materials).

#### *2.7. Drug Release of OligoG–Polymyxin Conjugates*

To compare the rate of degradation of OligoG–polymyxin conjugates, solutions were prepared (3 mg/mL) in either (i) PBS at pH 5, (ii) PBS at pH 7, or (iii) PBS at pH 7 containing alginate lyase from *Sphingobacterium multivorum* (1 U/mL) and incubated at 37 ◦C for 0, 2, 4, 6, 24 and 48 h. Upon collection, samples were immediately snap-frozen in liquid nitrogen and stored at−20 ◦C. Time-dependent changes in molecular weight and free polymyxin content were determined by SEC-RI and FPLC, respectively.

#### *2.8. Characterisation of In Vitro Toxicity*

A 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay was used to measure cell viability and proliferation of HK-2 cells. Cells were seeded into sterile 96-well microtitre plates at 1 <sup>×</sup> 105 cells/mL (100 <sup>μ</sup>L/well) and allowed to adhere for 24 h at 37 ◦C. The following day, the old medium was replaced with test compounds (0–1 mg/mL polymyxin base) dissolved in filter-sterilised K-SFM. After 67 h incubation at 37 ◦C, filter-sterilised MTT solution (20 μL of a 5 mg/mL solution in PBS) was added to each well and incubated for a further 5 h at 37 ◦C. Finally, the medium was carefully removed, and the formazan crystals were solubilised in DMSO (100 μL) for 30 min. Absorbance was measured at 550 nm using a Fluostar Omega microplate reader. The results are stated as percentage cell viability compared with the untreated control cells. Data are expressed as mean ± SEM (*n* = 18).

Release of the cytokine, tumour necrosis factor alpha (TNFα), by HK-2 cells (1 <sup>×</sup> 105 cells/mL) after exposure to free- and OligoG-conjugated antibiotic (0–1 mg/mL polymyxin base) was assessed using an enzyme-linked immunosorbent assay (ELISA) kit. After 72 h incubation, the 96-well microtitre plates were centrifuged (226× *g*, 3 min), the supernatant was collected, diluted with reagent diluent (1:1) and analysed with the TNFα ELISA kit according to the manufacturer's instructions (Fisher Scientific; Loughborough, UK). Plates were analysed spectrophotometrically at 450 nm. In parallel, 100 μL of K-SFM was added to the wells of the centrifuged plates containing cells and MTT assays were performed. A standard curve was used to calculate TNFα concentrations in the test samples, which were then multiplied by the dilution factor (×2) and divided by cell viability for each drug concentration (from the MTT assay). Two outliers were identified and removed using robust regression and the outlier removal (ROUT) method (Q coefficient = 0.2%). Data are expressed as mean ± SEM (*n* = 6).

#### *2.9. Antimicrobial Activity of OligoG–Polymyxin Conjugates*

The minimum inhibitory concentration (MIC) of colistin (as sulphate salt) and polymyxin B and their conjugates was determined using the broth microdilution method in MH broth in accordance with standard guidelines [28]. Test organisms were suspended in MH broth (100 <sup>μ</sup>L, 5 <sup>×</sup> 105 colony forming units (CFU)/mL) and incubated in 96-well microtitre plates in serial two-fold dilutions of the test compounds. The MIC was defined as the lowest concentration of test compound that produced no visible growth after 16–20 h. Results were expressed as mode (*n* = 3). For the purpose of calculating selectivity index (SI), MIC values lower than 0.008 were taken as the lowest concentration tested. Selective activities of the polymyxins and OligoG–polymyxin conjugates were calculated as follows:

#### Selectivity index (SI) = IC50 (μg/mL)/MIC (μg/mL).

To investigate whether alginate oligomer degradation is required for antimicrobial activity, MIC assays were also conducted in the presence of alginate lyase (1 and 10 U/mL), whereby alginate lyase was added to the MH broth during microtitre plate set up. In addition, alginate oligomer–colistin conjugates (3 mg/mL) were incubated in PBS at pH 7 containing bacterial alginate lyase (1 and 10 U/mL) at 37 ◦C for 24 h, before preparing microtitre plates as described above.

To more closely mimic in vivo environmental conditions, the antimicrobial activity of test compounds was studied in the presence of mucin, by supplementing MH broth with porcine stomach (type II) mucin (0.2 and 2% *w*/*v*) and used to set up the 96-well plates according to the standard MIC protocol. To account for turbidity caused by mucin, resazurin dye solution (30 μL, 0.01% *w*/*v* in dH2O) was added to each well and incubated for a further 3 h at 37 ◦C. Colour changes were observed and recorded.

To study the antimicrobial efficacy of test compounds under more clinically relevant conditions, the MIC protocol was performed using AS medium instead of MH broth. The plates were incubated with resazurin as described above.

A checkerboard assay was used to assess synergy of test compounds with azithromycin dihydrate. Here, stock solutions of test compounds (8 × MIC) and serial two-fold dilutions of azithromycin dihydrate (16–1/16 × MIC) were freshly prepared in MH broth. Test compound solutions (100 μL) were placed in the wells of row 1, then serially diluted along the ordinate with MH broth. Serially diluted azithromycin dihydrate solutions (50 μL) were then added to the wells in decreasing concentration along the abscissa. Each microtitre well was inoculated with the test organism (5 <sup>×</sup> <sup>10</sup><sup>5</sup> CFU/mL) and incubated at 37 ◦C for 20 h. The fractional inhibitory concentration index (FICI) was calculated by comparing the MIC values of the individual agents with the MIC value of the combined treatments [29]. Drug combinations were considered synergistic when the mean FICI was ≤0.5, additive when the FICI was between 0.5 and 2, indifferent when the FICI was between 2 and 4, and antagonistic when the FICI was ≥4 [30]. Results were expressed as median values (*n* = 3).

To study the effect of test compounds on bacterial pharmacokinetic profiles, 96-well microtitre plates were set up according to the standard MIC protocol, then placed in a Fluostar Omega Microplate Reader at 37 ◦C, and absorbance at 600 nm was measured hourly for 48 h. Results were expressed as mean values (*n* = 3). Unconjugated colistin plus OligoG, OligoG and the high molecular weight, biologically inactive alginate, PRONOVA, at equivalent concentrations used in amide-linked or ester-linked conjugates, were used as controls.

#### *2.10. Anti-Biofilm Activity of OligoG–Polymyxin Conjugates*

To analyse the effect of test compounds on biofilm formation, solutions of test compounds in MH broth were inoculated (1:10) with *P. aeruginosa* R22 (standardised to 107 CFU/mL) in a Greiner glass-bottomed optical 96-well plate. The plate was then wrapped in parafilm and incubated (37 ◦C, 20 rpm) for 24 h. The supernatant was carefully removed and replaced with 10% *v*/*v* LIVE/DEAD stain in PBS prior to imaging. CLSM of Syto 9 (λex/λem maximum, 480/500 nm) and propidium iodide (λex/λem maximum, 490/635 nm) was performed using a Leica SP5 confocal microscope with ×63 lens (under oil) and a step size of 0.79 μm. Z-stack CLSM images were analysed using COMSTAT image analysis software [31] and results were expressed as mean ± SEM (*n* = 15).

#### *2.11. Pharmacokinetic–Pharmacodynamic (PK–PD) Model*

A two-compartment static dialysis bag model (adapted from Azzopardi et al. [32]) was used to study the PK–PD profile of OligoG–colistin conjugates. First, the ability of the dialysis membrane to control diffusion of test compounds was assessed. The inner compartment (IC) contained OligoG–colistin or colistin sulphate (10 mg/mL colistin base; 5 mL) in PBS and the outer compartment (OC) contained sterile PBS (15 mL). The aseptically sealed beaker was incubated (37 ◦C, 70 rpm) for 48 h. Samples were collected at various time points from each compartment and stored at −20 ◦C prior to analysis of colistin content by BCA assay.

A modified experimental set-up was used to investigate the concentration- and time-dependent antimicrobial activity of test compounds using a TTK assay (48 h). Here, the total volume in the system was considered, with the IC containing test compounds at MIC (colistin sulphate 0.25 μg/mL; OligoG–A–colistin 0.125 μg/mL colistin base; OligoG–E–colistin 0.125 μg/mL colistin base) or 2 × MIC (OligoG–A–colistin 0.25 μg/mL colistin base; OligoG–E–colistin 0.25 μg/mL colistin base) in PBS. The OC contained MH broth inoculated with *A. baumannii* 7789 (5 <sup>×</sup> 10<sup>5</sup> CFU/mL). Samples were collected from the OC (0, 2, 4, 6, 24 and 48 h) and colony counts (CFU/mL) determined using drop counts. Treatments were considered bactericidal if the reduction in viable bacterial counts was ≥3 log10 CFU/mL (equivalent to 99.9% of the initial inoculum) and bacteriostatic if the decrease was <3 log10 CFU/mL [33,34]. Growth (no test compounds) and sterility (no bacteria) controls were also performed.

#### *2.12. Statistical Analysis*

GraphPad Prism (version 6.01, 2012; San Diego, CA, USA) was used for statistical analysis. Statistical significance was indicated by \*, where \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001 and \*\*\*\* *p* < 0.0001. Analysis of variance (ANOVA) was used to evaluate multiple group comparisons (*n* ≥ 3) followed by Dunnett's post hoc test to account for multiple comparisons.

#### **3. Results**

#### *3.1. Synthesis and Characterisation of OligoG–Polymyxin Conjugates*

The characteristics of the OligoG–antibiotic conjugates synthesised in this study are summarised in Table 1 and Table S2. Polymyxin B conjugates typically showed less drug loading (6.1–8% *w*/*w*) than the colistin conjugates (8.1–12.9% *w*/*w*). SEC-MALS, SEC-RI and FPLC analysis confirmed the presence of high molecular weight conjugates with <6% unbound drug (Figures S1 and S2, Table S3). The mean molecular weight of amide- and ester-linked conjugates (measured by SEC-MALS) was 8200–12,800 g/mol and 5200–6200 g/mol, respectively. The ninhydrin assay indicated that 2–4 amine groups were used for binding to OligoG via amide conjugation. Diffusion-ordered spectroscopy (DOSY) NMR confirmed covalent coupling of OligoG to colistin. Signals corresponding to OligoG and colistin in the samples had the same diffusion coefficient (1.26 <sup>×</sup> 10−<sup>10</sup> m2/s), indicative of covalent coupling (Figure S3). OligoG–A–colistin conjugate samples appeared to contain some free OligoG while the DOSY spectrum for OligoG–E–colistin conjugate showed the presence of both, free OligoG and colistin in the sample.


**Table 1.** Summary of the properties of the OligoG-polymyxin conjugates synthesised in this study.

Abbreviations: A, amide; E, ester; PDI, polydispersity index (given in brackets); SEC-MALS, size exclusion chromatography with multi-angle light scattering detection; N/A, not applicable.

#### *3.2. Stability of OligoG–Polymyxin Conjugates*

Both ester- and amide-linked conjugates of OligoG–colistin and OligoG–polymyxin B incubated in PBS at either pH 5 or pH 7 showed no significant decrease in molecular weight (Figure S4). Conjugates were slightly less stable at pH 7, compared to pH 5. Conversely, alginate lyase effectively triggered ~30% of colistin and ~90% of polymyxin B release (increase in % free drug) within 24 h from these conjugates at 1 U/mL. There was little difference in total drug release between amide- and ester-linked conjugates.

#### *3.3. Cytotoxicity of OligoG–Polymyxin Conjugates*

The concentration-dependent cytotoxicity of unmodified antibiotics, OligoG and OligoGpolymyxin conjugates in HK-2 cells is shown in Figure 1. OligoG was not cytotoxic at <10 mg/mL. Cytotoxicity was greatest for the free drugs (colistin sulphate half maximal inhibitory concentration (IC50 = 0.026 mg/mL), polymyxin B (0.011 mg/mL)); slightly reduced by ester conjugation (OligoG–E–colistin (IC50 = 0.057 mg/mL), OligoG–E–polymyxin B (0.032 mg/mL)); and significantly reduced for the amide-linked conjugates (OligoG–A–colistin (IC50 = 0.242 mg/mL), OligoG–A–polymyxin B (0.299 mg/mL)). ELISA showed that the unmodified antibiotics induced greater TNFα release than the conjugates (Figure 1c). OligoG–E–polymyxin B caused the highest release of TNFα, compared to the other conjugates, but this was still lower than the unmodified drug.

**Figure 1.** In vitro cytotoxicity of OligoG–polymyxin conjugates in human kidney proximal tubule cells (HK-2) cells. Cell viability determined by 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) assay after 72 h incubation. (**a**) Colistin sulphate. (**b**) Polymyxin B. Data are presented as mean % of untreated control ± SEM (*n* = 18). (**c**) Tumour necrosis factor alpha (TNFα) release in HK-2 cells after incubation with OligoG–polymyxin conjugates for 72 h (±SEM; *n* = 6). (**d**) Half maximal inhibitory concentration (IC50) values (±SEM) and fold-change (MTT assay) of tested compounds in HK-2 cells. Significance is indicated by \*, where \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001, \*\*\*\* *p* < 0.0001 compared to colistin sulphate or polymyxin B controls. Abbreviations: A, amide; E, ester.

#### *3.4. Antimicrobial Activity of OligoG–Polymyxin Conjugates*

The effects of conjugation on antimicrobial efficacy against a range of Gram-negative pathogens varied between the conjugates and antibiotic (Table 2). Whilst ester-conjugation resulted in similar (≤2-fold differences) or decreased MIC values for OligoG–colistin and –polymyxin B conjugates, the amide-linked conjugates demonstrated increased MIC values. This effect was particularly evident for the polymyxin B conjugates, where MICs were increased by 4- to 32-fold. OligoG conjugation did not improve the bactericidal efficacy of colistin in colistin-resistant strains (Table 2). Both OligoG–polymyxin conjugates exhibited substantially improved selectivity for Gram-negative bacteria in comparison to mammalian cells, compared to unmodified colistin sulphate (1.7–4.7-fold) and polymyxin B (2.3–4.1-fold) (Table S4).

The antimicrobial activity of the conjugates was also assessed in the presence of alginate lyase or following pre-incubation with alginate lyase (Table S5), where no significant change was observed for either amide- or ester-bonded conjugates.

In contrast, when mucin was added to the broth, antimicrobial activity of OligoG–polymyxin conjugates decreased in a dose-dependent manner (Table S6). The presence of unconjugated OligoG with colistin sulphate or polymyxin B did not alter the antimicrobial activity of the free antibiotic.


Differences in antimicrobial activity of both, free- and OligoG-bound antibiotics were observed in AS medium compared to MH broth (Table S7). In most cases, using the checkerboard assay, an indifferent or additive effect was observed when azithromycin dihydrate was combined with OligoG–colistin or colistin sulphate (Table S8). Generally, OligoG conjugation did not alter the efficacy of the antibiotic combination, except for *A. baumannii* 7789, where the additive effect of colistin sulphate + azithromycin dihydrate (FICI = 1.14) became indifferent when azithromycin dihydrate was combined with OligoG–colistin conjugates (FICI >2.5). However, the combination of azithromycin dihydrate with OligoG–E–colistin resulted in a synergistic effect for the *E. coli* National Collection of Type Culture (NCTC) 10418 isolate (FICI = 0.46).

Bacterial growth curves of *P. aeruginosa* MDR 301 (Figure 2) showed that the OligoG–colistin conjugates delayed bacterial growth in a concentration-dependent manner (indicated by the longer lag-phase), and exponential growth was similarly slower compared to the untreated control. Growth inhibition (up to >48 h) was noted for the OligoG–colistin conjugates at ≥2 × MIC, while higher equivalent concentrations of colistin sulphate were required (≥8 × MIC) to achieve comparable efficacy. Neither amide- nor ester-linked conjugates had an effect on time to onset of bacterial growth. Both, OligoG–A–colistin and OligoG–E–colistin conjugates, at their MIC (1 and 0.5 μg/mL colistin base, respectively), demonstrated a delayed lag phase of >24 h and >18 h, respectively. Typically, colistin covalently conjugated to OligoG showed similar activity to the combined mixture of unconjugated colistin and OligoG at equivalent concentrations. Furthermore, neither OligoG nor Pronova alone had any significant effect in reducing bacterial growth, when an equivalent concentration to that contained in OligoG–colistin conjugates was used.

#### *3.5. Anti-Biofilm Activity of OligoG–Polymyxin Conjugates*

CLSM of biofilms grown in the presence of free and OligoG-bound colistin (≥MIC) showed a marked effect on biofilm formation (Figure 3). For example, the OligoG–E–colistin conjugate, at its MIC (1 μg/mL colistin base), caused obvious bacterial clumping, disruption of biofilm structure, and cell death (as noted by increased numbers of red cells). COMSTAT analysis revealed a significant reduction in biofilm thickness for all treatments (≥MIC) (*p* < 0.05) and biofilm roughness was significantly increased by OligoG–colistin conjugates (≥MIC) and colistin sulphate (MIC) treatments (*p* < 0.05). Both OligoG–colistin conjugates (2 × MIC) significantly reduced biofilm biomass compared to untreated control (*p* < 0.05), whereas no significant change was observed with colistin sulphate (up to 2 × MIC).

#### *3.6. Pharmacokinetic–Pharmacodynamic (PK–PD) Model*

In the PK-PD model (Figure 4), when antibiotic was placed in the IC, colistin diffused more rapidly than the OligoG–colistin conjugates (1 mg/mL in the OC was reached at 2.83 h (colistin) < 10.47 h (OligoG–E–colistin) < 17.35 h (OligoG–A–colistin)). Colistin sulphate, at MIC (0.25 μg/mL) and the OligoG–E–colistin conjugate at 2 × MIC (0.25 μg/mL colistin base) caused substantial bacterial killing after 4 h (<3 log10 CFU/mL) (Figure 4b). Both treatments caused a reduction in viable bacterial counts compared to the control (~5-fold lower) and the initial starting bacterial concentration (~2-fold lower). However, no antimicrobial effect was observed with the OligoG–A–colistin conjugate up to 2 × MIC.

**Figure 2.** Bacterial growth curves for *P. aeruginosa* MDR 301 (48 h) in the presence of the following antimicrobials. (**a**) OligoG–A–colistin conjugate. (**b**) OligoG–E–colistin conjugate. (**c**) Colistin sulphate. (**d**) Colistin sulphate plus OligoG control for amide-linked conjugate. (**e**) OligoG control for amide-linked conjugate. (**f**) PRONOVA control for amide-linked conjugate. (**g**) Colistin sulphate plus OligoG control for ester-linked conjugate. (**h**) OligoG control for ester-linked conjugate. (**i**) PRONOVA control for ester-linked conjugate. Controls were used at the equivalent concentrations used in the corresponding amide- or ester-linked conjugates (*n* = 3). Abbreviations: A, amide; E, ester.

**Figure 4.** Pharmacokinetic–pharmacodynamic (PK–PD) model to compare diffusion of free- and OligoG-conjugated colistin. (**a**) Change in colistin concentration in the outer compartment over 48 h (measured by BCA assay). Data are expressed as mean ± SD (*n* = 3). (**b**) Viability count of *A. baumannii* 7789 in a "time-to-kill" (TTK) model. Data represent mean colony forming units (CFU) ± SD (*n* = 3). Dotted line, lower limit of detection (102 CFU/mL). Abbreviations: A, amide; E, ester.

#### **4. Discussion**

#### *4.1. Rationale for Development of OligoG–Polymyxin Conjugates*

In recent years, interest in developing polymyxin derivatives to improve the therapeutic index or provide activity towards bacterial strains that are not currently susceptible to polymyxins has grown considerably [35]. Structural modifications, such as removal or substitution of the N-terminal acyl chain, reduction in the number of positive charges, polymer conjugation and the introduction of hydrophobic residues, have all been explored in an attempt to improve activity, reduce adverse side effects and elucidate structure–activity relationships [36,37]. These studies have demonstrated the critical importance of the amphipathicity of polymyxin molecules for their antimicrobial activity, which stems from the charged Dab residues and hydrophobic tail. This study compared the activity and toxicity of OligoG–polymyxin conjugates containing reversible or irreversible linking chemistries, to improve the therapeutic index of the polymyxins.

To form an amide bond between OligoG and polymyxin antibiotics, the carboxyl groups of the polymer were first activated by EDC in the presence of sulfo-NHS to create a stable amine-reactive intermediate [38]. The resultant amine-reactive sulfo-NHS ester was then bound to primary amines on the polymyxins. We hypothesised that amide-linked conjugates would rely on degradation of the OligoG backbone by alginate lyase. There are no known mammalian enzymes capable of degrading alginate, but bacterial alginate lyase from *K. pneumoniae* and *P. aeruginosa* has been found in cystic fibrosis patients' lungs [39] and could provide an opportunity for site-specific release of a therapeutic payload from an alginate conjugated drug. Although alginate lyases from these bacteria are generally considered to be mannuronate-specific, it has been shown that they may also demonstrate moderate to low activity towards guluronate [40]. Non-biodegradable polymers whose molecular weight is below the renal threshold (<40,000 g/mol) are expected to be readily excreted from the body [41], so the conjugates synthesised in the present study should be readily excreted by the kidney. Furthermore, sugar residues and/or linker groups that remain attached to the antibiotic Dab residues after polymer degradation, may result in reduced antimicrobial activity, as observed with amide-linked dextrin–colistin conjugates [18]. In parallel, ester-linked OligoG–polymyxin conjugates were formed using Steglich esterification. Here, the polymer carboxyl groups were activated by DCC, with DMAP as a catalyst [42]. The resultant *O*-acylisourea intermediate was then bound to hydroxyl groups on the polymyxins. Addition of DMAP as a catalyst compensates for hydroxyls being poorer nucleophiles

than amines (which can cause spontaneous rearrangement of the *O*-acylisourea intermediate into undesirable *N*-acylurea) by reacting with *O*-acylisourea to form an acyl pyridinium species. Reaction of DMAP with *O*-acylisourea forms an acyl pyridinium intermediate that is unable to form intramolecular by-products but can react with a hydroxyl group to form an ester bond [43]. Ester-linkage permits complete release of the native antibiotic at low or high pH, and by enzymatic activity and reactive oxygen species at sites of infection [44]. Moreover, release of intact OligoG at the target site would restore intrinsic antimicrobial activity of the polymer, which would further enhance its antibiotic efficacy in vivo.

#### *4.2. Physicochemical Characterisation of OligoG–Polymyxin Conjugates*

Covalent attachment of OligoG to colistin was confirmed using several methods. DOSY is an indirect method that can detect if the polymer and peptide are chemically linked as they will have the same diffusion coefficient if bound together in solution [45] while SEC detects an increase in size caused by an increase in molecular weight. In a control experiment using SEC-MALS, OligoG with 10% *w*/*w* added colistin yielded almost identical RI chromatograms as pure OligoG, confirming that the observed shift in elution profile for the OligoG–colistin conjugates was not simply due to electrostatic interactions between OligoG and free colistin. DOSY confirmed successful conjugation using both amide and ester linkers. Nevertheless, DOSY also detected unbound colistin in the OligoG–E–colistin sample and SEC-MALS analysis did not show an increased molecular weight compared to free OligoG. This may be caused by hydrolysis of the ester linkage during measurement (in D2O at 25 ◦C), rather than insufficient removal of unreacted material, highlighting the importance of using multiple analytical methods to characterise these complex molecules.

#### *4.3. Biological Characterisation of OligoG–Polymyxin Conjugates*

Polymyxin antibiotics have been reported to cause severe nephrotoxicity in up to 53.5% of patients [46] due to extensive reabsorption of the drug by renal tubular cells [47]. OligoG–polymyxin conjugates in the present study exhibited a marked decrease in in vitro cytotoxicity in kidney cells when compared to unmodified antibiotics. As expected, ester-linked polymyxin conjugates were considerably more cytotoxic towards HK-2 cells compared to amide-linked conjugates. Since the positively charged Dab residues are known to mediate polymyxin toxicity, and 2–4 of these primary amines were used for irreversible (amide-linked) conjugation with OligoG, this result was unsurprising.

Antimicrobial efficacy of all OligoG–polymyxin conjugates was comparable to that of the parent antibiotic. However, attachment of OligoG in the bi-functional molecule was unable to overcome colistin resistance. For colistin-sensitive strains, conjugate MIC values were below the Clinical and Laboratory Standards Institute [48] and European Committee on Antimicrobial Susceptibility Testing [49] susceptibility breakpoints (≤2 μg/mL) for polymyxins. Importantly, ester-linked conjugates showed full retention of the antimicrobial activity of the free drug, while the antimicrobial activity of the amide-linked conjugates was reduced by more than two-fold, presumably because of residual sugars and/or linker groups on the antibiotic amine groups. Although these studies did not test the stability of the conjugates in bacterial growth medium, it is likely that hydrolysis of the ester bond would occur during the MIC assay incubation. This may explain the smaller decrease in antimicrobial activity seen with ester-linked conjugates compared to amide-linked ones. Greater selectivity of OligoG–A–polymyxin conjugates for Gram-negative bacteria in comparison to mammalian cells suggests better tolerability and reduced side effects in vivo and substantially improved efficacy at clinically relevant concentrations compared to the free antibiotic. In addition, studies with alginate lyase suggested that either OligoG degradation is not necessary for antibiotic activity, or that alginate oligomers are broken down by bacterial enzymes. Nevertheless, compared to dextrin–colistin conjugates described in previous studies [16], alginate oligomer–conjugates were significantly more potent (more than five-fold change). This may be due to the larger molecular weight of dextrin causing

steric hindrance or the charge difference between the two polymers, but more likely, it can be attributed to the inherent biological activity of OligoG itself [20].

Sustained antibiotic release was demonstrated by slower bacterial growth in the presence of OligoG–colistin conjugates, which was dose-dependent. In this study, lower concentrations of OligoG–colistin conjugates, compared to free colistin, were required to inhibit bacterial growth which was sustained for up to 48 h. OligoG–E–colistin delayed the onset of bacterial growth for much longer than amide-linked conjugates, suggesting that, after systemic administration, OligoG–E–colistin conjugates might achieve better therapeutic activity in vivo. Similarly, when PEG was attached to colistin via a labile ester bond, sustained drug release led to equivalent or better antimicrobial activity against *A. baumannii* and *P. aeruginosa* isolates [17].

In a clinical setting, binding of colistin to sputum biomolecules (e.g., mucin) in the airways could negatively impact antibiotic effectiveness and availability. Indeed, Huang et al. [50] demonstrated >100-fold increase in MIC values of both, colistin and polymyxin B, when mucin was added to the bacterial culture medium. A four-fold increase in the MIC of colistin was also reported when the assay was conducted in AS medium instead of MH broth, thought to be caused by bacterial growth disruption, structural modifications of lipopolysaccharides or direct colistin–mucin interactions [24]. Although recent studies have demonstrated the ability of OligoG to bind mucin [51], conjugation of OligoG to colistin and polymyxin B did not affect the ability of the antibiotic to bind mucin or alter the effect of nutrient-deficient medium.

In practice, patients with severe infections of MDR pathogens are usually treated with combinations of two or more antibiotics to overcome or prevent drug resistance. When we combined OligoG–colistin conjugates with azithromycin dihydrate, an antibiotic that has previously shown enhanced efficacy in combination with OligoG [20], antimicrobial activity of the drug was enhanced, but only additively. Similarly, He et al. [52] reported additive effects in *P. aeruginosa* when they combined a low molecular weight alginate oligosaccharide (Mw < 10 kDa) with azithromycin, suggesting that the alginate oligosaccharide component of OligoG–colistin conjugates may be responsible for the additive effects observed in our study.

Chronic airway infections by *P. aeruginosa* affect more than 80% of CF patients and contribute to a progressive decline in lung function [53]. Marked disruption of *P. aeruginosa* biofilm formation was observed when they were grown in the presence of both OligoG–colistin conjugates, although only the ester-linked conjugate induced bacterial clumping (≥MIC) which might be associated with the higher cationic charge of colistin. This is in keeping with the findings of Powell et al. [23] who showed that OligoG, at concentrations ≥0.5%, caused *P. aeruginosa* aggregation, while higher concentrations (≥2%) caused significant disruption of bacterial biofilm formation and growth.

#### *4.4. PK–PD Modelling*

Drug TTK profiles and colistin release rate from amide- or ester-linked OligoG–colistin conjugates were investigated using an in vitro two-compartment PK–PD model. Predictably, diffusion of colistin, which was mirrored by a time-dependent increase in drug concentration in the OC, was substantially faster than the OligoG conjugates. When the ester-linked OligoG–colistin conjugate was contained in the IC, diffusion of colistin was more pronounced than when the amide-linked conjugate was tested, presumably due to the unstable nature of the ester bond. *A. baumannii* is an opportunistic pathogen that can causes serious infections often associated with multidrug resistant strains and has an 8.4–36.5% mortality rate [54]. In the present TTK study, although colistin sulphate at MIC (0.25 μg/mL colistin base) and the OligoG–E–colistin conjugate at 2 × MIC (0.25 μg/mL colistin base) exhibited rapid initial antimicrobial efficacy, marked bacterial re-growth was observed at 24 h. Previous studies have reported the impact of hetero-resistance of *A. baumannii* clinical isolates to colistin that allowed significant bacterial re-growth at 24 h at 32 × MIC [55] and 64 × MIC [56]. The reduction of viable bacterial counts by <3 log10 CFU/mL compared to the initial inoculum was indicative of bacteriostatic activity only. Similarly, bacteriostatic activity of colistin, at its MIC,

towards *A. baumannii* clinical isolates has been demonstrated, indicating a 2-fold decrease in CFU/mL at 4–6 h post-dose [57]. Importantly, previous studies saw significant bactericidal activity of colistin when carbapenem-resistant *A. baumannii* isolates were treated with higher concentrations of the antibiotic (≥4 × MIC) [58]. Observations in the present study support the clinical limitations of conventional colistin therapy, due to concentration-dependent nephrotoxicity, which may limit the optimal dosing and efficacy of the antibiotic.

The findings of this study indicate that the ester-linked OligoG–colistin conjugate could be a suitable alternative to conventional colistin, as it demonstrated equivalent antimicrobial effectiveness (0.25 μg/mL colistin base) but exhibited significantly lower cytotoxicity in human kidney cells. Following systemic administration of colistimethate sodium (Colomycin®; prodrug of colistin), the plasma colistin concentration at steady-state is 0.5–4 μg/mL [59]. Clinically, nephrotoxicity is an important limiting factor to colistin dosing, therefore, a plasma concentration of 2 μg/mL is desirable to target bacterial pathogens with MIC values ≤1 μg/mL [60]. To avoid acute kidney injury, a maximum plasma concentration of 2.42 μg/mL is recommended [61]. Yet, the clinical susceptibility breakpoint for colistin against *Acinetobacter* spp. is 2 μg/mL [48,49], which gives it a narrow therapeutic index in vivo. Due to the EPR effect, OligoG–E–colistin conjugates are expected to accumulate within infected tissues, so higher antibiotic concentrations (>0.25 μg/mL colistin base, equivalent to >2 × MIC) could theoretically be achieved much quicker than with the unmodified antibiotic. Sustained release over 48 h and concentration-dependent antibacterial efficacy of colistin has been achieved using dextrin–colistin conjugates [32]. In that study, colistin was covalently linked to dextrin through an amide bond, so "unmasking" of antibiotic relied on α-amylase-mediated degradation of the polymer. The fact that the OligoG–A–colistin conjugate did not show any antimicrobial effect in the PK–PD model in this study could be attributed to the absence of alginate lyase in the culture medium or the presence of residual saccharides attached to colistin which would not be present on antibiotic released from the ester-linked conjugates. Recently, it has been demonstrated that, even after complete amylase degradation of dextrin in amide-linked dextrin-colistin conjugates, the colistin molecule was still attached to at least one linker with varying lengths of glucose units [18]. These findings suggest that complete "unmasking" or release of colistin is a pre-requisite for reinstatement of antibiotic activity.

Importantly, the therapeutic benefits of OligoG–colistin conjugates might have been underestimated by the in vitro assays. Passive accumulation of conjugates at sites of in vivo bacterial infection due to the EPR effect, alongside the local reduced pH, reactive oxygen species and esterase activity as well as alginate lyase could all promote the controlled release of the drug from the polymer, and might further enhance the efficacy of the drug in vivo and thus, reduce the doses required to eradicate infection.

#### **5. Conclusions**

This study has established, for the first time, the potential therapeutic benefits of using OligoG conjugation to reduce antibiotic toxicity, while maintaining antimicrobial activity against MDR Gram-negative bacterial pathogens. These studies also demonstrate that complete detachment of the polymer from the bioactive compound is required to restore its full biological efficacy, with residual sugars shown to impede complete regeneration of activity. As OligoG has been shown to enhance the antimicrobial activity of macrolides, tetracyclines and β-lactams antibiotics, against a range of MDR Gram-negative bacteria [20], OligoG conjugation might also improve the pharmacokinetics of other toxic, water-insoluble or otherwise undeliverable drugs. Polymer conjugates like the OligoG–polymyxins offer a novel approach to repurpose "old" antibiotics into safer, less toxic bi-functional compounds to meet the increasingly urgent need for new antimicrobial therapies.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/1999-4923/12/11/1080/s1, Figure S1: Size exclusion chromatography with multi-angle light scattering detection (SEC-MALS) analysis of OligoG-conjugates (three different batches of OligoG–A–colistin (OAC) and one batch of OligoG–E–colistin (OEC)), showing overlaid refractive index chromatograms and corresponding Mw-time calibration lines. The injected

mass was 250 μg for all samples. Abbreviations: A, amide; E, ester, Figure S2: SEC-MALS analysis of OligoG in the absence and presence of 10% *w*/*w* colistin, showing that they do not form strong complexes since the elution profile is identical, Figure S3: Diffusion-ordered spectroscopy (DOSY) of (a–c) three different batches of OligoG–A–colistin conjugates and (d) one batch of OligoG–E–colistin conjugate. The assignment of the unique signals for OligoG and colistin is indicated at the top of each panel and the red lines indicate the average diffusion coefficients of the molecules, Figure S4: Drug release of OligoG–polymyxin conjugates in phosphate buffered saline (PBS) at pH 5, pH 7 or pH 7 containing alginate lyase (AlgL). (a) Content of free polymyxin and (b) change in molecular weight were determined by fast protein liquid chromatography (FPLC) and size exclusion chromatography with refractive index detection (SEC-RI), respectively, over 48 h incubation. Abbreviations: A, amide; E, ester, Table S1: Gram-negative bacterial isolates used for characterisation of OligoG–polymyxin conjugates, Table S2: Physicochemical characteristics and batch details of OligoG–polymyxin conjugates used in this study, Table S3: Weight and number average molecular weights of OligoG and OligoG–colistin conjugates, Table S4: Selectivity index (SI) values of OligoG–polymyxin conjugates against a range of Gram-negative bacterial pathogens, Table S5: Microbiological efficacy (MICs) of OligoG–colistin conjugates in the presence of alginate lyase in Mueller–Hinton (MH) broth or after pre-incubation with alginate lyase against Gram-negative bacterial pathogens, Table S6: Microbiological efficacy (MICs) of polymyxins and antibiotic conjugates in the absence and presence of mucin against Gram-negative bacterial pathogens, Table S7: Comparison of the effect of growth medium (AS medium and MH broth) on antimicrobial activity (MIC determinations) of polymyxins and antibiotic conjugates, Table S8: Fractional inhibitory concentration index (FICI) values of OligoG–colistin conjugates or colistin in combination with azithromycin dihydrate, Methods: NMR spectroscopy.

**Author Contributions:** Conceptualization, J.S., E.L.F., D.W.T. and K.E.H.; Investigation, J.S., L.C.P. (participated in the confocal imaging experiments and analysis), O.A.A. and F.L.A. (SEC-MALS and DOSY experiments and analysis); Formal analysis, J.S., E.L.F., D.W.T., K.E.H.; Writing—Review and Editing, J.S., E.L.F., D.W.T., K.E.H. and P.D.R. All authors read and approved the final manuscript.

**Funding:** This work was supported by funding from the Research Council of Norway (228542/O30, 281920 and 226244), AlgiPharma AS, Sandvika, Norway and UK Medical Research Council (MR/N023633/1).

**Acknowledgments:** We thank Timothy Walsh (Department of Medical Microbiology and Infectious Disease, Cardiff University, UK) for the colistin resistant bacterial isolates. We thank Anne Tøndervik and Håvard Sletta for intellectual discussions about the research and Alexander Åstrand (AlgiPharma AS) for helpful comments on the manuscript.

**Conflicts of Interest:** This work was partly supported by funding from AlgiPharma AS, Sandvika, Norway who also provided the alginates used in the study. The authors (D.W.T. and K.E.H.) declare previous research funding from AlgiPharma AS. P.D.R. is CSO at AlgiPharma AS.

#### **References**


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## *Article* **Mixed Pluronic—Cremophor Polymeric Micelles as Nanocarriers for Poorly Soluble Antibiotics—The Influence on the Antibacterial Activity**

**Maria Antonia Tănase 1, Adina Raducan 1, Petru¸ta Oancea 1, Lia Mara Di¸tu 2, Miruna Stan 3, Cristian Petcu 4,\*, Cristina Scomoro¸scenco 4, Claudia Mihaela Ninciuleanu 4, Cristina Lavinia Nistor <sup>4</sup> and Ludmila Otilia Cinteza 1,\***


**Abstract:** In this work, novel polymeric mixed micelles from Pluronic F127 and Cremophor EL were investigated as drug delivery systems for Norfloxacin as model antibiotic drug. The optimal molar ratio of surfactants was determined, in order to decrease critical micellar concentration (CMC) and prepare carriers with minimal surfactant concentrations. The particle size, zeta potential, and encapsulation efficiency were determined for both pure and mixed micelles with selected composition. In vitro release kinetics of Norfloxacin from micelles show that the composition of surfactant mixture generates tunable extended release. The mixed micelles exhibit good biocompatibility against normal fibroblasts MRC-5 cells, while some cytotoxicity was found in all micellar systems at high concentrations. The influence of the surfactant components in the carrier on the antibacterial properties of Norfloxacin was investigated. The drug loaded mixed micellar formulation exhibit good activity against clinical isolated strains, compared with the CLSI recommended standard strains (*Staphylococcus aureus* ATCC 25923, *Enterococcus faecalis* ATCC 29213, *Pseudomonas aeruginosa* ATCC 27853, *Escherichia coli* ATCC 25922). *P. aeruginosa* 5399 clinical strain shows low sensitivity to Norfloxacin in all tested micelle systems. The results suggest that Cremophor EL-Pluronic F127 mixed micelles can be considered as novel controlled release delivery systems for hydrophobic antimicrobial drugs.

**Keywords:** mixed polymeric micelles; drug delivery; antibiotics; Pluronic F127

#### **1. Introduction**

Infectious disease treatments continue to impose the extensive use of antibiotics. Despite the remarkable advances made in the last century in the synthesis of new drugs with antimicrobial activity, there are many deficiencies, such as high toxicity, low solubility, reduced bioavailability, and inadequate release profile for both new and old antibiotics successfully used in the present therapies. The main issue, however, remains the multidrug resistance, which produce a huge burden on the global health system.

To overcome these drawbacks, in the last decades, nanosized drug delivery systems have been generate increasing interest of the scientific community, due to their unique physicochemical properties. The advantages of these nanoparticulate carriers in antimicrobial formulations include targeted delivery to the infection site, improved cellular

**Citation:** T ˘anase, M.A.; Raducan, A.; Oancea, P.; Di¸tu, L.M.; Stan, M.; Petcu, C.; Scomoro¸scenco, C.; Ninciuleanu, C.M.; Nistor, C.L.; Cinteza, L.O. Mixed Pluronic—Cremophor Polymeric Micelles as Nanocarriers for Poorly Soluble Antibiotics—The Influence on the Antibacterial Activity. *Pharmaceutics* **2021**, *13*, 435. https://doi.org/10.3390/pharmaceutics 13040435

Academic Editor: Umile Gianfranco Spizzirri

Received: 14 February 2021 Accepted: 22 March 2021 Published: 24 March 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

internalization and drug stability, higher solubility, and sustained drug release [1]. The quinolone and their derivatives fluoroquinolone antibiotics are the most efficient class of topoisomerase inhibitors, used to treat bacterial infections caused by both Gram-positive and Gram-negative bacteria [2]. Their mechanism of action consists in the inhibition of topoisomerase enzymes (DNA gyrase implicated in genome replication and transcription), which inhibits the relaxation of supercoiled DNA and promotes the breakage of double stranded DNA [3]. Their broad antimicrobial spectrum is due to their ability to cross bacterial cell wall and cytoplasmic membranes via passive diffusion mechanism, being is strongly dependent on the lipid composition [4]. Consequently, the antibacterial activity of fluoroquinolones appears to result from the combination of efficient cellular membrane penetration and DNA gyrase inhibiting activity. Norfloxacin (1-ethyl-6-fluoro-l,4-dihydro-4-oxo-7-(1-piperazinyl)-3-quinoline carboxylic acid) is one of the most used antibiotics from the class of fluoroquinolones [5]. Due to its complex structure, it shows superior antibacterial activity against both gram positive and gram-negative bacteria, and is used to treat a large variety of respiratory or urinary tract infections.

Norfloxacin is considered as a poorly soluble drug, despite of the experimental partition coefficient log P that is reported in literature ranging from −0.43 to −1.52, quite different from the value obtained from theoretical calculation (−0.92 to 1.44) [6]. However, the large value of negative thermodynamic potential of solvation in the apolar solvent explains the poor bioavailability (less than 40%) and short half-time (3–4 h) in serum [7]. Various drug delivery systems were proposed to prolong the release, and good results have been obtained. Encapsulation in polymeric material guar gum, sodium carboxymethyl cellulose, and hydroxypropyl cellulose lead to an extended the Norfloxacin release over a period of 7–12 h [8]. A sustained release of the drug was also reported using Norfloxacin loaded proniosomes and lipid polymer hybrid nanoparticles [9].

Taking into account the advantages of different nanostructured vectors, various drug delivery systems for Norfloxacin (NFLX) have been proposed to increase its solubility, stability to chemical and photodegradation, and to improve the therapeutic benefits. For example, Ahmad et al. [10] are proposing a series of liposomal preparations of Norfloxacin with variable concentrations of phosphatidylcholine, based on the charge transfer complex between drug and phosphatidylcholine molecules. In another study, Norfloxacin-loaded nanosponges based on cyclodextrin to maximize oral absorption were proposed [11]. Norfloxacin-stearic acid solid-lipid nanoparticles were also successfully used as an oral delivery formulation [12]. Another hybrid drug delivery system containing Norfloxacin loaded into TiO2 nanoparticles followed by encapsulation onto poly lactic acid was found to be a suitable carrier with high antibacterial activity [13].

Nanocarriers as delivery systems for antibiotics enhance drug solubility, modulate drug release characteristics, and are also valuable tools in fighting antibiotic resistance [14]. Advanced drug delivery systems are studied for the encapsulation of norfloxacin [15] and other quinolones [16], but very rare micellar systems are investigated [17].

Micelles dispersions, from classic surfactants or polymeric surfactants, are currently used in many pharmaceutical products with various applications, from infusible solutions with chemotherapeutics or antibiotics to ocular formulations. Most of the pharmaceutical products contains as solubilizers nonionic surfactants, such as Tweens or Cremophors. Cremophor EL is FDA-approved nonionic emulsifier, used as a solubilizing agent for many years [18], produced by reacting castor oil with ethylene oxide. Therefore, it contains a mixture of unmodified castor oil and a large variety of polyethylene glycols, polyethoxylated glycerols, polyethoxylated fatty acids, and mono-, di-, and tri-esters of glycerol that are polyethoxylated to different degrees [19]. Due to its excellent emulsifying capacity, Cremophor is used for solubilization, protection, and delivery of different lipophilic active pharmaceutical ingredients (API). Numerous studies highlight Cremophor toxicity, namely anaphylactic hypersensitivity reactions, allergic shock, lipoprotein patterns and hyperlipidemia, neurotoxicity and hypotension, also at low concentration [20]. Also, some papers

indicate that Cremophor can modify the toxicity profile of some active pharmaceutical ingredients [20].

Polymeric surfactants are considered a safer and more convenient alternative to classic surfactants, due to their low CMC values, good biocompatibility profile, and high drug entrapment efficiency since they possess large hydrophobic inner core. Among them, the most investigated class are polyethylene oxide-polypropylene oxide block copolymers (Pluronics or Poloxamers). These group of surfactants are FDA approved and used in various nanocarriers, in micelles, gels, and polymer stabilized nanoemulsions, to solubilize and improve the bioavailability of the poorly water-soluble active pharmaceutical ingredients. Their specific tri-block copolymer structure offers advantage regarding the modification of the release and are widely used as a carrier for controlled drug delivery [21]. Another significant advantage is that aqueous solutions of Pluronics in the presence of acids, alkalis, and metal ions are very stable [22]. Also, these polymeric micelles are more investigated because they do not show demicellization (micellar aggregate breaking) upon dilution in the presence of biological fluids such as blood and tearing.

Micellar carriers that contain Pluronic polymers exhibit good biocompatibility over a large concentration range, characteristic confirmed in many studies [23].

Most of the inconvenient of micellar systems as drug delivery systems are related to the intrinsic toxicity due to the large content of surfactant required to produce micellar aggregates even after dilution suffered after administration. In this respect, a mixture of surfactants is investigated as a possible solution to prepare mixed micelles with superior stability. When classic surfactants are added to Pluronic micelles depending on the molecular interactions taking place in the system, size and morphologies of the formed aggregates are modified [24], significantly modifying the encapsulation and release capacity of the drug. Forming mixed micelles with Pluronics is especially used to decrease the critical micellar concentration of the systems. Extensive research is available on the synergistic effects in the micellization process in mixtures containing various ionic surfactant and polymeric surfactants, while very few studies report synergism in mixed micelles of Pluronics and nonionic surfactants used in drug delivery systems formulation [25,26].

Mixed micelles are used especially to enhance the drug encapsulation and delivery parameters and have great potential as efficient drug carrier [27]. Beyond the reduction of CMC value, mixed micelles show other synergistic properties, such as increased drug loading capacity and micelle stability, higher than of the individual components. According to some reports mixed micelles that include Pluronic F127 exhibit higher solubilization capacity compared to pure F127 micelles [21]. In a study is reported a polymeric mixed micellar formulation at 1:1 Poloxamer 407/Pluronic P123 ratio, that exhibits a slower and controlled release of the drug, in contrast to the pure polymer micellar system. This behavior minimized the adverse effects associated with exceeding the safe concentration of the drug [28].

Although polymeric micelles are known to be very efficient drug carriers, little attention has been paid in using them as nanosized formulations for antibiotics. In a study carried by Khanal [29], a drug delivery system was developed by encapsulate the anionic drug cloxacillin sodium in a polyvinyl pyridine block of polystyrene-b-2-vinyl pyridine-bethylene oxide in order to investigate the possibility of micelles being a suitable antibiotic delivery system. Polymeric micelles were also proposed as carrier to ensure the drug transport across the blood brain barrier (BBB). Micelles prepared from cholesterol-conjugated PEG and anchored with a transactivator of transcription (TAT) peptide (TAT-PEG-b-Col) were prepared to encapsulate Ciprofloxacin and prove sustained antibacterial activity against *B. subtilis* and *E. coli*. The in vivo study evidenced that the polymeric micelles pass the BBB [30].

The aim of the present study was to prepare mixed micellar systems containing a well-known nonionic surfactant, i.e., Cremophor EL and Pluronic F127 to encapsulate a poorly soluble antibiotic Norfloxacin. The self-assembling properties in mixed surfactant systems were systematic investigated to prove the possibility of synergistic effects in micelle forming behavior. The optimization of the composition ensures the minimum surfactant amount in formulation and the use of reduced quantity of Cremophor EL, to decrease the side effects associated with it. The antibacterial activity of Norfloxacin loaded mixed micelles was evaluated, in order to consider these novel colloidal vectors as possible nanocarriers for poorly soluble antibiotics.

#### **2. Materials and Methods**

#### *2.1. Materials*

Micelle forming polymeric surfactants Pluronic® F127 (BioReagent MW = 12,600 g/mol), Cremophor EL (Millipore, Burlington, MA) and model drug Norfloxacin (≥98% TLC) were purchased from Sigma-Aldrich (Merck Group, Darmstadt, Germany). Pyrene (99% purity) and the solvents absolute ethanol (99.9%), dimethyl sulfoxide (DMSO), benzene (99% purity), chloroform (99% purity), phosphate Buffer Solution (99% purity), and hydrochloric acid (36.5 g/mol, 99% purity) were also obtained from Sigma-Aldrich (Merck Group, Darmstadt, Germany). All used reagents and chemicals were used as received without further modifications.

#### *2.2. Preparation of Micelles*

A simple thin-film rehydration method was used to prepare micelles of Pluronic F127 and Cremophor EL. The procedure is schematically presented in the Figure 1.

**Figure 1.** Schematic representation of the obtaining procedure for the Norfloacin loaded Pluronic F127—Cremophor EL mixed micelles.

Suitable amounts of surfactants or surfactant mixtures were dissolved in ethanol in a round-bottom flask. The vial was attached to a rotatory evaporator (Rotavapor-R-300®, Buchi Labortechnik AG, Flawil, Switzerland) and heated at 45 ◦C under vacuum to produce a thin film of micelle forming material deposited on the walls. After the total removal of the solvent, the required volume of distilled water was added to rehydrate the film under moderate magnetic stirring at 45 ◦C, for 30 min. The as prepared micellar dispersion was further filtered through sterile syringe filter Minisart® 0.2 μm (Sartorius, Gottingen, Germany). For the mixed micelles, the two surfactants were added in ethanol solvent to obtain molar ratio 0.2, 0.4, 0.6 and 0.8 in the final aqueous dispersions.

The drug loaded pure and mixed micelles were prepared following the same procedure and Norfloxacin was dissolved together with the polymeric surfactants in alcohol. The dissolution of the Norfloxacin in the deposited thin film before the rehydration step ensure the maximum solubilization of the drug inside the micelle core.

#### *2.3. Characterization of Mixed Micelles*

*CMC* values of the pure and mixed surfactant systems were evaluated using the fluorescence method with pyrene as fluorescence probe [31]. Briefly, the fluorescence spectra of pyrene incorporated into the studied micelles (pure Cremophor EL, Pluronic F127 or mixed in various molar ratio) were recorded. A sharp variation of the slope in the graphic representation of the I3/I1 ratio (first and third vibronic peaks) indicate the CMC. For the sample preparation, pyrene was dissolved in acetone, a required volume was added in a graduate flask and the solvent was removed. Then, the surfactant solution was added to pyrene to reach a concentration of 1 × <sup>10</sup>−<sup>6</sup> M. The concentrations of the analyzed solutions ranked from 2.5 × <sup>10</sup>−<sup>4</sup> M to 2.5 × <sup>10</sup>−<sup>6</sup> M for all surfactants and their mixtures. The spectrofluorimetric measurements were performed using a Jasco FP8200 instrument (JASCO Corporation, Tokyo, Japan). The excitation wavelength for pyrene was 334 nm and its emission was recorded between 350 and 600nm. The intensity ratio between the first peak (373 nm) and the third peak (384 nm) was plotted against the concentration of the analyzed solutions and analyzed for the calculation.

Particle Size, Polydispersity Index (PI), and Zeta Potential: The pure and mixed polymeric micelles size and size distribution were measured using the dynamic light scattering (DLS) method. The data was analyzed by number and intensity weighted distributions. The zeta potential was calculated by Laser Doppler Velocimetry (LDV). The measurements were performed on a Nano ZS ZEN3600 Zetasizer (Malvern Instruments Ltd., Malvern, UK) equipment. The measurements were carried out on pristine and diluted samples, and the dilution was made with distilled water or PBS. In order to evidence the stability against aggregation and detect the presence of precipitated drug nanocrystals the samples were measured before and after filtration.

The micropolarity of the micelles was evaluated from the value of the I3/I1 ratio calculated for pyrene spectra in micellar solutions, compared to the values obtained in water and nonpolar solvent as reference.

The stability of micellar systems against dilution was evaluated by changes in size and size distribution when sample were diluted 10 time in either water or PBS. The resistance of micellar aggregates at different pH was also investigated by DLS measurements in surfactant solutions at pH = 4, pH = 6 and pH = 7.4 values.

The physicochemical stability of drug-containing micelles was evaluated by measuring the variation of the size and size distribution on samples stored at room temperature over four weeks.

#### *2.4. Drug-Surfactant Interaction*

The interaction of Norfloxacin with micelle forming surfactants was investigated using FTIR. The measurements were performed as follows. The spectra were recorded with a Tensor 37 Bruker equipment (Woodstock, NY, USA), using 32 scans with 4 cm−<sup>1</sup> resolution in the 4000−400 cm−<sup>1</sup> spectral range.

The sample pellets were prepared by adding pure components (NFLX, Pluronic or Cremophor) and drug loaded micellar solution in KBr powder, further subjected to extensive drying procedure to remove the solvent.

#### *2.5. Drug Solubility and Entrapment Efficiency*

For the measurement of the maximum solubility of Norfloxacin in surfactant solutions pure and mixed micelles were prepared with Cremophor EL, Pluronic F127 and binary mixture of Cremophor and Pluronic F127 with the molar ratio α = 0.2, all samples at concentrations five time their CMC value. Excess amounts (2 mg) of Norfloxacin were added in flasks and dissolved with 1 mL of ethanol. The solvent was removed in a rotaevaporator and the drug was further dissolved in 1 mL of each micellar solution. The drug—micellar systems were left to equilibrate for 12 h and the obtained dispersions were centrifuged and filtered using 0.22 μm regenerated cellulose syringe filters.

The drug amount was quantified using fluorescence method adapted in our laboratory from literature [32]. Briefly, fluorescence spectra were recorded in acidic solution of 0.1 N HCl, with λex = 330 nm and λem = 450 nm. Concentration of Norfloxacin in micellar systems tested for calibration was: 100 μg/mL, 50 μg/mL, 25 μg/mL, 12.5 μg/mL, 6.25 μg/mL and 3.125 μg/mL. Calibration curves were obtained by plotting the fluorescence intensity at 450 nm against the concentration of Norfloxacine.

Good linearity was obtained for the concentration range 50–3.125 μg/mL for all micellar systems with R2 = 0.9961 for Cremophor micelles, R<sup>2</sup> = 0.9903. For Pluronic F127 micelles and R2 = 0.9908 for the binary mixture of Cremophor and Pluronic F127 with the molar ratio, α = 0.2.

The encapsulation efficiency (EE%) was calculated as the weight ratio of encapsulated drug NFLX to the drug in feed at drug-loaded micelles:

EE (%) = Experimental drug loading/Theoretical drug loading × 100

#### *2.6. In Vitro Drug Release*

The release of Norfloxacin from pure micelles of Cremophor and Pluronic and mixed micelles of binary mixture of Cremophor and Pluronic at selected molar ratio α = 0.2 was studied using the dialysis bag method under physiological conditions. A volume of 2 mL of Norfloxacin encapsulated (100μg/mL) in Pluronic F127 (1.5 × <sup>10</sup>−<sup>3</sup> M), Cremophor EL (8 × <sup>10</sup>−<sup>4</sup> M) and a binary mixture of Cremophor EL and Pluronic F127 with the molar ratio, <sup>α</sup> = 0.2 (1 × <sup>10</sup>−<sup>3</sup> M) were added to the pre-swelled dialysis bag with two ends sealed with plastic sealing clips. The systems were weighted before and after adding the NFLX micellar solution. Each bag was placed in PBS release medium at room temperature, under constant stirring (50 rpm). Aliquotes of 0.5 mL were withdrawn from the release medium at predetermined time intervals (15 min, 30 min, 45 min, 1–8 h, 24 h) and replaced with the same amount of fresh medium.

All aliquots were diluted 1:10 (*v*/*v*) with 0.1N HCl and quantified by means of fluorescence spectrophotometry. The spectrofluorimetric measurements were performed using a Jasco FP8200. The excitation wavelength was 330 nm, and its emission was recorded between 350–600 nm. The intensity of the peak at 450 nm was recorded and further used to calculate the cumulative % of drug release.

The influence of the temperature and pH of the media on drug release was tested and the experiment were performed at 25 ◦C and 37 ◦C, in PBS buffer with pH values of 7.4, 6 and 4.

#### *2.7. Cells Viability Assay*

*Cell culture:* Human lung fibroblasts MRC-5 (ATCC CCL-171) were grown in complete Eagle's minimal essential medium (Invitrogen, USA) containing 10% fetal bovine serum (Gibco, Carlsbad, CA, USA) at 37 ◦C in a humidified atmosphere with 5% CO2. The cells were seeded at a cell density of 5 × 104 cells/cm2 and left to adhere for 24 h. Then, the fibroblasts were incubated for the next 24 or 48 h with different concentrations of surfactants in the range 1 × <sup>10</sup><sup>−</sup>6–1 × <sup>10</sup>−<sup>3</sup> M, which were previously sterilized by filtration with 0.2 <sup>μ</sup><sup>m</sup> pore size filter membrane. Untreated cells were used as control for all in vitro experiments.

MTT assay: The cellular viability was measured using the 3-(4,5-dimethylthiazol-2 yl)-2,5-diphenyltetrazolium bromide (MTT; Sigma-Aldrich, St. Louis, USA) assay. After 24 h of incubation, the culture medium was removed, and the cells were incubated with 1 mg/mL MTT for 2 h at 37 ◦C. The purple formazan crystals formed in the viable cells

were dissolved with 2-propanol (Sigma-Aldrich, St. Louis, USA) and the absorbance was measured at 595 nm using a microplate reader (Flex Station, Molecular Devices).

Griess assay: The concentration of nitric oxide (NO) in the collected culture medium after the 24 h of incubation was performed with the Griess reagent, a stoichiometric solution (*v*/*v*) of 0.1% naphthylethylendiamine dihydrochloride and 1% sulphanilamide in 5% H3PO4). Increased levels of NO are related with cytotoxic effects as this molecule relates to inflammation and apoptosis. The absorbance of mix formed by equal volumes of medium supernatants and Griess reagent was read at 550 nm using the FlexStation 3 microplate reader and the NO concentration was calculated from the NaNO2 standard curve.

Statistical analysis: The in vitro assays were performed in triplicates and the results were presented as mean ± standard deviation (SD) of three independent experiments. The statistical significance was analyzed by Student *t*-test. A value of *p* less than 0.05 was considered significant.

#### *2.8. Antibacterial Activity*

The antimicrobial assays were performed using standard and clinical bacterial strains that were included in the microbial collection of University of Bucharest, Faculty of Biology, Microbiology Department: *Staphylococcus aureus* ATCC 25923 and *Staphylococcus aureus* MRSA clinical strain, *Enterococcus faecalis* ATCC 29213 and *Enterococcus faecalis* VRE clinical strain, *Pseudomonas aeruginosa* ATCC 27853 and *Pseudomonas aeruginosa* 5399 clinical strain, and *Escherichia coli* ATCC 25922 and *Escherichia coli* ESBL 135 clinical strain. To perform the experiment, two successive passages were made by passing the microbial strains on nutritious agar medium and incubating for 24 h, at 37 ◦C.

The qualitative screening of the anti-microbial properties was performed by an adapted spot diffusion method, according with CLSI standard (Clinical Laboratory Standard Institute, 2021). Bacterial suspensions of 1.5 × 108 CFU/mL (corresponding with 0.5 McFarland standard density) obtained from 24 h microbial cultures developed on Muller Hinton agar (MHA) were used in the experiments. Petri dishes with MHA were seeded with microbial inoculums and an amount of 10 μL solution of each sample was spotted, the calculated concentration of norfloxacin being 20 μg/mL. The standard disks with 30 μg of norfloxacin were used as control for the strain's sensitivity. The plates were left at room temperature to ensure the equal diffusion of the compound in the medium and then incubated at 37 ◦C for 24 h. Sensitivity was evaluated by measuring the diameters of the inhibition zones that appeared around the spot and expressed in mm.

For establishing the MIC (minimum inhibitory concentration) values of the obtained compounds we utilized a serial microdilution method performed in nutritive broth. The sterile broth was added in sterile 96 well plates and binary dilutions of each tested compound were performed in a final volume of 150 μL, starting with 20 μg/mL concentration calculated for Norfloxacin, in the first well. Further, 15 μL of microbial suspension adjusted to 1.5 × <sup>10</sup><sup>7</sup> CFU/mL, were added in each well. The MIC values were established by spectrophotometric measurement (absorbance reading at 600 nm using BIOTEK SYNERGY-HTX ELISA multi-mode reader). Each experiment was performed in triplicate and repeated on at least three separate occasions.

Statistical analysis: For biological tests, significant differences between the means of triplicate experiments and the control were determined by using one-way ANOVA statistical analysis (significance difference was noted as \* for *p* < 0.05, and \*\* for *p* < 0.01). All data are presented as mean values ± the standard deviations (SD).

#### **3. Results and Discussion**

*3.1. Mixed Micelles Preparation and Non-Ideal Behavior in Mixed Pluronic F127-Cremophor EL Aqueous Solutions*

In this study, a novel carrier from mixtures of two nonionic polymeric surfactants Cremophor EL and Pluronic F127 is proposed, in order to obtain a controlled drug delivery for Norfloxacin with extended release and higher permeability through cell membranes.

The adequate composition of the mixed polymeric micelles to ensure the minimum content of surfactants and a low value of CMC was evaluated from the non-ideal behavior of surfactant mixture, using Rubingh model. The self-assembling properties of the aqueous solution of Cremophor EL and Pluronic F127 have been studied and CMC values were calculated from the variation of pyrene fluorescence spectra, i.e, variation of the I3/I1 ratio with concentration of surfactant, as described in the previous section. The point where two linear fitting curves in premicellar (low concentration) and micellar region intersect is considered as CMC value. To evaluate the ideal or non-ideal behavior in mixed micelles, CMC theoretical values were calculated from the Clint Equation (1):

$$\frac{1}{C^\*} - \frac{a\_1}{C\_1} + \frac{1 - a\_1}{C\_2} \tag{1}$$

where *C*\* represents the CMC of the binary mixture of Surfactant 1 and 2, α is the mole fraction of surfactant 1 in the mixed solution and *C*<sup>1</sup> and *C*<sup>2</sup> are the individual CMCs of Surfactant 1 and Surfactant 2.

In order to calculate the intensity of interaction between the two surfactants from the binary mixture, one can use a parameter, β, calculated using Equation (2), from the model proposed by Rubingh [33]:

$$\beta\_{12} = \frac{\ln \frac{\alpha\_1 C^\*}{x\_1 C\_1}}{\left(1 - x\_1\right)^2} \tag{2}$$

where *α*<sup>1</sup> is the mole fraction of surfactant (Cremophor EL) in the mixed micellar solution, *C*\* is CMC of mixed micelles, *C*<sup>1</sup> is the CMC of Surfactant 1 and *x*<sup>1</sup> is the micellar mole fraction of Surfactant 1.

The micelle mole fraction *x*<sup>1</sup> can be calculated by iteratively solving Equation (3):

$$\frac{\mathbf{x}\_2^1 \cdot \ln \frac{a\_1 C^\*}{x\_1 C\_1}}{(1 - x\_1)^2 \cdot \ln \frac{(1 - a\_1) C^\*}{(1 - x\_1) C\_1}} = 1\tag{3}$$

The value of parameter of interaction β quantifies the interactions between the molecules of surfactant. When attractive forces between the two surfactants are present, the value of this parameter is negative, meaning that synergism is present in the mixed micelles. Positive values for parameter β indicate an antagonistic effect, whereas in the case of β = 0 the mixed micelles formation is considered ideal. The larger the value of β (both positive or negative) the stronger the interaction (repulsion or attraction) between the two surfactants. The determination of β values allow to select mixture where synergistic behavior is present, leading to the formation of micellar aggregates at lower concentration that corresponding pure surfactants. More stable micellar systems with low surfactant content could be obtained, with a certain advantage for pharmaceutical formulation.

In Table 1, the theoretical and experimental values of CMC, together with the molar fraction in mixed micelles and parameter of interaction β are summarized for the mixture containing Cremophor EL and Pluronic F127 in various molar ratio (α is expressed relative to Cremophor EL as Surfactant 1 and Pluronic F127 as Surfactant 2).

As it is expected for a mixture with nonionic surfactants, a moderate synergistic effect is observed [25], with low values of interaction parameter, ranging from −0.22 to −1.54.

Negatives values for β were obtained for all molar fractions of 0.2, 0.6 and 0.8, but the higher value is recorded for the molar fraction 0.2, where the higher synergistic effect appears. For further experiments mixture with molar ratio α = 0.2 was selected since contain the smallest amount of Cremophor EL and the CMC value for the mixture is very low.


**Table 1.** The micellization parameters in binary mixtures Cremophor EL and Pluronic F127 in various molar ratio obtained from the Rubingh model.

\* using Equation (1).

#### *3.2. Micelles Characterization*

The size and size distribution of the micelles formed in Cremophor EL, Pluronic F127, and their mixture in aqueous solution was evaluated from DLS measurements (Figure 2).

**Figure 2.** Size and size distribution of the micelles: (**a**) Cremophor EL, (**b**) Pluronic F127, (**c**) Mixed Cremophor EL-Pluronic F127 micellar system, (**d**) Norfloxacin loaded mixed Cremophor EL-Pluronic F127 micellar system.

At 25 ◦C the DLS diagram for 6 × <sup>10</sup>−<sup>4</sup> aqueous solution of Cremophor EL shows a single population of scattering units, the surfactant micelles, with an average diameter of 12.4 ± 2.4 nm and zeta potential −0.98 ± 0.4 mV. The polydispersity index PdI is 0.157 indicating high monodispersity of the sample. In contrast, the solution of Pluronic F127 exhibits trimodal distribution, with three signals in intensity mode representation. The main diameter of the empty micelles is 31.91 ± 4.4 nm, within the range of reported values [22]. The strong intensity signal at 6.88 nm is due to the presence of the polymeric macromolecules unassociated, in equilibrium with the micellar aggregates and the signal around 354 nm is probably due to some larger micelle—micelle aggregates, which have been reported in other papers [34]. The polydispersity index of 0.669 is consistent with a trimodal distribution. The sizes of the mixed micelles range from 14.90 nm to 27.54 nm with the increase of Pluronic F127 in composition, and the presence of Pluronic non associated polymeric chain is no longer evidenced. Also, the large aggregates disappear, that confirm the increase in micellization tendency in binary mixtures as it is observed from the synergistic behavior discussed in the previous section. The size of mixed micelles is

lower that is expected from an ideal mixing, and it is also consistent with the data obtained from Rubing model (Table 1), where the molar fraction of Cremophor EL in mixed micelle is always higher than the one in mixed solution.

The solubilization of the drug in mixed micelle in a concentration of 100 μg/mL produced a slightly increase in the micellar size, from 27.54 ± 8.4 nm for empty Cremophor EL—Pluronic F127 mixed micelles F127 at α = 0.2 molar ratio to 28.42 ± 7.9 nm for Norfloxacin loaded micelles. The solubilization of hydrophobic drugs inside the inner core of the micelles results in most of the case in more obvious increase of the loaded micelles [35]. Sometimes the encapsulation of the more complex molecule, such as Norfloxacin is simultaneously accompanied by a dehydration of the polymeric aggregate that produce a thinning of the hydrophilic polyoxiethylene corona [36], thus the modification of the micelle size is less evidenced.

The drug release from micelles do not produce observable changes in the size of mixed aggregates (average size before the experiment 26.42 ± 0.95 nm compared to samples after release experiment 26.90 ± 2.53 nm).

The micropolarity of the micelles was determined by using Pyrene as fluorescence probe, as recommended in literature [37]. The intensity of the vibronic bands in the fluorescence spectrum of monomeric Pyrene are reported to be very strong dependent with de polarity of the microenvironment. The ratio I3/I1 (where I1 is the first maximum emission peak at 372 nm and I3 the third one, at 384 nm) are considered as micropolarity index [38].

The I3/I1 value obtained from pyrene in water is very high, in the range 1.79–1.82, while smaller values in the range of 0.80–0.90 were determined for fluorescent probe inside micelles of various surfactants, indicating that pyrene molecules are located in a less polar environment, i.e., the hydrocarbonate core of the micellar aggregate.

For the evaluation of the microenvironmental polarity in the micelles, the ratio I3/I1 (first to third vibronic peaks) at the plateau region in the I3/I1 versus surfactant concentration curve was used, which is consistent with micellar domain.

From the spectra of pyridine recorded in pure Cremophor, pure Pluronic and mixed Cremophor-Pluronic micelles, the I3/I1 values were found 0.89, 0.96, and 0.90, respectively. The value of I3/I1 ratio of 0.89 for the Cremophor micellar system is consistent with the chemical structure of surfactant, which can produce a nonpolar core resembling to the hydrocarbon media to accommodate pyrene molecules. In contrast, the higher value, 0.96 found for Pluronic F127 micelles suggests that these micelles provide a microenvironment for the pyrene probe more polar than usual surfactants (in the domain of moderate polarity), due to the tendency of water penetration at the core-corona border [37]. The mixed Cremophor-Pluronic micelle exhibits an I3/I1 value similar to the pure Cremophor micelles, probably due to the higher molar ratio of Cremophor inside the mixed micelles than the actual molar ratio in the bulk surfactant solution.

The resistance of the micelles against dilution and pH changes was also checked.

The samples of pure and mixed micelles diluted 10 times show similar values of surfactant aggregates after dilution compared to concentrated ones. For drug loaded Cremophor EL-Pluronic F127 mixed micelles at α = 0.2, for example, average size of concentrated dispersion (1 × <sup>10</sup>−<sup>3</sup> M) in PBS was 26.41 ± 0.95 nm while after a tenfold dilution, the size was 26.33 ± 0.95 nm.

The pH value of the dispersion media is expected to show negligible influence on the micellar size and shape, since both surfactants are nonionic. Thus, the values of the average size of mixed micelles Cremophor EL-Pluronic F127 at α = 0.2 are found 26.97 ± 1.41 nm at pH = 4, 26.94 ± 2.54 at pH = 6, and 26.41 ± 0.95 nm at pH = 7.4, respectively. The mixed micelles prove to be resistant to dilution and no effect of pH variation in the range 4–7.4 affect the aggregation.

The DLS measurements were also applied to the samples after four weeks storage at room temperature. No significant changes were recorded in the size and size distribution of the void and NFLX.

#### *3.3. Drug—Polymeric Micelle Interactions*

To investigate the interaction of Norfloxacin molecules with the polymeric surfactants in micelles Fourier transformed infrared spectroscopy were used. In Figure 3, the FTIR spectra of pure Norfloxacin (1), Cremophor EL (2), Pluronic F127 (3), Norfloxacin in Cremophor EL micelles 100 μg/mL (4), Norfloxacin in mixed micelles Cremophor EL-Pluronic F127 (5) and Norfloxacin in Pluronic F127 micelles 100 μg/mL (6) are displayed.

**Figure 3.** FTIR spectra of (1) Norfloxacin, (2) Cremophor EL, (3) Pluronic F127, (4) Norfloxacin in Cremophor EL micelles 100 μg/mL, (5) Norfloxacin in mixed micelles Cremophor EL-Pluronic F127 and (6) Norfloxacin in Pluronic F127 micelles 100 μg/mL.

The FTIR spectrum of Norfloxacin shows specific absorption peaks, similar to those reported in literature [13] as follows: the wide band centered at 3438 cm−<sup>1</sup> is produced by both imino moiety of piperazinyl groups (–NH stretching vibration) and –OH group from acid and the band at 2881 cm−<sup>1</sup> correspond to C–H stretching vibrations. The absorptions at 1627 cm−<sup>1</sup> are characteristic for quinolones (–NH bending vibration) and the region between 1500–1450 cm−<sup>1</sup> for =O–C–O– group of acid (υ<sup>s</sup> stretching vibration). The bending vibration of –OH was found at 1272 cm−<sup>1</sup> and the strong absorption at 1114 cm−<sup>1</sup> was related to C–F group.

For Cremophor EL were registered a broad band at 3436 cm−<sup>1</sup> for the –OH group, the band at 2926 cm−<sup>1</sup> attributed to C-H stretch and the very small absorption band between 1715–1730 cm−<sup>1</sup> characteristic to C=O stretch for esters. The stretching band of C=C was found at 1642 cm−1, the band from 1101 cm−<sup>1</sup> was attributed to C-O stretch from alcohols and the wide absorption of 636 cm−<sup>1</sup> to =C-H bend, similar to reported data in the literature [39]. In the spectra of Pluronic F127 micellar systems specific signal were observed as broad band at 3439 cm−<sup>1</sup> for the –OH group, the bend of 2886 cm−<sup>1</sup> was attributed to C-H stretch, 1348 cm–1 (in-plane O-H bend) and 1112 cm–1 (C-O stretch) [40].

For the sample with Norfloxacin in Cremophor EL micelles, FTIR spectrum absorptions at 2923 cm−<sup>1</sup> attributed to C-H stretch, 1643 cm<sup>−</sup>1, stretching band of C=C, 1101 cm−<sup>1</sup> attributed C-O stretch from alcohols (identical to that of Cremophor), 950 cm−<sup>1</sup> attributed to *trans*-CH=CH- group and 669 cm−<sup>1</sup> attributed to =C-H bend. The most evident change is the broadening and increase intensity of the absorption band at 3438 cm<sup>−</sup>1, probably due to the formation of numerous hydrogen bonding with OH group from acid (Norfloxacin) and from alcohol (Cremophor) and other interactions with the imino moiety of piperazinyl groups (NH stretching vibration).

As a result of the encapsulation of the Norfloxacin in Pluronic F127 micelles no significant changes were observed in the peak intensities and positions, with the exception of the wide absorption band in the 3500 cm−<sup>1</sup> region, which is wider and more intense, due to the formation of hydrogen bonds between drug and polymer molecules. In the spectra of encapsulated Norfloxacin in the mixed micelles changes in shape and position of peaks could not be observed, except the same broadening and shifting to higher wavenumber of the peak at 3438 cm−<sup>1</sup> to 3464 cm−<sup>1</sup> and a minor shift to higher wavenumber of the peak corresponding to C-F bending. These changes are due to the interactions of the Norfloxacin with Cremophor and Pluronic F127, as a result of encapsulation of the Norfloxacin in the mixed micelles.

#### *3.4. Drug Solubility and Encapsulation Efficiency*

The maximum solubility of NFLX in micelles and encapsulation efficiency are tabulated in Table 2.

**Table 2.** Amount of Norfloxacin (NFLX) solubilized in polymeric micelles and encapsulation efficiency at 25 ◦C.


The concentration in the micellar dispersion were selected to represent 20 time the CMC value, in order to compare the solubility in micelle aggregates rather than conventional procedure versus weight of the polymeric material. The drug loading capacity expressed as % of NFLX from the weight of drug and micelle forming materials is 5% for Cremohor EL, 4.67% for Pluronic F127 and 4.89% for mixed micelles Cremohor EL-Pluronic F127 with molar ration α = 0.2 (thus low content of Cremophor). One can conclude that a high loading capacity is maintained in mixed micelles, even at a significantly decreased surfactant concentration at the molar ratio where the synergistic effect is present, due to the favored micellization process.

#### *3.5. In Vitro Drug Release*

The release kinetics of Norfloxacin from various micellar dispersions was evaluated using PBS as receiving media and the results are presented in Figure 4.

The drug release profile is significantly dependent to the composition of micelles. Norfloxacin encapsulated in Cremohor EL micelles exhibit a cumulative release less than 20% up to 48 h, probably due to the compact packing of the hydrophobic chains of the surfactant inside the inner core and higher tendency of drug retention. The release profile show a burst in the first hour, then a decrease in the release rate. The burst region is present also in the Pluronic F127 micellar dispersion for the first 2.5 h, but cumulative release is far more important, up to 64%. The Norfloxacin molecules could be located both in the hydrophobic inner region of polypropylene oxide (PPO) and embedded in the hydrophilic corona of polyoxyethylene (POE) groups in the Pluronic micellar aggregate and lead to a higher extent of drug released than from Cremophor EL micelles. The release from the mixed micelles retains the burst segment, less pronounced compared to the situation in Cremophor micelles, due to the small amount of this polymeric surfactant in α = 0.2 selected composition. The cumulative release is improved compared to Cremophor EL micelles, due to the influence of the large Pluronic, content up to 49% at 48 h.

**Figure 4.** In vitro drug release profile for NFLX in Cremohor EL, Pluronic F127 and CrEL-Pl F127 (α = 0.2) mixed micelles.

The influence of the temperature and pH on the release profile of NFLX in selected mixed micelles (Cremophor-Pluronic F127 at molar ratio 0.2) is presented in Figure 5.

**Figure 5.** In vitro drug release profile for NFLX in mixed CrEL-Pl F127 (α = 0.2) micelles at various temperature (**a**) and pH values (**b**).

Since both Cremophor EL and Pluronic F127 are nonionic surfactants the effect of temperature up to 50 ◦C is negligible on the size and morphology of the micellar aggregates, thus the release of Norfloxacin encapsulates in either pure or mixed micelles is not significantly affected by the increase of the temperature from 25 ◦C to 37 ◦C. The kinetic profile is similar for the two temperatures investigated up to 12 h, with a moderate increase (from 51.8% at 25 ◦C to 72.0% at 37 ◦C) at 24 h, probably due to the increase in the aqueous solubility of drug when rise the temperature.

The release profile at various pH values shows an increase of the cumulative drug release at 24 h up to 60.80% at pH = 4, while at the pH=6a decrease of the total NFLX release to 47% is observed. This unexpected variation could be the result of the complex equilibrium of the drug species inside the micelles and release media, due to the peculiar variation of the solubility of Norfloxacin with pH.

This behavior is not the result of changes in micelle properties, but of the intrinsic properties of the drug. Size and size distribution of the Cremophor-Pluronic F127 (α = 0.2) micelles mixed micelles, as well as those for pure Cremophor or Pluronic micelles is not affected by the modification of the pH, according to the DLS measurements presented in Section 3.2.

The chemical structure of Norfloxacin produces amphiphilic, ionized and neutral species with variation of the pH, i.e at pH = 7.4 existence of both neutral NFLX0 and zwitterionic NFLX± was evidenced, at pH=6a quasi-equimolecular mixture of zwitterionic NFLX<sup>±</sup> and ionic NFLX+ is present and at pH = 4 only the ionic NFLX<sup>+</sup> is observed. This ionization behavior is consistent with the dramatic changes in solubility of NFLX with the pH, from 0.3 M at pH = 5 to 2.9 × <sup>10</sup>−<sup>3</sup> M at pH = 6 and 1.3 × <sup>10</sup>−<sup>3</sup> M at pH = 7 [41].

From the significant increase of the aqueous solubility of Norfloxacin at pH = 4, one can expect a more obvious increase in the drug released. However, the fully charged species NFLX+ is stronger retained inside the micelles due to the interaction with the –OH and –COOH groups from the Pluronic and Cremophor molecules. At pH = 6 the coexistence of both zwitterionic and protonated Norfloxacin species results in a lower content of drug released.

The combination of the two polymeric surfactants results in a sustained release of NFLX form micelles that can ensure a long-term delivery of the antibiotic.

#### *3.6. Biocompatibility of Mixed Micelles*

The effect of surfactants on normal cells was studied in terms of cell viability by MTT assay and cellular inflammation and membrane damage with nitric oxide (NO) release test. The study was performed on human lung fibroblasts MRC-5 cells selected as model normal cells with moderate sensibility to dispersions. The results are shown in Figures 6 and 7.

As shown in Figure 6, the Cremophor EL micellar solution in the range 5 × <sup>10</sup>−6– <sup>2</sup> × <sup>10</sup>−<sup>4</sup> <sup>M</sup> did not affect the number of viable cells after 24 h of incubation compared with the control. Although no change in cell viability was measured for sample Cremophor EL even at concentration approximative 5 times higher the CMC, a significant decrease was noticed for the 3.2 × <sup>10</sup>−<sup>4</sup> M concentration, after both time intervals of incubation with surfactants.

**Figure 6.** *Cont*.

**Figure 6.** Cell viability results obtained by MTT assay after 24 and 48 h of cell growth in the presence of surfactants. Results are presented as mean ± standard deviation of three independent experiments (\* *p* < 0.05, \*\* *p* < 0.01 and \*\*\* *p* < 0.001 compared with control).

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**Figure 7.** *Cont*.

**Figure 7.** NO release measured by Griess assay after 24 and 48 h of cell growth in the presence of surfactants. Results are presented as mean ± standard deviation of three independent experiments (\*\* *p* < 0.01 and \*\*\* *p* < 0.001 compared with control).

The micellar dispersions based on Pluronic F127 show lack of cytotoxicity up to 1.5 × <sup>10</sup>−<sup>4</sup> M, as it is reported in most paper for low concentrated polymeric solutions below 8 × <sup>10</sup>−<sup>6</sup> M, in premicellar region [42]. An unexpected dose-dependent decrease was obtained for Pluronic concentrated micellar disperions from 3 × <sup>10</sup>−<sup>4</sup> M to <sup>6</sup> × <sup>10</sup>−<sup>4</sup> M, where the cellular viability decreased to 49% compared to control value after 48 h of exposure.

In the mixed micelle system with molar ratio Cremophor EL- Pluronic F127 α = 0.2, the dose dependent variation of the cytotoxicity is observed similar to Pluronic F127 micellar dispersion, probably due to the small amount of the Cremophor in the mixture. Significant decrease in the cellular viability is recorded after 24 and 48 h at higher concentration than <sup>2</sup> × <sup>10</sup>−<sup>4</sup> M of mixed surfactants.

Since the synergistic molar ratio <sup>α</sup> = 0.2 exhibits the low value of CMC <sup>4</sup> × <sup>10</sup>−<sup>5</sup> M, solutions with concentration 3-fold CMC will ensure the existence of the micellar aggregates and remains in the region without evident toxicity.

The amount of NO released in the culture medium was assessed as a valuable indicator of inflammation produced by the contact with surfactant solutions in premicellar and micellar regions. As it is shown in Figure 7, no significant changes in NO release after cell exposure to surfactants over whole range of studied concentrations could be observed. Thus, it was concluded that even in the case of the Cremohore EL presence, the surfactant micelles did not induce inflammation in human fibroblast cell cultures.

#### *3.7. Antibacterial Activity*

The qualitative screening of the antimicrobial activity evaluated the efficiency of the Norfloxacin encapsulated in Cremophor EL and Pluronic F127 micellar carriers, by measuring the diameters of the inhibition zone expressed by each tested bacterial strain. The value of the diameters of the inhibition zones demonstrated the achievement of a concentration gradient around the spot following the release of the antibiotic from the micellar carrier (20 μg in 10 μL micellar solution), by comparison with the concentration gradient achieved by the release of antibiotics from the standard disc (30 μg/disk) (Figure 8).

**Figure 8.** Aspect of the inhibition zones for microorganisms (**a**) *Staphylococcus aureus* ATCC 25923, (**b**) *Pseudomonas aeruginosa* ATCC 27853 and (**c**) *Escherichia coli* ATCC 25922. The samples are denoted P1 = NFLX in water; P2 = NFLX in DMSO; P3 = NFLX in Cremophor EL micellar solution; P4 = NFLX in Pluronic F127 micellar solution; P5 = NFLX in mixed micellar solution, while M1, M2 and M3 are empty micelles of Cremophor EL, Pluronic F127 and mixed micelles, respectively.

> It cannot be observed significant difference between Cremophor and Pluronic micellar carriers (P3 and P4) or combination of them (P5) on the growth of standard bacterial strains. In Figure 9 the values of the diameter of inhibition zone recorded for both clinical and standard bacterial strains are presented.

> As shown in Figure 9, different behaviors for the clinical and the standard bacterial strains exposed to NFLX are recorded, with the clinical strains expressing larger diameters of the inhibition zones in both cases standard disc and micellar carriers. The exception is the *P. aeruginosa* 5399 clinical strain which shows low sensitivity to Norfloxacin in all the tested systems. It seems that this bacterial strain is resistant to fluoroquinolone, given the diameter of the inhibition zone for the norfloxacin (16 mm < 18 mm, according with CLSI 2021) and by using the micellar carrier for antibiotic the sensitivity pattern has not changed when NFLX was encapsulated in Cremophor and mixed micelles compared to NFLX in aqueous solution. A rather higher sensibility is recorded to the NFLX encapsulated in Pluronic F127 (15 mm), probably due to the cellular membrane permeability induced by the tri-block copolymer micelles.

> For the *E. coli* strains, both standard and clinical strains, the diameter of inhibition zones produced by NFLX encapsulated in micelles were larger than the antibiotic in aqueous solution.

> The quantitative results expressed by minimal inhibitory concentration values allowed the quantitative evaluation of the carriers' efficiency in terms of antibiotic release capacity in the liquid medium (Figure 10).

**Figure 9.** Graphic representation of inhibition zone diameters (mm).

**Figure 10.** Graphic representation of minimal inhibitory concentration values for: (**a**) Gram negative tested strains and (**b**) Gram positive tested strains.

For *E. coli* ATCC 25922 strains the minimal inhibitory concentrations were similar for the reference aqueous solution of NFLX and the drug encapsulated in various micellar systems, with the exception of NFLX in mixed Cremophor EL- Pluronic F127 micelles, which exhibited a lower MIC of 0.039 μg/mL compared to 0.078 μg/mL. The clinical strain *E. coli* is less susceptible to NFLX encapsulated in Cremophor EL (MIC of 0.31 μg/mL) and again the highest activity was observed for the formulation with mixed surfactants (MIC 0.078 μg/mL).

In the case of *P. aeruginosa* ATCC 27853, a similar efficiency was observed for NFLX in aqueous solution and Pluronic micelles (MIC 0.625 μg/mL), while the presence of Cremophor EL in pure and mixed micelles reduced the MIC value to 0.312 μg/mL. As observed in the qualitative test, *P. aeruginosa* clinical strain proved to be more resistant to NFLX, and the encapsulation in micelles leads to an unexpected increase of the MIC values for all surfactants.

In the case of the Gram-positive microorganisms there were also differences in the results recorded for standard and clinical strains depending on the composition of the micellar carrier. For standard *E. fecalis* ATCC 29212 the lowest efficiency was demonstrated by the NFLX loaded Cremophor EL micelle dispersion (MIC value 0.312 μg/mL), while the encapsulation in Pluronic F127 and in the mixed micelles reduced the MIC value to quarter. The clinical *E. fecalis* VRE 2566 strain developed a high value of MIC in the case of NFLX in Pluronic micelles (MIC value 0.625 μg/mL). Again, the encapsulation in mixed micelles decreased the value of MIC up to 0.125 μg/mL.

For *S. aureus* standard strain ATCC 25923 the MIC values for NFLX in all micellar systems proved to be double compared to the value recorded for aqueous solution (0.625 μg/mL compared to 0.312 μg/mL).

In the case of resistant *S. aureus* strain, no significant improvement was observed with the Pluronic F127 micelles, on the contrary the MIC value obtained is the higher obtained in our study (1.25 μg/mL) for Gram positive microbial cultures. The presence of Cremohor EL as pure micelles or in combination with Pluronic F127 increased the antibacterial efficiency of the encapsulated drug.

The micelles of two polymeric surfactants used in this study had different effects on the Norfloxacin antibacterial efficiency on different microorganisms, and a variation in behavior could also observed between standard and clinical strains. A possible explanation is the specific interaction between the cellular membrane and micelle forming surfactants, that should be investigated in detail in order design drug delivery systems for NFLX with enhanced activity against resistant microbial strains.

Pluronic polymeric surfactants have begun to be investigated to elucidate their role in management of biofilm formation, but relevant data and explanations about the complex interactions with the microbial cellular membrane are still lacking [43].

#### **4. Conclusions**

Polymeric micelles were prepared as drug delivery systems for model poorly soluble antibiotic Norfloxacin in order to achieve better stability and controlled delivery of drug. Two polymeric surfactants Cremophor EL and POE-PPO-POE triblock copolymer Pluronic F127 and their mixtures in various molar ratio were studied. The non-ideal behavior of the self-assembling process in surfactant mixtures reveal the existence of synergistic effects over the whole range of composition, with a maximum value of interaction parameter in mixed micelle for the molar ratio α = 0.2. The micelles were characterized in term of size, size distribution and drug solubilization, and the selected composition with low content of Cremophor EL show suitable performance to be used as Norfloxacin carrier.

The mixed micelles selected exhibit resistance against dilution and pH changes in the range of 4–7.4. The encapsulation efficiency of NFLX in mixed CrEL—Pl F127 micelles was 52.2 ± 2.1% and do not significantly change the size of nanocarrier after the drug encapsulation.

The drug release profile exhibited an initial burst (more obvious in Cremophor EL micelles) in all micellar dispersion, with a cumulative release up to 51.8% in mixed Cremophor EL—Pluronic F127 system at 25 ◦C, intermediate from the value obtained for Cremphor EL and Pluronic F127 pure micelles. The total amount released in 24 h was increased to 72% at temperature of 37 ◦C. The results of cytotoxicity and inflammation tests on normal fibroblast cells indicate that both studied surfactants and their mixtures prove high biocompatibility for concentrations below 2–3 × <sup>10</sup>−<sup>4</sup> M.

The analysis of qualitative and quantitative results showed an improvement of the antibacterial properties of Norfloxacin against the majority of the tested bacterial strains, both Gram positive and Gram negative, when the polymeric micelles were used as carriers.

The sample with NFLX encapsulated in the novel carrier prove to be effective on both *E. coli* ATCC 25922 and clinical strains, with MIC value of 0.039 μg/mL and 0.078 μg/mL, respectively. The minimum inhibitory concentrations were found to vary greatly with nature and provenience of the microbial strain and with the surfactant composition of the

formulation. As a general conclusion, the drug loaded mixed micellar formulation exhibits good activity against clinical isolated strains, compared with the CLSI recommended standard strains, thus Cremophor EL-Pluronic F127 mixed micelles can be considered as novel controlled release delivery systems for hydrophobic antimicrobial drugs.

**Author Contributions:** Conceptualization, L.O.C. and C.P.; methodology, C.L.N.; validation, L.O.C. formal analysis, C.P.; investigation, M.A.T., A.R., P.O. spectroscopy, L.M.D. microbiology, M.S. biocompatibility, C.S., C.M.N. FTIR; writing—original draft preparation, L.O.C. and M.A.T.; writing review and editing L.O.C. and C.P., project administration, L.O.C. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Romanian Ministry of Education and Research, CCCDI— UEFISCDI, project number PN-III-P2-2.1-PED-2019-4657, within PNCDI III.

**Institutional Review Board Statement:** Not Applicable.

**Informed Consent Statement:** Not Applicable.

**Data Availability Statement:** Not Applicable.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

#### **References**


## *Article* **Hybrid Nanoparticles of Poly (Methyl Methacrylate) and Antimicrobial Quaternary Ammonium Surfactants**

#### **Beatriz Ideriha Mathiazzi and Ana Maria Carmona-Ribeiro \***

Biocolloids Laboratory, Departamento de Bioquímica, Instituto de Química, Universidade de São Paulo, Av. Prof. Lineu Prestes 748, 05508-000 São Paulo, Brazil; bemathi@usp.br

**\*** Correspondence: amcr@usp.br; Tel.: +55-011-3091-1887

Received: 23 March 2020; Accepted: 8 April 2020; Published: 10 April 2020

**Abstract:** Quaternary ammonium surfactants (QACs) are microbicides, whereas poly (acrylates) are biocompatible polymers. Here, the physical and antimicrobial properties of two QACs, cetyl trimethyl ammonium bromide (CTAB) or dioctadecyl dimethyl ammonium bromide (DODAB) in poly (methyl methacrylate) (PMMA) nanoparticles (NPs) are compared to those of QACs alone. Methyl methacrylate (MMA) polymerization using DODAB or CTAB as emulsifiers and initiator azobisisobutyronitrile (AIBN) yielded cationic, nanometric, homodisperse, and stable NPs. NPs' physical and antimicrobial properties were assessed from dynamic light scattering (DLS), scanning electron microscopy, and viability curves of *Escherichia coli*, *Staphylococcus aureus*, or *Candida albicans* determined as log(colony-forming unities counting) over a range of [QACs]. NPs were spherical and homodisperse but activity for free QACs was higher than those for QACs in NPs. Inhibition halos against bacteria and yeast were observed only for free or incorporated CTAB in NPs because PMMA/CTAB NPs controlled the CTAB release. DODAB displayed fungicidal activity against *C. albicans* since DODAB bilayer disks could penetrate the outer glycoproteins fungus layer. The physical properties and stability of the cationic NPs highlighted their potential to combine with other bioactive molecules for further applications in drug and vaccine delivery.

**Keywords:** hybrid nanoparticles; biocompatible polymer; antimicrobial amphiphiles; dynamic light scattering; scanning electron microscopy; cell viability from counting of colony-forming unities; antimicrobial activity of nanoparticles; *Escherichia coli*; *Staphylococcus aureus*; *Candida albicans*

#### **1. Introduction**

Biocompatible synthetic or natural polymers can improve body functions without interfering with its normal functioning or triggering side effects [1–7]. They have been useful in tissue culture, tissue scaffolds, implants, artificial grafts, wound dressings, controlled drug release, prosthetic replacements for bones, dentistry, etc [8]. Some examples of biocompatible polymers are poly (lactic-co-glycolic acid) [9], poly (ε-caprolactone) (PCL) [10], poly (lactic acid) [11], poly (3- hydroxybutyrate-co-3-hydroxyvalerate) (PHBV) [12], chitosan [13,14], cellulose [15], and poly (acrylates) including poly (methyl methacrylate) (PMMA) [6,16–18]. Among the poly (acrylates), several cationic latexes and coatings obtained from additives such as cationic surfactants, lipids, antimicrobial polymers or co-polymers have been described [19–25]. In the case of PMMA, the discovery of PMMA biocompatibility occurred when an optical technician placed a PMMA scleral lens on his eye and found that it could be well tolerated. The first PMMA patent appearing in 1948 [16] was acquired by Bausch & Lomb in 1972 [26].

In our laboratory, the good compatibility between PMMA and some antimicrobial quaternary ammonium compounds (QACs) was first described both for PMMA/QAC coatings [19] and

PMMA/QACs nanoparticles (NPs) [27]. The QACs were apparently able to impart their antimicrobial activity [28–31] to the PMMA/QACs assemblies [19,27]. In the absence of PMMA, QACs in water dispersion self-assemble to yield micelles or bilayers depending on the QAC chemical structure and molecular shape [32,33]. Dioctadecyl dimethyl ammonium bromide (DODAB) assembles in water solution as bilayer vesicles or bilayer fragments (BF) depending on the dispersion method; sonication disrupts DODAB vesicles producing BF [34–36]. Cetyltrimethylammonium bromide (CTAB) assembles in water solution as micelles above the critical micellar concentration [32]. Ion–dipole interactions between the quaternary ammonium in DODAB and the carbonyl moiety in PMMA plus other weak but cooperative van der Waals interactions led to hybrid materials where DODAB was found well dispersed in the polymeric matrix [19]. This allowed for combining the bactericidal DODAB property [29–31,37–42] with excellent DODAB immobilization and distribution in the PMMA polymeric network [19]. Later on, determining the effect of QAC chemical structure on the antimicrobial activity of PMMA/QAC coatings revealed that DODAB and CTAB behaved differently; CTAB diffused through the film and reached bacteria in the outer medium, whereas DODAB impregnation in the polymeric matrix killed bacteria upon contact without leakage [20].

In this work, PMMA/QAC NPs synthesized by emulsion polymerization of methyl methacrylate (MMA) using azo-bis-isobutyronitrile (AIBN) as initiator and DODAB BF or CTAB as emulsifiers were evaluated regarding their physical properties and antimicrobial activity as compared to the free QACs. Scheme 1 illustrates the emulsion polymerization of MMA in the presence of DODAB BF or CTAB using AIBN as the initiator.

Scheme 1. **Scheme 1.** Emulsion polymerization of methyl methacrylate (MMA) initiated by azobisisobutyronitrile (AIBN) using dioctadecyl dimethyl ammonium bromide (DODAB) or cetyltrimethylammonium bromide (CTAB) as emulsifiers. The cross sections of DODAB bilayer fragments (BF) and CTAB micelles were schematically represented as loaded with the MMA monomer (in red) before adding AIBN to initiate the polymerization.

Over a range of MMA or QAC concentrations, 0.4 M MMA and 2 mM QAC yielded optimal physical properties for the PMMA/DODAB and PMMA/CTAB NPs (nanometric size, low polydispersity, high and positive zeta-potential, high yield, and high colloidal stability). Inhibition halos by CTAB and PMMA/CTAB NPs against bacteria and yeast showed that PMMA/CTAB NPs behaved as reservoirs for the release of CTAB with time after dialysis. CTAB was able to move both through the dialysis

membrane and the agar to inhibit microbial growth. In contrast, DODAB incorporated in the PMMA polymeric matrix did not move in the agar and did not cross the dialysis membrane. For PMMA/DODAB NPs or DODAB BF, inhibition halos were not observed due to the lack of DODAB diffusion in the agar medium. The incorporation of QACs in the PMMA/QAC NPs reduced antimicrobial activity in comparison to the QACs in dispersion. CTAB was the most active microbicidal agent against the three microbes tested (*E. coli*, *S. aureus*, and *C. albicans*) reducing cell viability countings by 7 logs at submicellar concentrations. CTAB mobility in hydrated medium favored its electrostatic interaction and penetration through the microbes' cell wall and membrane, imparting a lytic effect on the cells. DODAB BF revealed an important fungicidal activity against *C. albicans* not described before; similar to rod-like copolymers assemblies, the disk-like cationic DODAB BF possibly entered the outer glycoproteins layer of the fungus penetrating the cell and causing a 5-log reduction in yeast viability. Besides applications in antimicrobial chemotherapy, PMMA/QAC NPs here described may find interesting uses also as immunoadjuvants due to their nanometric size, positive zeta-potential, narrow size distribution, and high colloidal stability. They are expected to combine well with oppositely charged antigens, such as proteins [43], peptides [44,45], or DNA [46–48] for subunit vaccines design as many cationic adjuvants do.

#### **2. Materials and Methods**

#### *2.1. Materials*

Methyl methacrylate (MMA), hexadecyltrimethyl ammonium bromide (CTAB) dioctadecyldimethyl ammonium bromide (DODAB), azobisisobutyronitrile (AIBN), agarose, Mueller-Hinton agar (MHA), D-glucose, NaCl, ethanol 99.9%, and cellulose acetate membranes with a molecular weight cutoff around 12,400 g/mol were obtained from Sigma-Aldrich (Darmstadt, Germany) and used without further purification.

#### *2.2. Preparation of CTAB or DODAB Dispersions in Water Solution*

CTAB and DODAB are quaternary ammonium amphiphiles (QACs) that were separately weighted and added to a 1-mM NaCl solution (pH 6.3) to yield a stock dispersion at 10 mM amphiphile. Whereas CTAB yielded a homogeneous and transparent dispersion, typical of its assembly in water as micelles, DODAB had to be dispersed ultrasonically with a macroprobe. The DODAB BF were obtained from the DODAB powder added to the aqueous 1 mM NaCl solution dispersed by sonication with tip at 85 W for 15 min above 47 ◦C, before centrifuging the dispersion for precipitation of titanium ejected from the tip (9300× *g* for 1 h at 4 ◦C). This yielded the DODAB BF as a somewhat turbid dispersion. These DODAB dispersions were previously characterized as containing open bilayer fragments that did not enclose a water compartment, were nano-sized, and able to incorporate hydrophobic drugs in the bilayer or antimicrobial peptides or polymers [34,35,44,45,49–54]. Both CTAB and DODAB dispersions were diluted from the stock dispersions to obtain the final desired concentrations. The analytical CTAB or DODAB concentrations were determined from halide microtitration [55,56].

#### *2.3. Synthesis of Waterborne PMMA*/*QACs Nanoparticles (NPs) by Emulsion Polymerization*

The PMMA/CTAB or PMMA/DODAB hybrid and polymeric NPs were obtained by emulsion polymerization of MMA using AIBN as the initiator of the MMA polymerization, similar to the synthesis previously described using potassium persulfate (KPS) as the initiator [27]. The synthesis protocol was similar to the one previously described for the synthesis of PMMA/PDDA NPs using AIBN as the initiator [21–23]. In order to eliminate oxygen, a flow of nitrogen gas was applied for a few minutes to 10 mL DODAB or CTAB dispersions in 1 mM NaCl added of the desired MMA concentration and 0.0036 g AIBN. The reaction mixture inside the glass tube was then closed with a cap, heated, and kept at 80 ◦C in a water bath for 1 h under periodic vortexing. Thereafter, the capped tube containing the reaction mixture was withdrawn from the water bath and allowed to reach room

temperature. The dispersions thus obtained were purified by dialysis against 2 L of ultrapure water (3×) for 24 h. Particle characterization took place after dialysis using dilutions of the original dispersion in 1 mM aqueous NaCl solution.

#### *2.4. Determination of Sizes, Zeta-Potentials, and Polydispersity of PMMA*/*QAC Dispersions by Dynamic Light Scattering (DLS)*

Size distributions, zeta-average diameters (Dz), and zeta potentials (ζ potentials) were obtained by dynamic light scattering (DLS) using a Zeta plus−Zeta potential Analyzer (Brookhaven Instruments Corporation, Holtsville, NY, USA) equipped with a 677-nm laser with measurements at 90◦. The polydispersity of the dispersions was determined by dynamic light scattering (DLS) following well-defined mathematic equations [57]. The mean hydrodynamic diameters (mean Dz) were obtained from the log-normal distribution of the light-scattering intensity curve against Dz. The ζ potentials were determined from the electrophoretic mobility (μ) and the Smoluckowski equation, ζ = μη/ε, where η and ε are the viscosity and the dielectric constant of the medium, respectively. Samples that underwent the DLS measurements were usually diluted from the original dispersions for optimal readings (50–100 μL PMMA/QAC dispersions in 2 mL of 1 mM NaCl). All measurements were performed in the DLS apparatus took place at 25 ± 1 ◦C.

#### *2.5. Visualization and Morphology of PMMA*/*QAC NPs from Scanning Electron Microscopy (SEM)*

Scanning electron microscopy (SEM) was performed using Jeol JSM-7401F equipment. Silicon <100> wafers were from Silicon Quest (Santa Clara, CA, USA) with a native oxide layer approximately 2 nm thick and used as substrates for casting the PMMA/QAC dispersions. The Si wafers with a native SiO2 layer were cut into small pieces of ca 1 cm2, cleaned with ethanol, and dried under an N2 stream. Samples of 0.050 mL PMMA/QAC dispersions were deposited on clean Si/SiO2 wafers and dried overnight in a desiccator. Then, the coatings were covered with a thin layer of gold before SEM analyses, as required for contrast and visualization. Mean diameters (D) for dry NPs were evaluated from ImageJ software Version 1.52u for 100 particles and presented as a mean value.

#### *2.6. Microorganisms Cultures and E*ff*ect of CTAB, DODAB, or PMMA*/*QAC NPs on Cell Viability in the Presence of the Cationic Amphiphiles Solutions or Dispersions*

*Escherichia coli* ATCC (American Type Culture Collection) 25922, *Staphylococcus aureus* ATCC 29213 or *Candida albicans* ATCC 90028 were cultured from previously frozen stocks (kept at −20 ◦C in the appropriate storage medium). Each microorganism was reactivated separately, seeded by streaking technique on the plates of Mueller-Hinton agar, and incubated for 18–24 h at 37 ◦C. The turbidity of either bacteria or fungus suspensions was adjusted according to tube 0.5 of the McFarland scale at 625 nm in isotonic 0.264 M D-glucose solution. The 0.264 M D-glucose solution was used instead of any culture medium because cationic molecules are inactivated by the relatively high ionic strength or negatively charged molecules, such as amino acids and polysaccharides. For the determination of cell viability, 0.1 mL of the cell suspensions (around 107–108 colony-forming unities per mL, CFU.mL<sup>−</sup>1) were mixed with 0.9 mL of NPs dispersions diluted in the same D-glucose solution and interacted for 1 h. Thereafter, aliquots of 0.1 mL were withdrawn and either directly plated or diluted 10 to 106 times before plating on MHA plates. The plates were incubated at 37 ◦C for 24 h. The CFU were counted and plotted in a logarithmic or percentage scale as a function of QAC concentration (mM). When no counting was obtained, since the log function does not exist for zero, the CFU counting was taken as 1 so that log CFU.mL−<sup>1</sup> could be taken as zero.

#### *2.7. Determination of Growth Inhibition Zones by PMMA*/*QAC NPs*

*Escherichia coli* ATCC (American Type Culture Collection) 25922, *Staphylococcus aureus* ATCC 29213, and *Candida albicans* ATCC 90028 from previously frozen stocks kept at −20 ◦C in an appropriate storage medium were grown as described above, and the bacterial suspension prepared in 0.264 M D-glucose had its turbidity adjusted according to 0.5 of the McFarland scale, as previously described. A softer growth medium containing 2.3% Muller-Hinton broth and 0.64% agar was prepared and sterilized in steam autoclave at 121 ◦C for 20 min. In 50 mL of this growth medium, 0.5 mL of the microorganism suspension was added and then carefully homogenized. Plates containing MHA were previously prepared and used to place micropipette tips with their bases positioned on the MHA to form wells with a 9 mm diameter. Before withdrawing the tips, ca. 20 mL of the microorganism culture in the soft MHA was allowed to harden. After agar hardening, the tips were removed and, in each well, 100 μL of the QAC dispersions at 0.01, 0.1, 0.2, 0.5, 1, 1.5, 2, and 2.5 mM or of the NPs dispersions were added for incubation 18–24 h at 37 ◦C for determining the inhibition zones surrounding the wells. From the comparison between inhibition zones for the standard QAC dispersions and the PMMA/QAC NPs, the QAC concentration in the NPs dispersions was estimated. 2020

#### *2.8. Determination of QAC Concentration from Halide Microtitration*

Bromide is the counterion of DODAB and CTAB. Similar to chloride, bromide can be determined by halide microtitration [55], as given in a detailed protocol for halide microtitration [56]. 3. Results and Discussion

#### **3. Results and Discussion**

#### *3.1. Synthesis of PMMA*/*QACs NPs by Emulsion Polymerization and their Physical Characterization from SEM and DLS*

PMMA/QAC NPs' morphology, size, and homogeneity were assessed by SEM. Two different PMMA/QAC NPs were synthesized and observed after drying under SEM (Figure 1). The QAC was DODAB (Figure 1a) or CTAB (Figure 1b). Both syntheses employed 0.4 M MMA, 2 mM QAC, 0.0036 g AIBN and 1 mM NaCl in 10 mL of reaction mixture. The NPs were spherical, displayed a narrow size distribution and were homo dispersed. PMMA/DODAB and PMMA/CTAB mean diameters after drying (D) were obtained for at least 100 particles from the ImageJ software as 56 ± 7 and 85 ± 11 nm, respectively (Figure 1).

Figure 1. a b **Figure 1.** Scanning electron micrographs (SEM) of original dispersions of poly methyl methacrylate /dioctadecyldimethylammonium (PMMA/DODAB) (**a**) or PMMA/CTAB nanoparticles (NPs) on silicon wafers (**b**). The magnification was either 25,000× (on the top) or 70,000× (on the bottom).

The mean diameter of dry NPs (D) was compared to the one of NPs in water dispersion given by dynamic light scattering (DLS) as the mean hydrodynamic diameter (Dz) (Table 1).

**Table 1.** Dry mean diameter (D) and mean hydrodynamic diameter (Dz) for PMMA/QAC NPs. D was from scanning electron microscopy (SEM) and Dz was determined by dynamic light-scattering (DLS). The nanoparticles (NPs) were obtained from 0.4 M methyl methacrylate (MMA), 2 mM dioctadecyl dimethyl ammonium bromide (DODAB) or cetyl trimethylammonium bromide (CTAB), and 0.36 mg·mL−<sup>1</sup> AIBN in 10 mL of 1 mM NaCl. The mean D was obtained using ImageJ software from 100 particles.


For NPs obtained from 0.4 M MMA, 2 mM DODAB, or CTAB using 0.0036 g AIBN, the mean hydrodynamic diameters (Dz) were 75 ± 1 and 81 ± 1 nm for PMMA/DODAB and PMMA/CTAB NPs, respectively. One would expect that the Dz values from DLS would be higher than D from SEM due to an eventual hydration layer surrounding each NP. Indeed, this occurred for the PMMA/DODAB NPs but did not occur for the PMMA/CTAB NPs. It is possible that the PMMA/CTAB NPs exhibited a less hydrophilic surface at the particle/water interface than the PMMA/DODAB NPs. This was possibly due to the mobile character of the CTAB molecules leaving the nanoparticles to the bulk water and the large affinity of the DODAB molecules for the PMMA polymeric matrix [19,20]. Whereas DODAB would improve the NP affinity for the surrounding water and the hydration layer, this would not be very significant for CTAB so that, for CTAB, the dry, dehydrated diameter D would not be significantly lower than the hydrodynamic diameter Dz. D was 56 ± 7 nm for PMMA/DODAB NPs and 85 ± 11 nm for PMMA/CTAB NPs.

Curiously, DODAB behaved as a better emulsifier than CTAB during the emulsion polymerization process remaining in the PMMA/DODAB NPs after polymerization due to its higher affinity for PMMA than the one exhibited by CTAB [19,20]. At this point, one should recall the nanostructures present in DODAB and CTAB dispersions in water: the DODAB bilayer fragments (DODAB BF) with Dz inside the 56–67 nm range, as determined by DLS or SEM [44,58,59] or CTAB micelles with 10–20 nm of diameter determined by small-angle neutron scattering (SANS) [60]. IT is possible that the higher amount of DODAB in DODAB BF as compared to the amount of CTAB in the micelle improved the emulsifying effect of DODAB and resulted in a smaller NP size for the PMMA/DODAB NPs than the one for the PMMA/CTAB NPs. Since CTAB has only one hydrocarbon chain, its hydrophilic–hydrophobic balance is larger than the one for DODAB with two long hydrocarbon chains. The resulting emulsifying effect during NP synthesis promoted by CTAB was inferior to the one promoted by the DODAB BF so that sizes for the resultant NPs were larger than those synthesized with DODAB BF.

#### *3.2. E*ff*ects of MMA Concentration, QAC Concentration, and Initiator Type on Physico-Chemical Properties of PMMA*/*QAC NPs Obtained by Emulsion Polymerization*

Over a range of [MMA] varying from 0.1 to 1 M MMA, PMMA/QAC dispersions exhibited variable colloidal stability after synthesis and before dialysis. Above 0.4 M MMA, at 2 mM QAC, aggregation and precipitation took place so that only the supernatants were dialyzed and used for DLS and solid contents analysis. From 0.1–0.4 M MMA, no precipitation took place after synthesis and before dialysis.

Table 2 and Figure 2 show the effect of [MMA] on the physical properties of PMMA/QAC NPs obtained by emulsion polymerization at 2 mM QAC.

The data on Table 2 reveal that increasing [MMA] from 0.1 to 0.4 M at 2 mM DODAB, slightly increased the Dz for the NPs, increased ζ, reduced P, increased the solid contents and conversion of monomer into polymer, and slightly increased number density (Np) for the NPs, in absence of

aggregation. Above 0.4M MMA, poor colloidal stability associated with low zeta-potential was depicted as precipitation at the bottom of the assay tubes (Table 2). For PMMA/CTAB NPs, similar behavior took place for the physical properties except for the zeta-potential that remained low and approximately constant 17–23 mV. These results for the compared zeta-potentials for PMMA/DODAB and PMMA/CTAB NPs, again, suggest that increasing [MMA] leads to increased incorporation of DODAB in the NPs, whereas the incorporation of CTAB remains low and practically unaffected by the increased amount of solid contents.

**Table 2.** Physical properties of PMMA/QAC NPs obtained by emulsion polymerization of MMA monomer over a range of [MMA] at 2 mM DODAB or CTAB using AIBN as the initiator. Zeta-average diameter (Dz), zeta potential (ζ), and polydispersity (P) were obtained by dynamic light scattering. The conversion of monomer into polymer was expressed in percentile. The particles' number density (Np), in mL<sup>−</sup>1, was calculated from nanoparticle size (Dz) and the mass of lyophilized dispersions. Solid contents, conversion, and Np were determined as previously described in [27].


Figure 2 gives an overview of the effect of [MMA] on the physical properties of the PMMA/DODAB and PMMA/CTAB NPs' dispersions. The green rectangle emphasizes the NPs obtained with desirable characteristics, such as low size, low polydispersity, high zeta-potential, high yield, high particle number density (Np), and high colloidal stability (absence of aggregates and precipitates). The synthesis performed at 0.4 M MMA was selected as the one yielding the optimal NPs to be analyzed regarding their antimicrobial properties. One should notice that at 0.1 M [MMA] in the presence of DODAB BF, the high P and large sizes obtained might be explained from the intrinsically high polydispersity of the DODAB BF; polymerization took place inside the bilayer fragments, and NPs acquired their polydispersity.

The effect of [QAC] on the physical properties of the PMMA/QAC NPs is shown in Table 3. Increasing [QAC] reduced Dz, decreased the zeta-potential of PMMA/DODAB NPs and increased the one of PMMA/CTAB NPs, increased the polydispersity, and barely affected conversion or particle number density. The highest zeta-potentials for PMMA/DODAB NPs occurred at the lowest [DODAB], meaning that DODAB imparted high positive charges to the NPs but preferred to interact with other DODAB BF in dispersion when [DODAB] increased. For PMMA/CTAB, similar behavior to the one of PMMA/DODAB NPs was found except for the zeta-potential that decreased with [DODAB] and increased with [CTAB]. This means that the low affinity of CTAB for PMMA again explained the low incorporation of CTAB to this polymer over a range of low CTAB concentrations. Only when [CTAB] increased did it become possible to cause an increase in the zeta-potential of the PMMA/CTAB NPs from 15 to 40 mV due to increased CTAB incorporation in the NPs (Table 3). Increasing [QAC], affected the polydispersity that also increased for both types of NPs. QACs indeed tend to increase the size of their aggregates with increase in their concentration or on the ionic strength of the medium [60–62]. The selected concentration of MMA was 0.4 M because, above 0.4 M MMA, poor colloidal stability

was depicted from a visual observation of precipitated material; in this case, the measurements were performed with the dialyzed supernatants. Below 0.3 M MMA, low MMA concentrations resulted in low conversion (yield %), high polydispersities (P), and comparatively low zeta-potentials (ζ). The green rectangle indicates the region of [MMA] yielding stable PMMA/QACs NPs at high conversion rates, maximal particles number densities (Np), low P (0.03–0.06), high ζ (20–50 mV), and low Dz (70–75 nm). <sup>2020</sup>

**Figure 2.** Figure 2. Physical properties of PMMA/QAC NPs obtained by emulsion polymerization over a range of [MMA] at 2 mM DODAB (-) or CTAB (Δ). All dispersions were exhaustively submitted to dialysis in pure water before measurements. Above 0.4 M MMA, poor colloidal stability was depicted from the visual observation of precipitated material; the measurements were performed with the dialyzed supernatants. Below 0.3 M MMA, low MMA concentrations resulted in low conversion (yield %), high polydispersities (P), and comparatively low zeta-potentials (ζ). The green rectangle indicates the region of [MMA] yielding stable PMMA/QACs NPs at high conversion rates, maximal particle number densities (Np), low P (0.03–0.06), high ζ (20–50 mV), and low Dz (70–75 nm).

**Table 3.** Physical properties of PMMA/QAC NPs obtained by the emulsion polymerization of MMA monomer at 0.4 M MMA over a range of [QAC] using AIBN as the initiator. Zeta-average diameter (Dz), zeta potential (ζ), and polydispersity (P) were obtained by dynamic light scattering. The conversion of monomer into polymer was expressed in percentile. The particles number density (Np), in mL<sup>−</sup>1, was calculated from nanoparticle size (Dz) and the mass of lyophilized dispersions.


Figure 3 showed the desired properties for the PMMA/QAC dispersions at 2 mM QAC (green line), which was the concentration selected for the evaluation of antimicrobial activity. At 2 mM DODAB or CTAB, there was low size, low polydispersity, high zeta-potential, high yield, high Np and good colloidal stability for the PMMA/QAC NPs.

Two initiators were compared for the NPs synthesis: potassium persulfate (KPS) and AIBN. The initiator effect on NP properties is shown in Table 4. The MMA concentration selected for the comparison was identical to the one previously used in [27], where NPs had been synthesized with 0.56 M MMA using KPS as initiator.

**Table 4.** Comparison between nanoparticles of PMMA/QAC synthesized using either potassium persulfate (KPS) or azobisisobutyronitrile (AIBN) as initiators, at 0.56 M MMA and 2 mM QAC.


<sup>1</sup> Results taken from the reference [27].

Imparting the positive charge on the NPs was a difficult task in the presence of the negative sulfate charges on the PMMA/ DODAB particles coming from the synthesis with KPS as the initiator; at 2 mM DODAB during the NPs synthesis, the zeta-potential was still negative (ζ = −10 ± 1 mV) and the particles were large (Dz = 1260 ± 43) and very polydisperse (P = 0.370), suggesting poor colloidal stability (Table 4). Naves and coworkers used large concentrations of the QACs, such as ca. 10 mM DODAB, to revert the KPS effect and obtain positively charged NPs [27]. AIBN was a much more convenient initiator since, when using only 2 mM DODAB, the dispersion contained small NPs (Dz = 89 ± 1) that were positively charged (ζ = +45 ± 2 mV) and homodispersed (P = 0.027 ± 0.010). Similar results were obtained from the comparison between PMMA/CTAB NPs synthesized with KPS and PMMA/CTAB NPs synthesized with AIBN (Table 4). In summary, at 2 mM QAC, the

comparison between NPs synthesized using KPS or AIBN indicated that the low sizes, the high positive zeta-potentials, and the low polydispersities occurred only using AIBN. <sup>2020</sup>

Figure 3. **Figure 3.** Physical properties of PMMA/QAC NPs obtained by emulsion polymerization over a range of [QAC] at 0.4 M MMA during particle synthesis where QAC is DODAB (-) or CTAB (Δ). All dispersions were exhaustively submitted to dialysis in pure water before measurements.

#### *3.3. Incorporation of QACs in the PMMA*/*QAC NPs*

Table 4. DODAB and CTAB have a different hydrophobic–hydrophilic balance, as depicted from their chemical structure. The double-chained DODAB tends to prefer more hydrophobic environments than the single-chained CTAB. The determination of QAC incorporation in the PMMA polymeric matrix of the NPs showed the higher incorporation of DODAB in comparison to the one of CTAB (Table 5).

NPs Initiator Dz /nm Ί /mV P

**Table 5.** Determination of QAC concentration [QAC] in PMMA/QAC NPs from halide microtitration in the supernatants of PMMA/DODAB and PMMA/CTAB NP water dispersions before and after dialysis. The PMMA/QAC dispersions were prepared in 2.0 mM QAC. The controls for the dialysis were QAC dispersions in water and 1 mM NaCl solution.


<sup>1</sup> Microtitration done for supernatants of dialyzed dispersions 3 days after dialysis.

A CTAB control solution with 2.5 mM CTAB before dialysis permeated the dialysis membrane almost completely, yielding 0.1 mM CTAB just after dialysis (Table 5). DODAB BF at 2 mM DODAB, on the contrary, did not permeate the dialysis bag. Similarl to CTAB, NaCl permeated the dialysis membrane almost completely. On the third day after dialysis, the supernatants of centrifuged PMMA/DODAB dispersions contained 1.3 mM DODAB, suggesting that 0.7 mM DODAB was still incorporated by the PMMA/DODAB NPs. PMMA/CTAB NPs dispersions had contents of QAC determined after dialysis and centrifugation, revealing the absence of CTAB in the supernatants just after dialysis but its presence in the supernatant 3 days after dialysis showing its low affinity for the PMMA/CTAB NPs. In contrast, 3 days after dialysis, 0.5 mM CTAB was determined in the supernatant of PMMA/CTAB NPs showing CTAB leakage from the PMMA/CTAB NPs just after dialysis (Table 5). In summary, DODAB incorporation in the NPs was substantial, whereas the one of CTAB was transient and possibly almost lost after dialysis.

The evaluation of inhibition halos by CTAB against bacteria and yeast is in Figure 4. Just after dialysis, CTAB was not found in the supernatants of PMMA/CTAB NPs. This contrasted with 0.5 mM CTAB found in the supernatant 3 days after dialysis (Table 5). PMMA/CTAB NPs behaved as a reservoir for the release of CTAB with time after dialysis. The experiment using CTAB to determine the inhibition halos also showed that CTAB is able to move through the agar to inhibit microbial growth. Due to this property, inhibition halos against seeded bacteria and fungus could be determined on Petri dishes as a function of [CTAB] over a range of [CTAB] (0.01–2.5 mM CTAB) (Figure 4). Inhibition halos occurred for PMMA/CTAB NPs before (B) dialysis but did not occur just after dialysis (A), confirming the momentaneous absence of CTAB in the NPs supernatant. Estimated [CTAB] in the NPs' supernatant before dialysis resulted from the similarity with halo 3 against *S. aureus* (Figure 4b) or halo 3 against *C. albicans* (Figure 4c), yielding 0.3 mM CTAB outside the NPs. If the added CTAB during particle synthesis was 2 mM, after synthesis, 1.7 mM was incorporated in the NPs and 0.3 mM was in the outer solution. However, the NPs dialysis after synthesis eliminated non-incorporated CTAB from the dispersions, as depicted from the absence of halo in A (Figure 4). Regarding the antimicrobial activity that will be determined just after dialysis, one will have to consider 1.7 mM as the CTAB concentration in the NPs.

**Figure 4.** Figure 4. a b c (**a**) *E. coli*, (**b**) *S. aureus*, and (**c**) *C. albicans* inhibition halos induced by CTAB alone (numbered from 1 to 8) or by PMMA/CTAB dispersions after (A) or before dialysis (B). From 1 to 8, [CTAB] inside the wells was 0.01, 0.2, 0.3, 0.5, 1.0, 1.5, 2.0, and 2.5 mM.

For PMMA/DODAB NPs, the experiment based on inhibition halos was not possible since DODAB BF are not able to diffuse in the agar medium as reported before [23].

#### *3.4. Antibacterial and Antifungal Activity of QACs and PMMA*/*QAC NPs*

A proper evaluation of the antimicrobial activity involves determining CFU counting over a range of [QAC] and expressing the CFU countings on a logarithmic scale so that the effective potency of the QACs becomes evaluated over ample range of magnitude.

Figure 5 shows the cell viability of *Escherichia coli* in the presence of CTAB (a), PMMA/CTAB NPs (b), DODAB BF (c), and PMMA/DODAB NPs (d). Just after dialysis, the inhibition halo experiment yielded an absence of free CTAB in the supernatant of PMMA/CTAB NPs and 1.7 mM CTAB in the NPs. This [CTAB] initially incorporated in the NPs decreased *E. coli* viability by 1.5 logs (Figure 5b), in contrast to free CTAB that reduced viability by 7 logs (Figure 5a). CTAB exhibited a remarkable activity against *E. coli* that could not be identified before from experiments expressing only % cell viability (limited to only 2 logs at most). CTAB incorporation in the PMMA/CTAB NPs reduced substantially its activity but may be useful for controlled release in biomedical prosthetic devices [63] or agar-based hydrogels [64].

Figure 5. a b c d **Figure 5.** Viability of *Escherichia coli* in the presence of CTAB (**a**), PMMA/CTAB (**b**), DODAB (**c**), and PMMA/DODAB dispersions (**d**). The NP dispersions of PMMA/CTAB and PMMA/DODAB were used after dialysis. The counting of colony-forming unities (CFU) was expressed on a logarithmic scale.

The CTAB mechanism of action involves reaching the bacterial cell membrane causing its disruption and cell lysis [65]. Therefore, the large reduction in CTAB activity was due to its location in the PMMA/CTAB NPs instead of moving freely in the bulk solution to interact with the bacteria. In another report, CTAB adsorbed by BiOBr nanosheets showed lower toxicity than the same level of free CTAB, which was attributed to the adsorption or hindering effect of BiOBr nanosheets [66]. PMMA/CTAB NPs in this work released 0.5 mM CTAB on the third day after dialysis from an initial concentration in the NPs of 1.7 mM CTAB (Table 5; Figure 4). This behavior was similar to the one of the inorganic BiOBr nanosheets incorporating CTAB that exhibited a slow and sustained release of CTAB or benzalkonium chloride for8h[66].

Among the biocompatible polymers, PMMA was used in combinations with CTAB or DODAB to prepare spin-coated films able to kill bacteria upon release to the medium (CTAB) or upon contact on the surface of the coating (DODAB) [19]. In pharmaceutics, polymers are often be used as reservoirs of the active principle or drug so that the polymer can control the release of the active molecule over time in vivo [3,67,68].

DODAB BF reduced the viability of *E. coli* by 2 logs (Figure 5c), whereas PMMA/DODAB NPs reduced viability by 0.5 logs CTAB (Figure 5d). Possibly, the lack of mobility of the DODAB BF across the bacterial cell wall to reach the cell membrane resulted in the much lower activity of DODAB in comparison to the one of CTAB. In fact, no leakage of intracellular contents was detected for DODAB BF interacting with *E. coli* in comparison to the pronounced leakage determined for CTAB and the anionic sodium dodecylsulfate (SDS) [29].

The high affinity of DODAB for PMMA determined its incorporation in the NPs, which was 0.7 mM remaining 1.3 mM in the supernatant after dialysis and centrifugation (Table 5). For the PMMA/DODAB NPs, the activity was lower than the one of DODAB BF (Figure 5c,d). A possible explanation for this is that the DODAB incorporated in the NPs was not leaving them to kill the microbia; this meant that only DODAB BF outside the NPs was effective.

Figure 6 shows the compared activity of CTAB (Figure 6a) and PMMA/CTAB NPs (Figure 6b) against *S. aureus*. Similar to the effect of PMMA/CTAB NPs against *E. coli*, before dialysis there was

0.3 mM CTAB outside the PMMA/CTAB NPs with 1.7 mM incorporated in the NPs. The CTAB reservoir effect of the NPs reduced its effect against the bacteria, as compared to free CTAB. The compared effect for DODAB BF (Figure 6c) and PMMA/DODAB NPs (Figure 6d) revealed a similar and low activity. *Staphylococcus* sp. developed a sensor system for cationic antimicrobial peptides based on a sensor consisting of a short and negatively charged extracellular loop of amino acid residues able to interact with cationic antimicrobial peptides [69]. The transduction of this interaction signal would trigger the d-alanylation of teichoic acids and the lysylation of phosphatidylglycerol, resulting in a decreased negative charge of the bacteria. It is possible that DODAB BF (but not CTAB molecules) were possibly able to trigger this resistance mechanism in *S. aureus* due to the multipoint attachment of DODAB BF to the negatively charged loop. CTAB, on the other hand, would kill microorganisms as individual molecules below its critical micellar concentration (1 mM) [28,30], rapidly and extensively penetrating the cell wall to reach the cell membrane.

Figure 6. a b c d **Figure 6.** Viability of *Staphylococcus aureus* in the presence of CTAB (**a**), PMMA/CTAB (**b**), DODAB (**c**), and PMMA/DODAB dispersions (**d**). The NP dispersions of PMMA/CTAB and PMMA/DODAB were used after dialysis. The counting of colony-forming unities (CFU) was expressed on a logarithmic scale.

 Against *Candida albicans*, free CTAB showed remarkable activity (Figure 7a) in contrast with the reduced activity of PMMA/CTAB NPs (Figure 7b). The NP dispersion just after dialysis was used to interact with the fungus so that no CTAB molecules were available outside the NPs to interact with the cells. However, some CTAB leakage from the NPs was probably taking place to yield the 1.5-logs reduction observed in Figure 7b. On the other hand, DODAB BF was surprisingly active against *C. albicans*, resulting in a reduction of ca. 5 logs at 1.2 mM DODAB after a 1-h interaction (Figure 7c). Neither Campanhã and coworkers [40] nor Vieira and coworkers [31] could observe the good antifungal activity of DODAB BF since they employed percentiles of viable cells to express the viability curves as a function of DODAB concentration. Consistently, Fukushima and coworkers [70] also reported a superior activity of supramolecular assemblies made of poly(lactide) (interior block) and cationic polycarbonates (exterior block); upon testing the spherical and rod-like morphologies for antimicrobial properties, they found that only the rod-like assemblies were effective against *Candida albicans*. This showed that the shape of the antimicrobial supramolecular assembly was important to determine antifungal activity. In the present case, the disk-like shape of the DODAB BF assemblies resulted in good antifungal activity that was not described before (Figure 7c). This can possibly be ascribed to the

improved penetration of rod-like or disk-like nanostructures through the outer layer of glicoproteins on the fungus cell wall as compared to vesicles, liposomes, or nanoparticles [42,52,71].

Figure 7. a b c d **Figure 7.** Viability of *Candida albicans* in the presence of CTAB (**a**), PMMA/CTAB (**b**), DODAB (**c**), and PMMA/DODAB dispersions (**d**). The NP dispersions of PMMA/CTAB and PMMA/DODAB were used after dialysis. The counting of colony-forming unities (CFU) was expressed in a logarithmic scale. The superior activity of QACs by themselves as compared to QACs in NPs is depicted from comparisons between (**a**,**b**) or (**c**,**d**). CTAB was active as a monomer at submicellar concentrations (**a**). DODAB was active as bilayer fragments (**c**).

In Figure 7d, the reduced activity of PMMA/DODAB NPs in comparison to the one of DODAB BF (Figure 7c), again, can be understood from the incorporation of DODAB in the NPs reducing its availability to interact with the fungus; there was 1.3 mM DODAB BF outside the NPs, whereas, in the DODAB BF, dispersion was 2.0 mM DODAB (Figure 7c; Table 5).

In Figure 8, the cell viability of *Candida albicans* as a percentile of viable cells (%) was obtained as a function of CTAB (Figure 8a) or DODAB concentration (Figure 8b) and compared with previous data from the literature [31]. There was a dependence of the viability curve on the initial concentration of viable cells: around 10<sup>6</sup> CFU.mL<sup>−</sup>1, the sigmoidal curve occupied a range of lower QAC concentrations than the one around 10<sup>7</sup> CFU.mL−1. This was consistent with the lower amounts of QAC required to kill lower concentrations of cells. In addition, one should notice that achieving a reduction of two logs in viable cells counting did not allow to discriminate whether the effect of the QACs reduced the counting by more than 2 logs. It is necessary to express CFU counting on a logarithmic scale in order to determine the complete effect of any antimicrobial, as done against *C. albicans* in Figure 7a,c. Consequently, the CTAB and DODAB BF effects were fully revealed, showing important reductions in CFU countings over 7 and 5 logs, respectively (Figure 7a,c).

Regarding the PMMA/CTAB NPs, the percentiles of viable countings plotted as circles in Figure 8c did not reveal the 5-logs reduction observed against *S. aureus* in Figure 6b. Only in the case of a small antimicrobial effect were the percentiles of viable cells sufficient to determine antimicrobial activity as was the case of the 1.0-logs reduction caused by PMMA/DODAB NPs in Figure 8d corresponding to Figure 6d expressed on a logarithmic scale.

Figure 8. (a) (b) (c) (d) **Figure 8.** *Candida albicans* viability (%) (-) over a range of CTAB (**a**) or DODAB concentrations (**b**) in QAC dispersions in water as compared to data from [31] (-). *Staphylococcus aureus* viability (%) (-) over a range of CTAB (**c**) or DODAB concentrations (**d**) in the presence of PMMA/QAC NPs, as compared to data from [27] (-). QAC dispersions in water are CTAB micelles or DODAB BF. In (**c**,**d**), PMMA/QAC NPs were obtained by the emulsion polymerization of MMA in the presence of QACs. These NPs were obtained either using AIBN (-) or KPS (-) as the initiator. [MMA] and [QAC] used during particle synthesis were 0.56 M and 4.0 mM, respectively, for squares in (**c**), 0.4 M and 2.0 mM, respectively, for circles in (**c**,**d**), and 0.56 M and 10 mM, respectively, for squares in (**d**).

#### 4. Conclusions **4. Conclusions**

Over a range of MMA or QAC concentrations, 0.4 M MMA and 2 mM QAC yielded optimal physical properties for the PMMA/DODAB and PMMA/CTAB NPs in dispersion, such as nanometric size (Dz < 100 nm), low polydispersity (<0.05), high and positive zeta-potential (>20 mV), high yield (>70%), high particle number density (>1013 particles.mL<sup>−</sup>1), and high colloidal stability (absence of aggregates and precipitates). Among two initiators (KPS or AIBN), AIBN was the best one to obtain optimal properties for the NPs synthesized at 2 mM QAC and 0.56 M MMA; the low sizes, high positive zeta-potentials, and low polydispersities occurred only using AIBN.

Inhibition halos by CTAB and PMMA/CTAB NPs against bacteria and yeast showed that PMMA/CTAB NPs behaved as reservoirs for the release of CTAB with time after dialysis; CTAB was able to move both through the dialysis membrane and the agar to inhibit microbial growth, highlighting hydrogels as good vehicles for CTAB. In contrast, DODAB preferred to remain incorporated in the PMMA polymeric matrix, did not move in the agar, and did not cross the dialysis bag membrane; for PMMA/DODAB NPs or DODAB BF, inhibition halos were not observed due to the lack of DODAB BF diffusion in the agar medium.

The incorporation of QACs in the PMMA/QAC NPs reduced antimicrobial activity in comparison to the QACs dispersions. CTAB was the most active microbicidal agent against the three microbes tested (*E. coli*, *S. aureus*, and *C. albicans*), reducing cell viability countings by 7 logs at submicellar concentrations. The controlled release of CTAB from PMMA/CTAB NPs, however, would be a promising strategy for CTAB delivery. CTAB mobility in hydrated medium favored its electrostatic interaction and penetration through the microbes' cell wall and membrane, imparting a lytic effect on the cells. DODAB BF revealed an important fungicidal activity against *C. albicans* not described before; similar to rod-like, cationic supramolecular assemblies, the disk-like cationic DODAB BF possibly entered the outer glycoproteins layer of the fungus penetrating the cell and causing a 5-logs reduction in yeast viability.

The real potency of QACs antimicrobials should not be evaluated in percentiles of initial CFU counting; potent agents often display reduction in microbial cell viability larger than the 2 logs of the initial counting of viable cells.

The PMMA/QAC NPs here described still require further evaluation regarding their cytotoxicity against mammalian cells before fulfilling their potential in drug and vaccine delivery. Due to their nanometric size, positive zeta-potential, narrow size distribution, and high colloidal stability, they are expected to combine well with oppositely charged antigens (proteins, peptides or DNA) for subunit vaccines' design, as many cationic adjuvants do. CTAB leakage and lytic property against eukaryotic cells, such yeast *C. albicans*, would not recommend the PMMA/CTAB NPs for vaccines. DODAB, on the other hand, remains embedded in the PMMA matrix, imparting a positive charge to the NPs without leaking to the outer medium. These considerations suggest that PMMA/DODAB NPs should be kept in perspective for further testing and prospective uses in drug and vaccine delivery.

**Author Contributions:** Conceptualization, A.M.C.-R.; Data curation, A.M.C.-R.; Formal analysis, B.I.M., A.M.C.-R.; Funding acquisition, A.M.C.-R.; Investigation, B.I.M., A.M.C.-R.; Project administration, A.M.C.-R.; Resources, A.M.C.-R.; Supervision, A.M.C.-R.; Writing—original draft, B.I.M., A.M.C.-R.; Writing—review & editing, A.M.C.-R. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research and the APC were funded by Conselho Nacional de Desenvolvimento Científico e Tecnológico (CNPq), grants 302758/2019-4 and 302352/2014-7. B.I.M. was the recipient of undergraduate fellowships of the Programa Unificado de Bolsas da Universidade de São Paulo granted to the Project "Cationic Supramolecular Assemblies and their Films" by A.M.C.-R.

**Acknowledgments:** The technical support of Rodrigo Tadeu Ribeiro is gratefully acknowledged.

**Conflicts of Interest:** The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in the decision to publish the results.

#### **References**


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