An adapted methodology was used for the synthesis of N,O-CMCS by alkalization followed by carboxylation [45]. CS (2.00 g) was mixed in 20 mL of a sodium hydroxide (20%) solution and stirred at room temperature (RT) for 12 h. The resulting CS of higher degree of deacetylation was separated by filtration. Next, 15 mL of isopropanol was added to the previously treated CS. Then, monochloroacetic acid (1.43 g) dissolved in isopropanol (10 mL) was slowly added dropwise into the treated CS isopropyl alcohol solution. The mixture was agitated for an additional 30 min. The flask was then put into a heated oil bath and stirred for 4 h. Then, the content was poured into a 50 mL beaker. Under continuous stirring condition, acetic acid (50%) was slowly added until the pH value reached 9. The product was in its salt-based form (N,O-CMCS-Na) due to the alkaline reaction medium. Finally, the reaction mixture was filtered, and the solid product was rinsed 3 times with a 200 mL ethanol solution, then once with absolute ethanol. The resulting solid was subsequently dried in an oven (50˚C) for three days. After, 1 g of N,O-CMCS-Na was suspended in 80% ethyl alcohol aqueous solution (100 mL), and then 10 mL of hydrochloric acid (37%) was added dropwise and stirred for 30 min to get the neutralized form. The resulting solid N,O-CMCS was filtered and rinsed using ethanol (70% - 90%) prior being dried out under vacuum overnight at RT [20].

The Fourier transformed infrared (FT-IR) and proton nuclear magnetic resonance (¹H-NMR) spectroscopy were used to confirm the addition of carboxymethyl groups on the CS amino and primary hydroxyl sites of the CS. Analyses were performed on an FT-IR spectrometer (FTIR Thermo iS10) at wavenumbers ranging from 600 to 4000 cm⁻¹. ¹H-NMR spectra of CS were obtained in D₂O, and N,O-CMCS in D₂O/HCl (100:1 v/v) using an NMR spectrometer (OXFORD NMR) under a static magnetic field of 200 MHz.

To determine the zeta potential (ZP), a 1 M (10 mL) solution of N,O-CMCS was dissolved in distilled water for 1 h with moderate shaking at room temperature. NaOH (0.01 M) was used to neutralize the pH of the solution. The zeta potential in solution was determined using a ZetasizerNano (Malvern Instruments Ltd., model ZEN 3600).

Stock solutions of N,O-CMCS and PEO were prepared in distilled water at three specific concentrations: 2.5, 3.3 and 8.0 wt% for N,O-CMCS, and 1.5, 3.0 and 8.0 wt% for PEO. To ensure a full and homogeneous dissolution, both solutions were kept under agitation at RT (e.g. 2 h, N,O-CMCS; 20 h PEO). Then, appropriate amounts of N,O-CMCS and PEO were mixed at various ratios to prepare electrospinning solutions. Electrospinning solutions used in this study are summarized in Table 1.

All electrospinning solutions were magnetically stirred for 2 h, then transferred in an ultrasonic bath for 15 min to remove air bubbles. Finally, the mixture was rested for 3 h before being used in the electrospinning setup. The N,O-CMCS/PEO solution was poured into an electrospinning plastic syringe or

kept in a refrigerator at 4˚C.

A syringe pump device (KD Science, model 100), a high-voltage power supply (Gamma High Voltage Research USA), and a metallic wireframe as a collector were part of the electrospinning system. For the conservation and stabilization of the membranes, two laboratory ovens (Fisher Scientific IsotempOven, Thermo Scientific HERAThermOven) were used. A schematic diagram of the electrospinning set-up used for the membrane preparation is depicted in Figure 1. Each electrospinning solution (Table 1) was poured into a 5 mL syringe mounted with a 20-gauge needle and attached to a syringe pump providing a slow and steady flow rate of liquid (0.4 mL/h). A distance (6 - 8 cm) was set between the tip of the needle and the collector. A metallic frame was used for the collector. The voltage used ranged between 10 and 15 kV. Various electrospinning conditions were attempted to obtain an optimal jet as well (stable with minimal drop projections) and beadless nanofibers mats by varying the flow rate, distance, and electrical current. Electrospinning was carried out in a tailor-made enclosure, at RT and relative humidity ranging between 30% - 50%. The nanofibrous mat was removed from the frame at the end of the electrospinning process and dried for 24 h in an oven at 80˚C.

N,O-CMCS/PEO nanofibers are soluble in water following the electrospinning process due to ionizable chemical functionalities of the material. After electrospinning, the nanofiber mat was dried overnight in an oven at 80˚C to remove residual solvent. A stabilization cycle was then completed by soaking resulting membranes in Na₂CO₃ (0.1 M) at RT for 3 h [43]. Finally, to improve the nanofiber mat stability in aqueous media, a heat treatment at 140˚C for 30 min was applied. The material stability was determined by an immersion in water for up to 6 h. Effects of water on nanofiber structure and morphology were studied by SEM images.

The degree of substitution (DS) of the sample was calculated using a potentiometric titration method [46]. Briefly, dried N,O-CMC (0.20 g) was dissolved in 0.1 M hydrochloric acid solution (20 mL). Methyl orange was used as indicator for the end-point determination. A standard 0.1 M sodium hydroxide solution was used during titration [31] [46]. The cumulative DS of the sample was determined using Equations (1) and (2):

DS = 161 A M CMCS − 58 A (1)

A = V NaOH ⋅ C NaOH (2)

where M_CMCS is the mass of CMCS (g), V_NaOH and C_NaOH are representing respectively the volume and molar concentration (M) of NaOH, and finally 161 and 58 are associated to the molecular weights of the chitosan (glucosamine) and the CMCS group [31].

Morphologies of both nanofibers CS/PEO and N,O-CMCS/PEO were assessed using a scanning electron microscope (SEM) (Hitachi SU1510 Scanning Electron Microscope). The average diameter of electrospun nanofibers was estimated using SEM images and Image J software. In order to obtain an average value, the diameter of 50 nanofibers collected on three separate images was calculated (e.g. total of 150 nanofibers per sample).

Some infrared (FT-IR) spectra analyses of the electrospun nanofiber mats were performed using a Fourier transform infrared spectrometer (Nicolet iS10, Thermo Scientific) in the 400 to 4000 cm⁻¹ wavenumbers range. In addition, ¹H-NMR spectra of CS and N,O-CMCS in D₂O/DCl were recorded using a 200 MHz NMR spectrometer (Varian).

The adsorption of FLX by the electrospun nanofibrous mat was carried out in batch mode. A 25 mg sample of the membrane was inserted into one Erlenmeyer flask containing 50 mL of an FLX solution (50 ppm) and 5% of methanol. The flask was then agitated on an orbital shaker (ORBIT Environ-shaker, Lab-Line) at RT. Before insertion of the membrane into the solution, an aliquot of 500 µL was sampled to determine the initial FLX concentration. Batch tests were conducted over a period of 150 min to ensure that equilibrium was achieved prior to the collection of a 500 µL aliquot. The adsorption capacity at time t (Q_t) was determined using the following Equation (3):

Q t = ( C 0 − C e ) V m (3)

where C_e is the concentration of the contaminant (ppm) at time t (min), C₀ is the initial concentration of contaminants (ppm), V is the volume of the solution (L), and m is the mass of adsorbents (g).

In order to determine the residual concentration of FLX in water samples during batch adsorption tests, a high-performance liquid chromatography (HPLC) system (Shimadzu Prominence I-series) coupled with a diode array detector (DAD) was used. The chromatographic separation was achieved using a C18 reverse-phase column (XB-C18, 100 Å, 150 × 3 mm, 2.6 μm particle size, Phenomenex, Kinetex®). The residual concentration of FLX in aqueous solution was determined by HPLC-UV DAD according to a method developed by Camiré et al. 2018 [44].

Kinetic curves were obtained by collecting medium samples (500 μL) at predetermined periods (0, 5, 10, 15, 20, 25, 30, 40, 50, 60, 75, 90, 120, and 150 min). Tests were carried out at RT with an initial FLX concentration of 50 ppm. Concentrations of FLX in aqueous phase were determined using an HPLC-UV DAD system. All tests were performed in triplicate to provide mean and standard deviation values.

Adsorption kinetic models are used to characterize the adsorption process (e.g. the movement of the adsorbate to the adsorbent’s external surface, the internal diffusion of the adsorbate to the active sites, and the real sorption to the adsorbent’s external surface) [47] [48]. Two models are mostly used in kinetics studies by researchers. Hence, it was decided to use nonlinear forms of pseudo-first-order (PFO) (Equation (4)) and pseudo-second-order (PSO) (Equation (5)) models in order to get the best-fit experimental data.

Q t = Q e ( 1 − exp − k 1 t ) (4)

Q t = k 2 Q e 2 t 1 + k 2 Q e t (5)

where Q_e is the amount adsorbed (mg/g) at equilibrium, Q_t is the amount adsorbed (mg/g) at time t (min), k₁ is the PFO adsorption rate constant (min⁻¹), and k₂ is the PSO adsorption rate constant (g/g min).

Kinetic models are typically used to examine the mechanism involved during the adsorption process, as well as the location at the surface of adsorption, the chemical reaction involved, and/or the processes of diffusion. As the process of prevalence, the PFO model assumes a physical adsorption, while a chemical adsorption is inferred by the PSO model [49]. Kinetic parameters (maximum adsorption potential and kinetic constant) were determined for both models using software: the kinetic equation solve feature for Microsoft Excel, and nonlinear regression analysis (MATLAB).

Infrared spectra of CS and N,O-CMCS are depicted in Figure 2. As shown, additional bands are well supporting the chemical structure of N,O-CMCS (DS 1.15). The broad band at 3301 cm⁻¹ (axial stretching of the O-H and N-H bonds secondary amine) is present in FT-IR spectra (Figure 2(b)); this evidence suggests the formation of the modified compound at NH position of the CS. In

addition, the stretching vibration of C-O located in CH₂COOH group can be associated to peaks 1250 cm⁻¹ and 1245 cm⁻¹. The peak at 2887 cm⁻¹ corresponds to the axial stretching of the C-H bonds, and the peak 1730 cm⁻¹ is related to the carbonyl group C=O from the newly formed COOH groups, which was not part of the initial CS spectrum. Bands at 1150 - 897 cm⁻¹ of C-O and C-O-C appear in all spectra. Characteristic bands that belong to CS are also identified: 3357 cm⁻¹ and 3287 cm⁻¹ (O-H and N-H bonds), 1650 cm⁻¹ and 1589 cm⁻¹ (NH₂ primary amino group of CS prior to its chemical modification to a secondary amine). Reported FT-IR peaks and bands confirming the carboxymethylation conversion of CS to N,O-CMCS are consistent with other published studies [37] [55].

The Zeta potential (ZP) is used to measure the effective electric charge on the membrane’s surface; it can provide useful information on possible electrostatic interactions between N,O-CMCS and FLX molecules. Based on the degree of

substitution (DS) of the N,O-CMCS calculated using a potentiometric or acid-base titration method, this measure provides information about the ionization state of the grafted carboxylic acid moieties on the surface of the modified polymer. Figure 4 shows the ZP (mV) as a function of pH. As the pH increased, the ZP is decreasing to negative values. This indicates that N,O-CMCS becomes highly ionized with negative charges due to larger amount of deprotonated carboxyl groups (-COO⁻). Under alkaline conditions, an electrostatic sorption mechanism is possible and favors the attraction of positive ions toward the negative surface of the modified biomaterial N,O-CMCS. However, at pH < 4.5 the - RNH 3 + group (from protonated secondary amine) is predominant along with positive ZP values. Around pH 4.5, an isoelectric point (pI) is generated by the presence of both charged species (e.g. - RNH 3 + /-RCOO⁻) on the surface. Similar observations are reported in literature [31] [47] [55].