E-ATTA (Additional file 1: Fig. S1a): 3363 cm−1 (O–H stretching), 3148 cm−1 (N–H stretching vibrations), 2927 cm−1 (C-H stretching vibrations (aliphatic)), 1708 cm−1 (C=O stretching vibrations (acid)), 1686 cm−1 (C=O stretching (amide)), 1560 cm−1 (C=N stretching vibrations), 1452 cm−1 (HC=CH stretching vibrations), 1309 cm−1 (C–O stretching vibrations), 1088 cm−1 (S–C–S stretching vibrations). Cs (Additional file 1: Fig. S1b): 3400–3460 cm−1 (O–H & N–H stretching (amide II), and intra-molecular hydrogen bond of –OH⋯O on saccharine ring), 2929 cm−1 and 2880 cm−1 (C–H, asym. and sym. stretching vibrations), 1659 cm−1 (C=O stretching vibration (amide I) and C=O⋯H–N hydrogen bond), 1596 cm−1 (N–H bending vibration (amide II)), 1430 cm−1 (-CH2– bending and rocking vibrations), 1380 cm−1 (–CH– asymmetric bending vibration), 1080 cm−1 and 1030 cm−1 (C–O stretching vibrations), and 895 cm−1 (stretching vibration of saccharine ring) [53, 54]. Cs-EATT (Additional file 1: Fig. S1c): 3441 cm−1 (O–H and N–H stretching vibration), 2924 cm−1 (C–H stretching vibration (aliphatic)), 1726 cm−1 (C=O stretching vibration (amide)), 1714 cm−1 (C=O stretching vibration (amide)), 1632 cm−1 (C=N stretching vibration), 1555 cm−1 (HC=CH stretching vibration), 1083 cm−1 (S–C–S stretching vibration), 1125 cm−1 (C–O–C asymmetric stretching vibration of glucosamine), and 1064 and 1023 cm−1 (C–O stretching vibration of glucosamine).
B-ATTA (Additional file 1: Fig. S2a): 3605 cm−1 (O–H stretching), 3201 cm−1 (N–H stretching), 3059 cm−1 (C–H stretching (aromatic)), 2971 cm−1 (C–H stretching (aliphatic)), 1693 cm−1 (C=O stretching (acid)), 1678 cm−1 (C=O stretching (amide)), 1572 cm−1 (C=N stretching), 1553 cm−1 (C=C stretching), 1300 cm−1 (C–O stretching), 1053 cm−1 (S–C–S stretching). Cs-EATT (Additional file 1: Fig. S2c): 3415 cm−1 (O–H and N–H stretching vibration), 3001 cm−1 (C–H stretching vibration (aromatic)), 2928 cm−1 (C–H stretching vibration (aliphatic)), 1796 cm−1 (C=O stretching vibration (amide)), 1733 cm−1 (C=O stretching vibration (amide)), 1632 cm−1 (C=N stretching vibration), 1538 cm−1 (C=C stretching vibration), 1133 cm−1 (C–O–C asymmetric stretching vibration of glucosamine), 1070 cm−1 (S–C–S stretching vibration), and 1053 and 1011 cm−1 (C–O stretching vibration of glucosamine).
1H & 13C NMR and mass analysis
E-ATTA (Additional file 1: Fig. S3a,b): 12.61 (s, 1H, O–H, exchangeable by D2O), 12.23 (s, 1H, NH, exchangeable by D2O), 3.17 (q, 2H, CH3CH2–S–), 2.66 (t, 2H, CH2-COOH), 2.53 (t, 2H, CH2–CONH–) and 1.3 (t, 3H, CH3CH2–S–).13C-NMR (Additional file 1: Fig. S3c): 173.56 (C=O (acid)), 170.80 (C=O (amide)), 158.61 (–S–C–S), 158.45 (–S–C–NH), 29.89 (–CH2–COOH), 28.37 (CH3CH2–S–), 28.15 (–CH2–CONH–) and 14.80 (CH3CH2–S–). The molecular formula of E-ATTA was further confirmed by mass spectrometry analysis (Fig. 1) showing the peak at m/z 261 (2.62%) corresponding to the molecular ion, which losses H2O, CH2=CH2, CO, HC≡CH, CO, S, and HCN successively to produce ion fragment peaks at 243.01 (17.17%), 215 (13.29%), 187 (3.84%), 161 (7.68%), 133 (21.39%), 101 (9.2%), and 74 (18.31%), respectively as shown in Fig. 1, and the base peak was observed at 55 (100%) correspond to vinyl carbonyl (C3H3O+) fragment ion.
B-ATTA (Additional file 1: Fig. S4a): 12.62 (s, 1H, O–H), 12.23 (s, 1H, NH), 7.39–7.24 (m, 5H, Ar–H), 4.46 (s, 2H, Ph–CH2–S–), 2.67 (t, 2H, CH2–COOH) and 2.54 (t, 2H, CH2–CONH–). 13C-NMR (Additional file 1: Fig. S4b): 173.51 (C=O (acid)), 170.81 (C=O (amide)), 158.92 (–S–C–S), 157.88 (–S–C–NH), 136.77,129.01, 129.01, 128.61, 128.61, 127.63 (aromatic carbons), 37.61 (CH2–S–), 29.83 (–CH2–COOH) and 28.30 (–CH2–CONH–). Further, the resulting compound was confirmed by mass spectrum (Fig. 1) which showed the molecular ion peak at m/z = 323 (0.45%) which losses H2O, C6H4, CH2, CO, HC≡CH, CO, S, and HCN successively to produce ion fragment peaks at 305 (13.3%), 229 (0.05%), 215 (0.13%), 187 (0.06%), 161 (0.07%), 133 (0.30%), 101 (0.49%), and 74 (0.52%), respectively, and the base peak observed at m/z = 91 (100%) which attributed to tropylium ion.
Cs-EATT and Cs-BATT (Fig. 2a, b): configuration of Cs-EATT was as well, confirmed by the Nuclear magnetic resonance technique. 1H-NMR of chitosan  elucidated the signals at (δ ppm): 1.32 ppm integrating for three protons of the methyl of the N-acetyl group in the Cs. In addition, the signals of H2-H2′ in the glucosamine unit are at 2.80 ppm and 3.36 ppm (3.28–3.40 ppm). The numerous signals in the 3.6–4.15 ppm range due to the hydrogens of H3-H6 and the signals at 4.76, 5.11 ppm ascribe to the hydrogens of H1-H1′ in the glucosamine moiety. As in Fig. 2a, the new signals at 1.17 ppm, 2.55 ppm, 2.68 ppm, 3.21 ppm, 9.83 ppm, and 12.64 ppm which were attributed to the resonance of the three protons of (CH3CH2–S), two protons of (CH2–CONH–), two protons of (CH2-COOH), two protons of (CH3CH2–S–) and two protons of (2NH) respectively, confirmed the formation of Cs-EATT. Also, NMR-d6 of Cs-BATT (Fig. 2b), the absence of the singlet signal corresponding to O–H proton of acid, and the appearance of a new N–H signal at δ 9.17 ppm indicate the formation of the modified chitosan, with other requisite numbers of protons in its 1H-NMR spectrum, supported the formation of the assigned structure as shown in Fig. 2b.
The elemental analysis results for Cs-EATT and Cs- BATT were presented in Table 2 and used to determine the degree of substitution (DS) according to Eq. 7 .
where (C/N)b is the carbon/nitrogen ratio of Cs-EATT and Cs-BATT, (C/N)b is the carbon/nitrogen ratio of the chitosan. x and y are the numbers of nitrogen and carbon atoms, respectively, introduced into chitosan after modification with E-ATTA and B-ATTA compounds. The degree of substitution (DS) value was observed to be 84.7% for Cs-EATT and 44.1% for Cs- BATT.
The TGA was conducted to investigate the thermal behavior of the two functionalized chitosan, Cs-EATT and Cs-BATT, and the data obtained were tabulated in Table 3 on the basis of the plot of mass loss (%) versus temperature, as presented in Fig. 3. From the figure, both samples undergo a decrease in mass as the temperature increases. Meaning that the stability of both samples is temperature-dependent. Comparing the data presented in Table 3 indicates that the thermal stability of Cs-BATT is higher than that of Cs-EATT. This may be due to the presence of phenyl groups in Cs-BATT which restrict the mobility of the chains, hence the stability increases. The initial loss in the sample mass with temperature corresponds to the desorption of the moisture. The mass loss propagation is due to the decomposition of the heterocyclic moieties followed by the decomposition of the d-glucosamine rings. As shown in Table 3, the T50 (The temperature at which the sample losses half of its mass) of Cs-EATT is lower than that of Cs- BATT by 67 °C. This confirms the result that the thermal stability of Cs-BATT is higher than that of Cs-EATT.
Recently, studies of functionalized chitosan with new active compounds have increased to provide a wide range of biomedical and biotechnological chitosan applications [35, 56, 57]. The chemical modifications of chitosan to form new functional characteristics are important steps to overcome the low solubility of chitosan that hinders its applications . The solubility of chitosan was increased in the presence of hydrophilic thiadiazole derivatives, as reported previously . Therefore, the antibacterial and antifungal activities of newly synthesized functionalized chitosan with ethyl and benzyl thiadiazole, Cs-EATT, and Cs-BATT were evaluated against Bacillus subtilis, Staphylococcus aureus, Escherichia coli, Pseudomonas aeruginosa, and Candida albicans by the agar well diffusion method. The DMSO (solvent system) did not exhibit any antimicrobial activity against any of the tested bacterial or fungal strains. For B. subtilis, the differences between the zone of inhibition formed due to treatment with 300 µg mL–1 are non-significant between Cs-EATT (19.8 ± 0.3 mm) and Penicillin (19.7 ± 0.6 mm), while it was highly significant (p ≤ 0.001) as compared with the zone formed due to Cs-BATT (14.7 ± 0.7 mm) (Fig. 4a). In contrast, the zones of inhibition formed due to the treatment of S. aureus, P. aeruginosa, and C. albicans with the highest concentration (300 µg mL–1) of functionalized Cs-EATT and Cs-BATT represent a significant difference. Data analysis showed that the activity of Cs-EATT was higher than that of Cs-BATT against all tested organisms except C. albicans (Fig. 4e). This phenomenon can be attributed to the presence of thiadiazole attached to the benzyl moiety. The current study is in harmony with those recorded by Li and co-author , who showed that the highest activity of functionalized chitosan with thiadiazole, methyl thiadiazole, and phenyl thiadiazole against three phytopathogenic fungi was achieved at a concentration of 1000 µg mL–1. The authors attributed the activity of functionalized chitosan against phytopathogens to the presence of different thiadiazole derivatives.
To integrate the synthesized chitosan derivatives into various biomedical applications, they should be able to detect the lowest concentration that inhibits the growth of pathogenic microbes, which is known as a minimum inhibitory concentration (MIC). Therefore, the efficacy of different concentrations (200, 100, 50, 25, and 12.5 µg mL–1) of two synthesized polymers, Cs-EATT and Cs-BATT, to control the growth of pathogenic Gram-positive bacteria, Gram-negative bacteria, and C. albicans was investigated. Data analysis showed that the activity of synthesized polymers is dependent on the concentration used. The activity decreases as the concentration is decreased. At 200 µg mL–1, the zone of inhibition was decreased to (16.6 ± 0.6, 13.0 ± 0.0, 15.0 ± 0.0, 15.3 ± 0.6, and 15.9 ± 0.1 mm), (16.0 ± 0.9, 12.6 ± 0.3, 12.5 ± 0.5, 12.9 ± 0.2, and 12.9 ± 0.1), and (12.7 ± 0.6, 9.7 ± 0.1, 11.1 ± 0.2, 11.5 ± 0.5, and 14.3 ± 0.6 mm) for positive control, Cs-EATT, and Cs-BATT against B. subtilis, S. aureus, P. aeruginosa, E. coli, and C. albicans, respectively. For Gram-positive bacteria, Cs-EATT was more active at low concentration as compared with positive control and Cs-BATT (Fig. 4a and b), whereas the functionalized chitosan with benzyl thiadiazole (Cs-BATT) was more active against C. albicans at low concentration as compared with control and Cs-EATTA (Fig. 4e). The MIC values for positive control against B. subtilis, S. aureus, P. aeruginosa, E. coli, and C. albicans were 50, 50, 25, 25, and 50 µg mL–1, respectively, whereas they were 25, 25, 50, 50, and 100 µg mL–1 for Cs-EATT and 100, 200, 100, 100, and 25 µg mL–1 for Cs-BATT with varied clear zones (Fig. 4a–e).
In our recent study, the functionalized chitosan with aminothiazole and imidazole carboxamide groups showed antimicrobial activity against Gram-positive bacteria (B. subtilis and S. aureus), Gram-negative bacteria (P. aeruginosa and E. coli), and C. albicans with MIC values ranging between 50 and 100 µg mL–1 . The antimicrobial activity of functionalized chitosan with active moieties was higher than that recorded by non-functionalized chitosan, as reported previously. For instance, chitosan loaded with ZnO nanoparticles showed stronger antimicrobial activity against E. coli and S. aureus than the activities recorded by chitosan without modification . Moreover, the activity of composite magnetite/guar gum/chitosan to inhibit the growth of Gram-positive and Gram-negative bacteria was higher than that recorded by non-magnetite ones .
As reported previously , chitosan is characterized by its broad-spectrum activity against different pathogenic microbes, and their activity is increased by grafting with thiadiazole and other active moieties. The inhibitory effect of functionalized chitosan can be attributed to the various hypothesis, one of these hypotheses is related to cationic nature. The chitosan with a low molecular weight can easily penetrate the microbial cell walls and hence react with DNA and ultimately block the transcription process . Whereas chitosan with a high molecular weight has the efficacy to bind with cell wall components with a negative charge and form an impermeable layer surrounding the cell, which leads to a change in selective permeability function . The grafting of chitosan with hydrophilic thiadiazole increases the solubility of chitosan and hence increases the electrostatic attraction between functionalized chitosan and the bacterial cell wall, which enhances cell mortality . Another important inhibitory mechanism can be due to the production of reactive oxygen species as a result of accumulating the functionalized chitosan into the microbial cells and hence increasing cell mortality [66, 67]. Due to the disruption of the sterol profile existing in C. albicans cell wall because of negative impacts of treatment on the ergosterol synthesis pathway is considered another inhibitory mechanism of unicellular fungi .
Antimicrobial activity comparison study
Various attempts have been accomplished to investigate the antimicrobial activity of chitosan after modification or functionalization with various derivatives. Among these derivatives, are hydrophilic and hydrophobic thiadiazole (Table 4). For instance, chitosan was modified with different thiadiazole derivatives, including 1,3,4-thiadiazole; 2-methyl-1,3,4-thiadiazole, and 2-phenyl-1,3,4-thiadiazole and exhibited high growth inhibition percentages of different phytopathogenic fungi . The highest growth inhibitions were achieved for modified chitosan with 2-methyl-1,3,4-thiadiazole with percentages of 75.3, 82.5, and 65.8%, respectively, against Colletotrichum lagenarium, Phomopsis asparagi, and Monilinia fructicola. Moreover, the antimicrobial activity of modified chitosan with 5-amino-1,3,4-thiadiazole-2-thiol; 5-phenyl-1,3,4-oxadiazole-2-thiol; and 5-(4 chlorophenyl)-1,3,4-thiadiazole-2-thiol against Gram-positive bacteria, Gram-negative bacteria, unicellular fungi, and multicellular fungi was higher than virgin chitosan . In the current study, a new functionalized chitosan polymer with ethyl/benzyl thiadiazole derivatives showed high antibacterial activity against S. aureus, B. subtilis, E. coli, and P. aeruginosa as well as anti-Candida activity. Interestingly, there is no significant difference between the activity of modified chitosan and the positive control, which indicates the promising activity of modified chitosan to control the growth of pathogenic microbes.
Detection of λmax for Cs-E/BATT
The wavelength of the prepared Cs-E/B-ATT was scanned at a range of 200–400 nm. The obtained data showed that the maximum absorbance of prepared Cs-EATT and Cs-BATT was observed at λmax of 285 nm and 280 nm respectively.
Evaluation the quality of the prepared film dressing
The main criteria that confirm the successful formation of films are transparency, uniformity, homogeneity, elasticity, and texture properties . At least one of these criteria is accomplished, as shown in Table 5.
Moreover, the thickness of the prepared film dressing was varied based on the modifications and film composition. As shown in Table 6, the thickness was varied in the ranges of 0.092 ± 0.01–1.12 ± 0.07 mm and 0.094 ± 0.01–1.06 ± 0.04 mm for Cs-EATT and Cs-BATT, respectively, according to the film forming agent used. The change in thickness of the prepared film dressing followed the order of PVA > CMC > HEC and this phenomenon is compatible with the published study . The weights of prepared film dressing were in the range of 4.5 ± 0.9–10.6 ± 1.2 mg and 4.3 ± 0.8–10.4 ± 1.3 mg for Cs-EATT and CS-BATT, respectively (Table 6). As shown, the highest weight was recorded for formulations of E1 and B1. The uniform distribution of the drug polymers and plasticizers is indicated by the weight uniformity of different films of the same batch. The folding endurance was correlated with the film weight. This means the folding endurance was increased by increasing the film weight. The maximum folding endurance was recorded for the formulations of E1 (84 ± 2) and B1 (93 ± 3) and this is due to their highest weights (Table 6). Tensile strength is used to investigate the mechanical properties of the prepared film dressing. Data showed that the highest tensile strength was recorded for the formulations of E1 and B1 due to the presence of polyvinyl alcohol as a film-forming agent. The obtained data are compatible with those recorded by Asrofi and co-authors, who reported that the tensile strength of film-forming using PVA as the film-forming agent was higher than those formed using bengkuang starch and this attributed to the high crystallinity of PVA . The PVA tends to form an intramolecular network between its chains, hence producing good mechanical characteristics .
The moisture percentage loss was in the range of 7.7 ± 0.3, 11.2 ± 0.9, and 12.6 ± 0.5% for formulations E1, E2, and E3, respectively, whereas it was 6.8 ± 0.6, 9.5 ± 0.7, and 14.3 ± 0.7% for formulations B1, B2, and B3, respectively. The decrease in moisture percentage loss was noted to follow the order of CMC > HEC > PVA formulations due to hydrophobic characteristics. On the other hand, the moisture percentage absorption was highly varied between formulations and was (12.3 ± 0.4, 17.3 ± 0.3, and 22.6 ± 1.2%) and (14.7 ± 0.7, 15.4 ± 0.7, and 20.9 ± 2.02%) for (E1, E2, and E3) and (B1, B2, and B3), respectively (Table 6). The moisture absorption of the films followed the order CMC > HEC > PVA formulations, and this is correlated with the hydrophilicity of the film forming agents. The main advantage of the different film dressings prepared in the current study was that their pH values were satisfactory (6.1–6.9) (Table 6), thereby avoiding the risk of skin irritation upon their applications. The drug content in each prepared film was assessed as shown in Table 6. The highest drug content was recorded for formulations E2 and B2 with percentages of 92.5 ± 26.5% and 94.9 ± 27.4% respectively, whereas the lowest drug content was recorded for formulations E1 (85.9 ± 16.1%) and B1 (86.7 ± 16.5%). These variations can be attributed to the film-forming agent used .
In-vitro drug release studies
The in-vitro release of Cs-E/B-ATT for the different formulas of (E1, E2, and E3) and (B1, B2, and B3) films was investigated using the dialysis membrane as the released medium. As shown in Fig. 5, the release of the Cs-E/B-ATT from its different formulae can be ranked in the following descending order; (E2 > E3 > E1) and (B2 > B3 > B1). It’s clear that the drug release varied from (83.88–93.2%) and (87.7–97.35%) for Cs-EATT and Cs-BATT, respectively. The maximum of Cs-E/B-ATT release was achieved from the HEC-based film (E2, B2). This phenomenon can be attributed to the molecular weight of both formulas (E2/B2) which is bigger than others, which leads to a higher release rate of the film dressing. An increase in polymer macromolecule cross-linking is stated to be a direct result of an increase in polymer molecular weight coupled by a decrease in polymer dissolution rate. As a result, the water and drug diffusion coefficients drop, and drug release decreases .
The in-vitro release studies of Cs-E/ B-ATT as topical film dressings were represented in Table 7. Various kinetic models, including zero-order (cumulative % drug release vs. time), first-order (log cumulative % drug remaining vs. time), and diffusion models (Figs. 6 and 7) were applied to obtain the best fit for the results. Data showed that the in-vitro release of Cs-E/ B-ATT films followed zero transport. Therefore, the release of drugs from the formulated films is controlled by the swelling of the polymer, followed by drug diffusion through the polymer and slow erosion of the polymer, representing constant drug release in zero-order kinetics from the wound dressing films. The zero-order release model represents ideal drug delivery to maintain constant drug release as in transdermal patches and matrix tablets .
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