Introducing the antibacterial and photocatalytic degradation potentials of biosynthesized chitosan, chitosan–ZnO, and chitosan–ZnO/PVP nanoparticles – Scientific Reports

Characterization techniques

UV–Vis spectroscopy

Figure 3 displays the results of the analysis of the UV–Vis absorption spectra of several materials, including chitosan, ZnO NPs, chitosan–ZnO NPs, and chitosan–ZnO/PVP. In the case of chitosan, two initial absorption band is observed at 270 nm (Fig. 3a1). This band is attributed to the n–π* transition of the amino group present in chitosan. This means that the electrons in the amino group undergo an electronic transition from the non-bonding (n) orbital to the anti-bonding (π*) orbital upon light absorption at 270 nm²⁶. Figure 3b1 represents the UV–Visible absorption spectrum of ZnO-NPs synthesized using of plant extract of Solanum nigrum L.²⁷. This spectrum showed the absorption maximum at 356 nm which is characteristic to ZnO NPs. The shape of the UV–Visible spectrum is quite similar to the spectrum reported in a previous study²⁸. As it could easily recognized from Fig. 3c1, the conjugation of ZnO with chitosan in chitosan–ZnO NPs causes a blue shift of the ZnO absorption maximum to a lower wavelength (310 nm compared with 356 nm). The reason for this is suggested to be due to the contact between ZnO and chitosan is lower (approximately 372 nm) compared to macrocrystalline ZnO²⁹. Similarly, Fig. 3d1 shows another blue shift produced by Chitosan–ZnO/PVP where the absorption maximum was oberved at 312 nm. These last couple of observations are in line with the previous report by Kubo theory³⁰. Kubo reported that as the nanoparticle’s diameter decreases, the absorption band will be blue-shifted. The peaks observed in the UV–Vis absorption spectra of chitosan, ZnO NPs and Chitosan–ZnO NPs are associated with specific electronic shifts of the functional groups or substances present in the samples³¹.

The band gap energy of Chitosan, ZnO NPs, chitosan–ZnO NPs, and chitosan–ZnO/PVP was calculated by plotting (hv)² versus energy (eV) as shown in Fig. 3a2,b2,c2,d2. The band gap energies of Chitosan, ZnO NPs, chitosan–ZnO NPs and chitosan–ZnO/PVP are 3.98, 3.48, 3.33 and 3.20 eV, respectively. These results confirm that the use of chitosan and PVP to encapsulate ZnO NPs significantly affected the optical properties of ZnO NPs and reduced the energy band gap to a minimum.

FTIR spectroscopy

Figure 4a–f shows the FTIR spectra of S. nigrum extract, chitosan, ZnO NPs, PVP, chitosan–ZnO and chitosan–ZnO/PVP nanocomposite in the respective order. The peaks appear at the following absorption bands 3488, 2991, 2802, 1695, 1417, and 1055 cm⁻¹ in Fig. 4a, which are due to the presence of the following functionalities, NH₂, CH₃, CH₂, C=O, C–N and C–O–C^32,33,34 respectively. Figure 4b shows that chitosan has a lot of functions in common with the plant extract with a new peak appearing at 644 cm⁻¹ specific to CH₃³⁵. Figure 4c shows the FTIR spectra of ZnO NPs, where a characteristic absorption peak appears at 514 cm⁻¹ for Zn=O. As shown in Fig. 4d–f, a new peak between 524 and 529 cm⁻¹ appears in the spectra of chitosan–ZnO and chitosan–ZnO/PVP nanocomposite when compared with the infrared spectrum of chitosan. This peak corresponds to the symmetric stretching vibration of ZnO³⁶. The characteristic peak of the NH group was shifted to a lower wavelength frequency of 3451 cm⁻¹³⁷. The presence of hydrogen bonds between ZnO and chitosan was confirmed by weakening the band at 1626 cm⁻¹. The characteristic bands in Figs. 4d,f moved to lower wavenumbers, indicating conjugation between the hydroxyl, amino and amide groups of chitosan and zinc oxide³⁸.

The FTIR spectrum of PVP (Fig. 4b) showed a band 1734 cm⁻¹, which was attributed to the C=O intermediate stretching of the pyrrolidone ring^39,40,41. The interaction between ZnO molecules and functional groups of chitosan and PVP was studied by Fourier transform infrared spectroscopy.

X-ray diffraction

Three diffraction peaks at 10.4°, 19.9°, and 29.1° are visible in the diffractogram of chitosan Fig. 5a–e. These peaks corresponded to the (020), (110), and (100) planes, respectively, of the crystal lattice. The peak at 10.4° had a lower intensity as compared to the intensity of the peak at 19.9°. This was likely due to the formation of the intramolecular hydrogen bonds during the deacetylation process, since these hydrogen bonds have an impact on the chitosan crystallinity and its structure⁴².

The crystallinity index was determined using the following formula:

Here, I_am is a measure of the strength of the amorphous diffraction peak centered at 2θ = 10.4°, while I₁₁₀ represents the maximum diffraction intensity at 2θ = 19.9°⁴³. The degree of crystallinity in chitosan was indicated by the calculated crystallinity index value of 74.5%. Regarding the XRD pattern of ZnO NPs, several diffraction peaks were observed at the following angles: 32.1°, 34.9°, 36.8°, 47.2°, 56.7°, 62.3°, 66.5°, 67.7°, 68.8°, and 72.9°. These peaks corresponded to the (100), (002), (101), (102), (110), (103), (200), (112), (201), and (004) planes, respectively⁴⁴, and indicated the presence of the well-crystallized ZnO structure of the hexagonal wurtzite crystal phase (JCPDS card no: 01-079-0205)⁴⁵.

In the XRD pattern of chitosan–ZnO NPs, the characteristic peaks of chitosan at 10.4° and 19.9°were less prominent. The other diffraction peaks observed in the chitosan–ZnO NPs pattern corresponded to the (100), (002), (101), (102), (110), (103), (200), (112), (201), and (004) planes⁴⁶. These peaks were sharper and stronger, indicating the high crystallinity of ZnO NPs. Based on the XRD analysis, it was concluded that the chitosan–ZnO complex was successfully formed at the nanoscale, as evidenced by the presence of the characteristic ZnO peaks and the change in the intensity of peaks in the difraction pattern of chitosan⁴⁷. The XRD pattern showed distinct crystalline structures of PVP, ZnO, and chitosan in the chitosan–ZnO/PVP nanocomposite. This suggests that the chitosan–ZnO composites were entirely embedded in the PVP polymer matrix. So, the fact that the intensity peaks have been diminished indicates that the Chitosan–ZnO nanocomposite has been effectively combined with the polymeric chains. The results corroborate those of the UV–Visible and FTIR analyses.

The crystallite size was calculated using Scherrer’s equation

where k = constant = 0.91, λ is X-ray wavelength = 1.5418 Å, θ = Bragg’s angle, and β = The Full Width at Half Maximum (FWHM) corresponding to the highest intensity peak⁴⁴.

The XRD diffraction data were utilized to deter-mine the crystallinity index and average crystallite size. As shown in Table 1, the crystallinity (%) values for the ZnO NPs, chitosan–ZnO and chitosan–ZnO/PVP were found to be 85.7%, 78.4% and 80.20%, respectively. The calculated crystallite size values were 15.3 nm, 17.8 nm and 30.1 nm for ZnO NPs, chitosan–ZnO and chitosan–ZnO/PVP, respectively.

SEM analysis

Chitosan, ZnO NPs, chitosan–ZnO NPs, and chitosan–ZnO/PVP were analysed using scanning electron microscopy (SEM) to look at their shape and size distribution. The results of these examinations are displayed in Fig. 6a for chitosan, Fig. 6b,c for ZnO NPs, Fig. 6d,e for chitosan–ZnO NPs and Fig. 6f for chitosan–ZnO/PVP.

It was established that chitosan derived from shrimp shells had a rough surface with many ridges. This shape was determined by various parameters, including the degree of deacetylation, the crystallinity index, the level of polymerization, and the temperature. ZnO NPs were spherical and elliptical agglomerates with few widely dispersed solitary NPs with an average diameter of 12 nm. Various parameters, such as the amount of the Zn(II) ions, the amount of chitosan, the temperature, and the pH, likely affected the resultant form of ZnO NPs. Finally, ZnO NPs coated with chitosan had variable shapes, as they were in the form of agglomerates of diverse forms, and this was possible due to the encapsulation of the ZnO NPs agglomerates with chitosan, which resulted in spherical, oval, and crystalline shapes. The encapsulation was successful upon comparing the morphology of ZnO NPs and chitosan–ZnO NPs. The average diameter of the latter NPs reached 21 nm.

According to the elemental analysis (Table 2), the atomic fractions of the elements in the different materials were as follows: In chitosan, carbon (C), oxygen (O), and nitrogen (N) were present at 44.19%, 34.38%, and 21.43%, respectively. In ZnO nanoparticles (NPs), zinc (Zn), oxygen (O), and carbon (C) were found at 73.42%, 24.76%, and 1.82%, respectively. For the chitosan–ZnO composite, the atomic fractions were 37.71% for carbon, 24.30% for oxygen, 15.16% for nitrogen, and 22.83% for zinc. Lastly, in the chitosan–ZnO/PVP composite, carbon, oxygen, nitrogen, and zinc were present at 40.57%, 26.12%, 16.78%, and 16.53%, respectively.

Thermal stability

Figure 7 depicts the thermal stability curves of chitosan, ZnO NPs, chitosan–ZnO NPs, and chitosan–ZnO/PVP. The first weight loss occurred for all materials at temperatures less than 100 °C due to the moisture loss, followed by the second weight loss between 200 and 600 °C due to the loss of retained crystalline water in the case of ZnO NPs and chitosan–ZnO NPs. The initial disintegration of chitosan caused the weight loss at a temperature greater than 200 °C is due to the detachment of functional groups from the chitosan backbone. The process of decomposition of chitosan continued up to 800 °C, when its carbonization was completed and thermal stability was achieved. The thermal stabilities of ZnO NPs and chitosan–ZnO NPs were achieved at about 450 °C and 600 °C, respectively. Significant changes in the thermal properties of chitosan, ZnO NPs, and the composite material comprising chitosan and ZnO NPs were observed under conditions of elevated temperatures. The thermal breakdown temperature of chitosan was found to be lower than that of ZnO NPs and the composite material of chitosan–ZnO NPs. This observation suggests that chitosan exhibits a higher degree of heat sensitivity. The rapid breakdown of the polysaccharide constituents inside chitosan molecules in its structural framework leads to enhanced sensitivity. Polysaccharides, which are characterized by their elongated sugar chains, have a high susceptibility to heat degradation. The decomposition of chitosan molecules at elevated temperatures is attributed to the fracture or chemical interactions between N-acetylglucosamine and glucosamine units. In identical conditions, it was shown that both ZnO NPs and the chitosan–ZnO NP composite exhibited greater thermal stability. This suggests that the addition of chitosan did not significantly alter the thermal properties of the nanoparticles. The results show how different chitosan is when it comes to how it reacts to heat and what that could mean for materials that are sensitive to heat.

It was obvious that the thermal constant of ZnO NPs produced with the aid of chitosan was quite high, which led to the enhancement of chitosan thermal stability when ZnO NPs were coated with it. Another research also observed similar results in the case of chitosan–ZnO⁴⁸.

The thermal characteristics of the nanocomposites were investigated through Thermogravimetric Analysis (TGA), and the thermo-gravimetric responses of the chitosan–ZnO/PVP nanocomposites are depicted in Fig. 7. The initial weight loss is observed within the temperature range of room temperature to 100 °C, attributed to the dehydration of water molecules present in all Chitosan and PVP-based materials⁴⁹.

The thermal behavior of the chitosan–ZnO/PVP nanocomposites revealed two distinct stages of weight loss occurring between 100–410 and 410–590 °C. These stages are associated with the carbonization of PVP and polysaccharides. Notably, the weight derivative of the Chitosan–ZnO/PVP nanocomposite indicates a 41% weight loss. This suggests a significant enhancement in the thermal stability of the chitosan–ZnO nanocomposite⁵⁰.

Photocatalytic degradation

Photocatalytic degradation of MB and TB dye

The best approach to the elimination of the synthetic dyes from wastewaters is their adsorption. For several metal ions and organic pigments, a high amino functionality content of chitosan offers an attractive adsorption characteristics. Chitosan functional groups can interact with MB and TB molecules through the covalent, electrostatic, and hydrogen bonding interactions. The greater acetylation of chitosan, its grafting (the introduction of functional groups), or crosslinking with other polymers may improve the adsorption properties toward various contaminants in wastewaters, and increase its resistance to harsh media conditions. The adsorption capacity of chitosan is large when the DD value is high, hence the DD of chitosan is important.

In the present work, the effectiveness of ZnO NPs, and chitosan–ZnO NPs, and Chitosan–ZnO/PVP in removing MB was 97.4%, 99.6%, and 100%, respectively, within 120 min as seen in Table 2 and Fig. 8. In the case of TB, the removal efficacy of this dye for ZnO NPs, chitosan–ZnO NPs, and Chitosan–ZnO/PVP was 96.8%, 99.5%, and 100%, respectively, also within 120 min as showed in Table 3.

Table 3 compares the efficiency of chitosan and ZnO NPs in removing the AZO dyes from diverse sources. When exposed to light, ZnO NPs can break down some dangerous chemical molecules⁵¹. The properties of ZnO NPs and the arrangement of the active species produced in the reaction media dictate the beneficial photocatalytic process. To examine the potential function of various active species in the photodegradation of Azo dyes, ZnO NPs were applied in radical scavenging tests. When the Azo dye is exposed to light, its breakdown is discovered to involve several substances, including isopropanol, formic acid, oxalic acid, and ascorbic acid⁵². However, in the present work, no extreme scavengers were used, just model organic pollutants, i.e., the MB and TB dyes, were utilized to assess the photocatalytic activity of ZnO NPs when exposed to UV radiation. Both the MB and TB dyes are toxic, carcinogenic, and non-biodegradable, posing serious issues for the human health and the environment⁵³. Therefore, the development of the effective and ecologically friendly procedure for their degradation and removal in wastewaters is of special importance. In particular, the photocatalytic degradation is recommended because it benefits from the complete mineralization of the pigments into direct, non-toxic species and much lower processing costs. Comparing the concentration of MB or TB dye in the aqueous solution after the photolysis test allowed researchers to determine how much adsorption had⁵⁴.

The MB and TB dyes were catalytically reduced when the UV light was used. The photocatalytic degradation efficiency (%) of both dyes was calculated using the equation⁵⁵.

where C_t is the current concentration, and C₀ is the starting concentration of MB or TB.

In the process of photocatalysis, the electrical structure of zinc oxide nanoparticles (ZnO NPs) is the most important thing. Zinc oxide (ZnO) is a semiconductor because it has two energy bands that can be easily identified: the valence band (VB) and the conduction band (CB). In the valence band (VB), electrons are usually in lower energy states and are attached to zinc (Zn) and oxygen (O) atoms. When ZnO is exposed to light, photons with energy equal to or greater than the band gap can be absorbed. This makes it easier for electrons to move from the valence band (VB) to the conduction band (CB). The process creates “conduction band electrons” and makes “valence band holes” inside the valence band at the same time. During photocatalysis, the presence of charged particles, such as CB electrons (e⁻) and VB holes (h⁺), is a key part of how reactive species (VB), h⁺, O₂, and OH are made⁴¹. On the surface of the ZnO nanoparticles, these charged particles help make a number of oxidation and reduction processes happen (Eqs. 1–8). As a result of the energy being higher than the band gap of ZnO, the CB electrons (e) and the VB holes (h+) are encouraged to be developed. The adsorbed MB and TB dyes may be directly oxidized by the photogenerated holes on the VB or it may react directly with the hydroxyl (OH). In the CB, the photoelectrons can change O₂ adsorbed on the surface of ZnO NPs into the superoxide radicals (O₂). For this reason, MB and TB dyes can be degraded by the photocatalysis with the production of both OH and O₂⁵⁶ as illustrated in (Fig. 8).

Recycling performance

A photocatalyst’s efficacy in water remediation applications is dependent on its separability and reusability⁶³.

After drying, the ZnO, chitosan–ZnO, and chitosan–ZnO/PVP photocatalysts were re-used in a second photocatalysis cycle with identical parameters to determine their recyclability. Figure 10e,j,o for MB dye and Fig. 9e,j,o for TB dye show the results for ZnO, chitosan–ZnO, and chitosan–ZnO/PVP in terms of the photocatalysts’ recyclability across 10 consecutive cycles. In terms of degrading MB and TB dyes, the results show that the generated ZnO, chitosan–ZnO, and chitosan–ZnO/PVP photocatalysts are very effective and may be reused. Still, after 10 cycles, the photocatalytic activity started to drop somewhat. Reasons for this might include catalyst depletion during centrifugation and washing, or the adsorption of intermediate species formed during photocatalysis⁶⁴.

After 10 photocatalytic cycles, the XRD results showed that the ZnO, Chitosan–ZnO, and Chitosan–ZnO/PVP photocatalyst maintained its critical XRD diffraction peaks, as shown in the figures. So, the ZnO, chitosan–ZnO, and Chitosan–ZnO/PVP photocatalysts’ diffraction peaks were unaltered by the catalytic material.

Effect of pH

One of the main factors that influence the rate of degradation of some organic compound pollutants is the pH value since it dictates the surface charge properties of the catalyst and size of aggregates it forms⁶⁵.

In this study, Figs. 9a–o and 10a–o represent a photodegradation efficiency, the reaction time, PH, cycles, DXR after 10 cycles the MB and the TB dye degradation percentage, where Figs. 9 and 10a–e ZnO NPs; Figs. 9 and 10f–j chitosan–ZnO NPs; and Figs. 9 and 10k–o chitosan–ZnO/PVP.

The effect of pH, on the degradation of MB and TB dyes was conducted with pH values of 3, 5, 7, and 9. The highest performance of the degradation of MB and TB dyes was achieved at pH 3, followed by pH 5, pH 7, and pH 9, as shown in Figs. 9c,h,m and 10c,h,m for MB and TB dyes respectively. As the pH value increases, the percentage of MB and TB degradation will be decreased. Semiconductor metal oxide usually exhibits an amphoteric behavior, which will influence the surface-charge (pzc) properties of the catalyst when the reactions occur on the surface of the semiconductor⁶⁶.

Thus, since MB and TB are negatively charged anions, they tend to accumulate on the surfaces of ZnO, chitosan–ZnO, and chitosan–ZnO/PVP catalysts when the pH of the solution is low. This is because these catalysts are naturally positively charged at pH values lower than pHpzc. This happens because the electrostatic interaction between the MB and TB dye molecules and the ZnO, chitosan–ZnO, and chitosan–ZnO/PVP catalysts increases, leading to faster dye degradation at low pH values.

The results showed that chitosan–ZnO/PVP had a greater effectiveness in removing both the MB and TB dyes from water than the effectiveness of chitosan–ZnO or ZnO NPs.

Antibacterial bioassay

Table 4 lists the results of the antibacterial action of chitosan, ZnO NPs, chitosan–ZnO NPs, and Chitosan–ZnO/PVP against the Gramme (+) bacteria Staphylococcus aureus and Staphulococcus aureus and the Gramme (−) bacteria Pseudomonas aeruginosa and Salmonella typhimurium (Fig. 11). It was established that when the chitosan concentration increased, the inhibitory zone of the compound widened. The antibacterial action of chitosan is mostly influenced by the type of bacteria, their stage of development, their concentration, their molecular weight, the pH and temperature of the solution, and their molecular weight⁶⁷. It is recognized that the negatively charged molecules on the bacterial cell membranes interact with the positively charged protonated amino groups of chitosan, killing the bacterial cells⁶⁸. Silva et al.⁶⁹ observed a complete antibacterial barrier after applying Buriti oil to chitosan films, while Devliger et al.⁷⁰ calculated the antibacterial efficacy of commercial chitosan against a number of putrefactive microorganisms with a high percentage of deacetylation (94%) and a lower wt. The researchers noted that while Gram (+) bacteria diffferently responded to chitosan, the Gram (−) bacteria were particularly susceptible to it.

ZnO NPs was found to havee a significant antibacterial effect against the Gram (+) bacteria but against the Gram (−) bacteria it was a lower efficacy (Table 4). The kind of the cell wall, the binding sites for ZnO NPs on the cell surface, and how ZnO NPs interact with the internal cell components decide about the antimicrobial effect of this nanomaterial. The Gramme (+) bacteria responded to the ZnO NPs antibacterial properties more readily than the Gramme (−) bacteria. This likely resulted from the contrast between Gramme (+) and Gramme (−) microorganisms in the outer membrane⁷¹.

Based on the given results, there is no doubt that chitosan exhibited the antibacterial properties, but it was less potent than ZnO NPs. Chitosan–ZnO NPs and chitosan–ZnO/PVP were characterised by a much higher antibacterial activity than chitosan alone, with the largest increase in the antibacterial activity against the Gram-positive or Gram-negative bacteria. The increase in the biological activity of both was likely associated with both the ZnO–chitosan-, and PVP-induced inhibition of the bacterial growth. However, compared with either chitosan alone, the presence of a small amount of ZnO with chitosan was sufficient to increase the antibacterial activity. Several literary reviews⁷² reported that ZnO NPs exhibit an antibacterial effect, but the specific mechanism is still under discussion. It was demonstrated that the release of the Zn(II) ions during the dissolution process is the primary mechanism responsible for the antibacterial activity of ZnO NPs⁷³. The microorganisms are also damaged and killed or destroyed by active free radicals that are formed on the surface of ZnO.

The Gram-positive bacterial strain Staphylococcus aureus was the most susceptible to the antibacterial action of the studied nanoparticles. Several critical factors likely affected this effect and its magnitude. The changes in the chemical composition and the structure of the cell membranes, and particularly the characteristics of the cell walls were possibly responsible for the variations in the antibacterial activity of the NPs against the two different types of bacteria studied here. The periplasmic region, which acts as a barrier between the outer and inner membranes and allows the entry of the inhibitory compounds, is easily solubilized by the thin outer membrane (Fig. 12)⁷⁴.

The present study observed that the antibacterial efficacy of chitosan and zinc oxide nanoparticles exhibited similar results to those reported in earlier research. Additionally, it was seen that the antibacterial activity of chitosan-coated zinc oxide nanoparticles surpassed the outcomes reached in prior investigations, as shown in Table 5. The aforementioned conclusions were drawn from the data that was gathered. This finding illustrates the successful integration of chitosan mechanisms. The bactericidal properties of zinc make it a potent factor in the efficacy of antibacterial activity.

Radical scavenger

Figure 13 shows the results of ZnO NPs, chitosan–ZnO, and chitosan–ZnO/PVP in inhibiting ABTS free radicals, where the maximum inhibition capacities of each were 63%, 67%, and 87%. These values reveal their ability to effectively neutralize ABTS radicals. It was observed that as concentrations increased, the absorbance values decreased and the inhibition percentage increased (Fig. 13a). This indicates a relationship that depends on concentration: the higher the concentration, the greater the inhibition rate (Fig. 13b).

Comparison of the radical scavenging ability of ZnO NPs, chitosan–ZnO, and chitosan–ZnO/PVP with ascorbic acid showed that all samples exhibited significant radical scavenging ability. This result is consistent with previous studies highlighting the scavenging potential of chitosan and zinc oxide nanoparticles⁸⁰.

The free radical and DPPH antioxidant activities of ZnO NPs, chitosan–ZnO, and chitosan–ZnO/PVP at different concentrations were also examined (Fig. 14a). The maximum inhibition percentages were 51%, 56% and 72%, respectively. These values indicate their effectiveness in neutralizing DPPH radicals, it was observed that as concentrations increased, the absorbance values decreased and the inhibition percentage increased (Fig. 14b). A concentration-dependent relationship was similarly observed for DPPH, with decreased inhibition capacities at lower concentrations.

The IC₅₀ results (Table 6) were as follows: 0.092, 0.079, 0.059, and 0.045 mg/mL for ZnO NPs, chitosan–ZnO, chitosan–ZnO/PVP, and ascorbic acid, respectively, in the ABTS test, and 0.095, 0.083, 0.061, and 0.064 mg/mL in the DPPH test, respectively. Incorporation of zinc into chitosan matrices and subsequently into PVP matrices significantly improved the free radical removal efficiency due to the synergistic effects of zinc, chitosan, and PVP.

Zinc exerts its inhibitory effect on ABTS and DPPH radicals through electron donation and redox reactions. This unique property, attributed to the high surface area and reactivity of zinc, enables zinc nanoparticles to donate electrons to ABTS and DPPH radicals, reducing them to stable, non-radical forms and preventing oxidative damage⁸¹. In the ABTS assay, reduction of ABTS radicals by zinc nanoparticles results in a decrease in color intensity, indicating ABTS inhibition. Likewise, scavenging of DPPH radicals reduces the intensity of the purple color of the DPPH solution, turning it yellow, indicating antioxidant activity⁸².

Chitosan shows its inhibitory effect on ABTS and DPPH radicals through its antioxidant properties and its ability to scavenge free radicals. As a biopolymer derived from chitin, chitosan is known for its antioxidant activity, possessing amino groups that effectively react with free radicals, including ABTS radicals. These amino groups (–NH2) act as electron donors, neutralizing radicals⁷⁹. Moreover, chitosan can undergo a hydrogen atom transfer process, which enhances its scavenging ability.

Incorporating chitosan–ZnO into the PVP matrix enhances its efficiency in neutralizing free radicals by combining the antioxidant properties of chitosan and zinc oxide with the excellent dispersing and stabilizing capabilities of PVP. This integration ensures uniform distribution of active sites, improves solubility and bioavailability, and increases electrostatic interactions and hydrogen bonding with free radicals. In addition, the photocatalytic activity of zinc oxide is stabilized within the matrix, resulting in more effective and sustained free radical scavenging.

Predictions of organ-specific toxicity using ProTox II

The toxicity of the screened products was examined at three distinct levels. Some toxicity endpoints include hepatotoxicity, carcinogenicity, immunotoxicity (inhibition of the B cell development), mutagenicity, and cytotoxicity. Different toxicological pathways (Aryl hydrocarbon Receptor (AhR), Androgen Receptor (AR), Aromatase, and Estrogen Receptor Alpha (ER)). Moreover, Additionally, ATPase family AAA domain-containing protein 5 (ATAD5) and mitochondrial membrane potential (MMP)) were used to evaluate the stress response pathways. Table 7 and Fig. 15 display the forecasts’ outcomes. Based on comparison of data with relevant toxicity indicator databases, products are coded as “inactive” and “active” for “non-toxic” and “toxic” respectively. According to the toxicity evaluation, the three samples examined were found to be safe and did not have any organ-specific toxicity. According to the in silico toxicity assessment, all the examined samples are predicted to be safe and to have no organ-specific toxicity. The in silico toxicity prediction with ProTox II suffers a number of limitations. These include the uncertainity and the limited reliability of these results. Accordingly, additional in vitro and/or in vivo studies are recommended to know the real toxicity upon adminstration of the studied nanoparticles.

Based on the comparison between in vivo toxicity studies and toxicity profiles generated using the ProTox-II method (Table 8), several insights can be gleaned regarding the ability of ProTox-II to predict toxicity across different endpoints.

Based on the comparison between previous studies (Table 9), it can be concluded that toxicity prediction using ProTox-II shows results consistent with experimental results in toxicity assessment.

In evaluating carcinogenicity, immunotoxicity, mutagenicity, and cytotoxicity, ProTox-II demonstrates promising predictive capabilities, aligning closely with experimental findings in many cases. For instance, compounds like isoeugenol and formaldehyde exhibit strong toxicity in both in vivo studies and ProTox-II predictions, while weaker compounds such as geraniol and cinnamic alcohol show negligible toxicity in both assessments. However, there are instances where discrepancies arise, particularly with compounds like para amino benzoic acid (PABA), where ProTox-II predicts activity in immunotoxicity despite in vivo studies suggesting otherwise. Overall, while ProTox-II shows promise in predicting toxicity across various endpoints, further refinement and validation are necessary to enhance its accuracy and reliability, ultimately aiding in the early identification of potentially hazardous chemicals and guiding safer product development processes. The use of diverse molecular descriptors and machine learning algorithms, though resulting in some variability, reflects a rich and adaptable platform capable of evolving with scientific advancements. The current models’ performance metrics, such as accuracy, highlight the platform’s robust foundation, upon which further refinements can be built. Addressing species-specific and inter-individual genetic differences represents an exciting frontier for future updates, promising even more personalized and precise toxicity predictions. Additionally, the ongoing commitment to updates ensures that ProTox-II will continue to integrate the latest data and scientific insights, expanding its predictive capabilities to include new endpoints like genotoxicity, nephrotoxicity, neurotoxicity, and cardiotoxicity. Notably, several studies have successfully utilized ProTox-II, demonstrating its practical accuracy and reliability in real-world applications. This continuous evolution positions ProTox-II as a dynamic and forward-looking tool, ever-improving to meet the complex demands of drug discovery and risk assessment^83,84.

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• Source: https://www.nature.com/articles/s41598-024-65579-z