Fungal xylanases‐mediated synthesis of silver nanoparticles for catalytic and biomedical applications

Abstract

Green synthesis of nanoparticles has fuelled the use of biomaterials to synthesise a variety of metallic nanoparticles. The current study investigates the use of xylanases of Aspergillus niger L3 (NEA) and Trichoderma longibrachiatum L2 (TEA) to synthesise silver nanoparticles (AgNPs). Characterisation of AgNPs was carried out using UV–Vis spectroscopy, Fourier transform infrared spectroscopy (FTIR), and transmission electron microscopy, while their effectiveness as antimicrobial, antioxidant, catalytic, anticoagulant, and thrombolytic agents were determined. The colloidal AgNPs was brownish with surface plasmon resonance at 402.5 and 410 nm for NEA‐AgNPs and TEA‐AgNPs, respectively; while FTIR indicated that protein molecules were responsible for the capping and stabilisation of the nanoparticles. The spherical nanoparticles had size of 15.21–77.49 nm. The nanoparticles significantly inhibited the growth of tested bacteria (63.20–88.10%) and fungi (82.20–86.10%), and also scavenged DPPH (37.48–79.42%) and hydrogen peroxide (20.50–96.50%). In addition, the AgNPs degraded malachite green (78.97%) and methylene blue (25.30%). Furthermore, the AgNPs displayed excellent anticoagulant and thrombolytic activities using human blood. This study has demonstrated the potential of xylanases to synthesise AgNPs which is to the best of our knowledge the first record of such. The present study underscores the relevance of xylanases in nanobiotechnology.

1 Introduction

Nanobiotechnology is an offspring of nanotechnology which is at the meeting point of nanotechnology and biology [1]. The evolution of nanobiotechnology has led to the fusion of life sciences with nanoscience, nanotechnology, and engineering. In general, nanotechnology deals with the synthesis and stabilisation of nanoparticles of various origin [2] using different types of physical and chemical methods, with the drawbacks of the use of hazardous chemicals and procedures, high consumption of energy, and being expensive. Therefore, biological method of synthesis is a substitute, which has presented promising solutions to the various problems caused by physical and chemical methods [3]. The green synthesis of nanoparticles has attracted immense attention in recent times as a result of its advantages such as cost‐effectiveness, biocompatibility, simplicity, eco‐friendliness, and extensive applications. Biomolecules of different kinds from plant and microbial origin have been effectively employed to synthesise nanoparticles. However, there is rise in recent times in the biofabrication of metallic nanoparticles using enzymes, agrowastes, and microbial and plant‐derived pigments which have been used for several biomedical applications [4].

Metallic nanoparticles, for instance, silver and gold nanoparticles, have immense applications in optics, biomedicine, and drug delivery [5]. These nanoparticles have been effectively synthesised using various agrowastes like Cocos nucifera coir and wheat bran [6, 7]. These agrowastes contain bioactive compounds such as proteins, flavonoids, and phenolics that are capable of reducing metallic ions to nanoparticles. In our laboratory, we have expanded the frontier of agrowastes in the green route approach to synthesise AgNPs using the extracts of kola seed shell [8], kola pod [9], and cocoa pod husk [10].

Several works have documented microbial and plant resources [11, 12, 13, 14, 15, 16] to synthesise AgNPs. In addition, we have recently explored cell‐free extract of Bacillus safensis LAU13 for synthesis of AgNPs [17, 18] as well as strains of Enterococcus species for synthesis of AgNPs [19]. Also, metabolites derived from arthropods such as spider cobweb [20] and paper wasp nest [21] for the synthesis of AgNPs have been documented [22].

Moreover, nanoparticles can be synthesised by both enzymatic and non‐enzymatic processes. Durán et al. [23] described the participation of thiol groups and disulphide bridge moieties of enzymes as the reaction sites of formation of nanoparticles. Additionally, it was also reported that the S‐H and S‐S moieties of denatured enzymes could convert metallic ions to nanoparticles. The use of enzymes for the synthesis of AgNPs has been reported, but there is insufficient research carried out in this area of nanobiotechnology, especially when compared with the use of agrowastes [4].

In the literature, α‐amylase was employed for synthesis of AgNPs, which were hexagonal and triangular in shape with sizes ranging from 22 to 44 nm [24]; while spherical AgNPs synthesised using laccase displayed potent antimicrobial activities [25, 26]. In addition, keratinase from a keratinolytic bacterium [27, 28] biofabricated spherical‐shaped AgNPs of 5–30 nm with potent antibacterial activities [29]. Also, nitrate reductase and cellulase have been used for synthesis of spherical‐shaped AgNPs [30, 31]. However, there is dearth of information on the involvement of xylanase to synthesise nanoparticles.

Over the years, prospects of industrial applications of xylanases in the manufacturing of pulp and paper have progressively increased, and some have been employed commercially. The most efficient usage of xylanase has been in pre‐bleaching process of kraft pulp to appreciably lessen usage of harsh chemicals in the processing of pulp [32]. Xylanases are also used as food additives in poultry feeds, enhancements of dough rising in bread making, and clarification of fruit juices [33, 34, 35]. The aim of the present study was to investigate the applications of crude fungal xylanases of Aspergillus niger L3 and Trichoderma longibrachiatum L2 previously isolated in our laboratory [35] for the synthesis of AgNPs, and examine their antimicrobial, antioxidant, catalytic, anticoagulant, and thrombolytic properties. This is in an attempt to further expand the scope of applications of xylanases to nanobiotechnology. Until now, there is no such report of the nanobiotechnological applications of xylanases.

2 Materials and methods

2.1 Microorganisms

Non‐aflatoxigenic fungal strains of A. niger L3 (NEA) and T. longibrachiatum L2 (TEA) used in the enzyme production and other fungal strains of A. niger, Aspergillus fumigatus, and Aspergillus flavus used for the antifungal studies were previously isolated from some stored food products [36]. Clinical bacterial strains that include Escherichia coli, Klebsiella granulomatis, Pseudomonas aeruginosa, and Staphylococcus aureus which were used for the antibacterial studies were originally sourced from LAUTECH Teaching Hospital, Ogbomoso, and obtained from the stock culture of Laboratory of Industrial Microbiology and Nanobiotechnology, LAUTECH, Ogbomoso. All the microbial strains were regularly subcultured as appropriate.

2.2 Enzyme

Crude xylanases were synthesised by non‐aflatoxigenic fungal strains of A. niger L3 (28.69 U/ml) and T. longibrachiatum L2 (22.13 U/ml) as previously reported by Elegbede and Lateef [35] and maintained at 4°C until use.

2.3 Biosynthesis of AgNPs

The xylanases synthesised by fungal strains of A. niger L3 and T. longibrachiatum L2 [35] were used for synthesis of AgNPs as previously described [26, 29]. About 1 ml of crude enzyme was reacted with 50 ml of 1 mM silver nitrate (AgNO₃) solution for synthesis of AgNPs at ambient temperature (30 ± 2°C). The biofabrication of AgNPs was monitored by observing the change of colour.

2.4 Characterisation of AgNPs

Characterisation of the AgNPs was carried out by measuring the absorbance spectrum using UV–Vis spectrophotometer. Moreover, Fourier transform infrared (FTIR) spectroscopy analysis was done according to Bhat et al. [1] to establish the biomolecules responsible for the synthesis, capping, and stabilisation of AgNPs. Energy diffraction X‐ray (EDX) was carried out to determine the prevalent element in the biosynthesised AgNPs. Also, transmission electron microscopy (TEM) micrographs were obtained to elucidate shapes and sizes of the biosynthesised AgNPs as previously reported [29].

2.5 Antibacterial and antifungal activities of AgNPs

The antibacterial effectiveness of NEA‐AgNPs and TEA‐AgNPs were investigated against strains of clinical bacterial isolates; E. coli, K. granulomatis, P. aeruginosa, and S. aureus, using the adapted liquid culture method of Prasannaraj and Venkatachalam [37]. The test bacterial strains were introduced into nutrient broth, and incubated at 37°C for 24 h to tune of 0.5 McFarland standard. Then, 1 ml each of gradient concentrations (10–100 µg/ml) of the AgNPs was added into test tubes containing 8 ml of peptone water inoculated with 1 ml of the test bacterial suspension and incubated at 37°C for 24 h. The control contained 1 ml of each bacterial suspension without the addition of AgNPs. The bacterial growth was monitored by measuring the optical density at 600 nm using UV–Vis spectrophotometer. The percentage inhibition of growth was calculated using the methods of Salem et al. [38] as follows:

$$\text{\%}\mathrm{Growth}\mathrm{inhibition}=\left\{\left(\mathrm{Absorbanc}{\mathrm{e}}_{\mathrm{control}}-\mathrm{Absorbanc}{\mathrm{e}}_{\mathrm{test}}\right)/\mathrm{Absorbanc}{\mathrm{e}}_{\mathrm{control}}\right\}\times 100\text{\%}$$

The antifungal activities of AgNPs were investigated against the test fungal isolates (A. niger, A. flavus, and A. fumigatus) as earlier reported [39]. Fresh fungal culture plug of 6 mm diameter were placed on potato dextrose agar medium impregnated with AgNPs (100 and 150 µg/ml), while the control lacked nanoparticles. All the plates were incubated for 48 h at 30 ± 2°C, after which the fungal radial growth was measured to determine the growth inhibition as follows:

% Growth inhibition = {(D _control − D _test)/D _control } × 100%, where D represents diameter of fungal growth.

2.6 Dye degradation activities of AgNPs

The degradation of malachite green and methylene blue using AgNPs was monitored via decolourisation, whereby 1 ml of the biosynthesised AgNPs at concentrations of 20–100 µg/ ml was mixed with 9 ml of 40 ppm of the dyes. The control experiment contained only 10 ml of the dye. All the reaction vessels were agitated on a rotary shaker at 100 rpm for 24 h at 30 ± 2°C. At specific intervals, absorbance readings of the reaction mixtures were measured using UV–Vis spectrophotometer at wavelength of 619 and 690 nm for malachite green and methylene blue, respectively. The percentage biodegradation of the dyes due to bioreduction capability of the nanoparticles in each of the reaction vessels were estimated [40] as follows:

% dye degradation = {(A _control −A _test)/A _control } × 100%; where A represents the absorbance value.

2.7 DPPH‐free radical‐scavenging activities of AgNPs

This was determined by using 2,2‐diphenyl‐1‐picrylhydrazyl (DPPH) (Sigma‐Aldrich, Germany) following the method reported by Williams et al. [41]. About 1 ml of gradient concentrations (10–100 µg/ml) of the AgNPs was added to 4.0 ml methanolic solution of 0.1 mM DPPH, agitated and then incubated in the dark for 30 min at 30 ±  2°C. The blank contained 1.0 ml of methanol mixed with 4.0 ml of 0.1 mM methanolic DPPH. The absorbance after incubation was obtained at 517 nm on a UV–Vis spectrophotometer and the inhibitory percentage of DPPH was calculated consequently [42]

$$\mathrm{Percentage}\mathrm{inhibition}=\left\{\left(A_{\mathrm{blank}}-A_{\mathrm{sample}}\right)/A_{\mathrm{sample}}\right\}\times 100\text{\%}$$

2.8 Hydrogen peroxide scavenging activities of AgNPs

The modified methods of Bhakya et al. [43] were employed, whereby 4 ml of graded concentrations (1–40 µg/ml) of the AgNPs was reacted with 0.6 ml of 40 mM H₂ O₂ prepared in phosphate buffer (pH 7.4) at 30 ± 2°C for 20 min. While H₂ O₂ solution only was used as the control, distilled water served as the blank, and the absorbance taken at 610 nm. The percentage peroxide scavenging activity was estimated thus:

peroxide scavenging activity = {(A _control −A _sample)/A _control } × 100%; where A is the absorbance.

2.9 Anticoagulant and thrombolytic activities of AgNPs

The anticoagulant assay was carried out in accordance with the established procedures [14, 44]. Blood samples of a healthy voluntary donor were collected in EDTA bottle and also in clean Eppendorf tubes mixed with xylanase, and silver nitrate solution (control). For the test experiment, 1 ml of 100 µg/ml AgNPs was interspersed separately with the fresh blood in equal proportions in Eppendorf tubes. The experiments and control were monitored visually for up to 4 h and all the Eppendorf tubes were then inverted to demonstrate anticoagulation or otherwise. Also, content of each of the tubes was smeared on a clean microscopic slide, and examined with optical microscope to visualise the red blood cells. The capabilities of the AgNPs in the management of blood clots (thrombi) were investigated in accordance with the scheme of Harish et al. [7]. The clots were formed in vitro by smearing healthy blood of a voluntary donor on a clean, grease‐free glass slide which served as the negative control, while the positive control experiments included of fresh blood combined with EDTA (anticoagulant). Other controls were set up through the addition of xylanase, and AgNO₃ solution to preformed blood clots on clean glass slides. About 0.2 ml of 170 µg/ml of AgNPs was added in a drop wise manner to the blood clots and allowed to react for 5 min for the test experiments. The thrombolytic activities of the AgNPs on the blood clots were visually inspected and also through optical microscopy.

3 Results and discussion

3.1 Biosyntheses of AgNPs

Crude xylanases produced by strains of A. niger L3 and T. longibrachiatum L2 biosynthesised AgNPs (NEA and TEA) in 5 min under ambient conditions of temperature (30 ± 2°C) with typical dark brown colour (Fig. 1) which further deepened over time and eventually stabilised within 15 min. The dark brown colour of biosynthesised AgNPs have been extensively reported [8, 9, 17, 18, 20, 44].

Fig. 1.

Course of biosynthesis of AgNPs using xylanase of A. niger L3 (NEA) and T. longibrachiatum L2 (TEA)

(a) Salt solution, (b) Crude xylanase, (c) Reaction mixture, (d) Initial colour change, (e) Stabilised nanoparticles

3.2 Characterisation of biosynthesised AgNPs

Maximum absorbance occurred at the wavelengths of 402.5 and 410 nm for NEA and TEA‐AgNPs, respectively (Fig. 2). The values obtained are within the range of 391–460 nm established for AgNPs [26, 29, 45, 46, 47, 48]. FTIR spectra for the crude xylanases synthesised by both A. niger L3 and T. longibrachiatum L2 showed strong bands at 3311, 3307, and 1635 cm⁻¹ (Figs. 3 a and b). Also, FTIR absorption spectra revealed distinct and prominent bands at 3277, 3294, and 1641 cm⁻¹ for both biosynthesised AgNPs (NEA and TEA) (Figs. 3 c and d). Other minor peaks were shown at 2387, 2304, 2187, 2156, 1514, 1367, 513, and 457 cm⁻¹ for both AgNPs. The broad peaks at 3292–3336 and 1641 cm⁻¹ relate to N‐H bond of amines, and C = C stretch of alkenes or C = O stretch of amides, respectively [49]. Also, the broadness of the band may be as a result of overlap of both O‐H and N‐H bond stretching of 1° and 2° amines. The peaks at 1367–1369, 1514, 1641, and 2154–2187 cm⁻¹ were ascribed to the 3° O‐H vibration of alcohols, aromatic nitrogen compounds, N‐H bend of 1° amine, and C≡C stretch of alkynes, respectively. It was apparent from these bands that biomolecules rich in amine (N‐H) and hydroxyl (O‐H) groups from the crude xylanases were accountable for the reduction in Ag⁺, as well as capping and stabilisation of the AgNPs [50]. The shift of the major peak from 1635 cm⁻¹ of the crude xylanase to 1641 cm⁻¹ in the biosynthesised AgNPs presupposes the chelation between Ag⁺ of the synthesised AgNPs and C = O from the peptide in the enzyme.

Fig. 2.

UV–Vis absorption spectra of the biosynthesised AgNPs

(a) NEA, (b) TEA

Fig. 3.

FTIR spectra of crude xylanase

(a) NE, (b) TE; and AgNPs, (c) NEA, (d) TEA

The TEM micrographs revealed that AgNPs synthesised have shapes ranging from spherical, cylindrical, and oval shapes having sizes of 18.11–77.49 nm for NEA‐AgNPs and 15.21–54.03 nm for TEA‐AgNPs (Fig. 4 a). The energy‐dispersive X‐ray analysis (Fig. 4 b) revealed prevalence of silver [8, 14, 17, 38], while the ring shape of selected area electron diffraction (SAED) pattern (Fig. 4 c) also revealed the face‐centred cubic crystalline nature of the biosynthesised AgNPs [29, 44, 49].

Fig. 4.

Characterisation of the nanoparticles: TEM micrographs

(a) EDX spectrum, (b) and SAED, (c) of the biosynthesised AgNPs

3.3 Antibacterial activities of AgNPs

Bacterial growth inhibition was enhanced with increase in concentrations of nanoparticles applied. Both biosynthesised AgNPs displayed outstanding antibacterial activities against the test bacterial strains; E. coli, K. granulomatis, S. aureus, and P. aeruginosa.

For NEA‐AgNPs, the activities were in the tune of 87.1, 84.2, 82.4, and 63.2% correspondingly at 100 µg/ml, while TEA‐AgNPs also displayed activities in the tune of 69.2, 88.1, 80.1, and 80.1% correspondingly at 100 µg/ml (Fig. 5 a). The least activities obtained were at 10 µg/ml for P. aeruginosa and S. aureus for NEA‐AgNPs and TEA‐AgNPs, respectively. MIC₅₀ values of 10, 20, 60, and 80 µg/ml were obtained for E. coli, K. granulomatis, S. aureus, and P. aeruginosa, respectively, for NEA‐AgNPs, while values of 20, 60, 80, and 80 µg/ml were obtained for E. coli, P. aeruginosa, S. aureus, and K. granulomatis, respectively, for TEA‐AgNPs. High growth inhibition by AgNPs in broth might be as a result of improved distribution and contact of bacterial cells with silver ions than in solid medium. Also, small sized AgNPs can easily infiltrate bacterial cell when growing in the liquid medium and this could be one of the apparent reasons why they could bring about improved antibacterial effect in liquid media [37]. Similarly, antibacterial activities of AgNPs have been connected to the interaction of AgNPs with sulphur and phosphorus containing constituents of the bacterial cell to instigate cell killing by attacking the cell division and respiratory chain [21]. It has been reported that silver nanoparticles (AgNPs) synthesised from Ficus sycomorus inhibited growth of bacteria [38]. Also, exposure of AgNPs biosynthesised from the leaf extract of Ocimum sanctum in liquid medium displayed elevated percentages of antibacterial activity [51]. Furthermore, Prasannaraj and Venkatachalam [37] reported that AgNPs biosynthesised using leaves and barks of medicinal plants displayed considerable antimicrobial activities in liquid medium when tested against P. aeruginosa, K. pneumoniae, E. coli, S. aureus, P. vulgaris, and S. epidermidis with growth inhibition percentage of 24.2–86.9%.

Fig. 5.

Antimicrobial activities of nanoparticles: Antibacterial growth inhibition

(a) and antifungal activities, (b) by NEA‐AgNPs and TEA‐AgNPs

3.4 Antifungal activities of AgNPs

Both biosynthesised AgNPs appreciably subdued the growth of A. niger, A. flavus, and A. fumigatus at tested concentrations of 100 and 150 µg/ml by 82.20–86.10% (Fig. 5 b) with MIC established at 100 µg/ml for both AgNPs against all the fungal isolates. These antifungal activities are in line with reports of previously published researches [10, 15, 21, 39, 52]. The inhibitory capabilities of nanoparticles can be correlated with the initiation of attack on the cell wall, which destroys fungal spores, thus resulting in the seepage of intracellular constituents which ultimately leads to cell death. In addition, acidification of intracellular environment resulting from the inhibition of H⁺ ‐ATPase in yeast cells have been associated with the potent antifungal activities of AuNPs [53].

3.5 Dye degradation activities of AgNPs

The biosynthesised nanoparticles degraded malachite green and methylene blue to varying degrees as shown in Figs. 6 a and b. Both biosynthesised AgNPs at different concentrations noticeably decolourised the tested dyes within the range of 64.30–78.97% for malachite green, and 14.8–25.3% for methylene blue after 24 h of reaction. It was observed that as the reaction time increased, there were reduction in the intensities in the colours of the nanoparticles‐treated dye, alluding to the potential of biosynthesised nanoparticles in biodegradation of dye as previously reported [15, 21, 54, 55, 56, 57, 58]. However, in the present study, malachite green was better degraded than methylene blue. Catalysis of dye degradation by nanoparticles is achieved through redox reaction and this is because nanoparticles can act as electron transfer mediators between the biomolecules on the surface of dye and particles. The technique of using nanoparticles in the treatment of wastewaters holds more promise for efficiency, as against conventional physical and chemical methods that basically concentrate or transfer the dyes from one phase to another.

Fig. 6.

Catalytic degradation of dyes by biosynthesised nanoparticles:

(a) malachite green, (b) methylene blue

3.6 Antioxidant activities of AgNPs

3.6.1 DPPH‐free radical scavenging activities

The biosynthesised nanoparticles displayed elevated DPPH radical scavenging potentials with activities ranging from 37.48 to 79.42% for AgNPs at 10–100 µg/ml (Fig. 7 a). The scavenging activities of the AgNPs were comparable to those previously reported [9, 10, 16, 17, 43]. The free radical scavenging potentials of nanoparticles have been credited to the functional groups of bioreductant molecules, whose ability to stick to the surface of the nanoparticles may result in amplified surface areas for activity [43].

Fig. 7.

Antioxidant activities of nanoparticles: DPPH free radical

(a) and hydrogen peroxide, (b) scavenging activities of AgNPs

3.6.2 Hydrogen peroxide scavenging activities

An impulsive clearance of the cloudy solution of H₂ O₂ that was prepared in phosphate buffer was obtained once reacted with AgNPs yielding activities of 20.50–96.50% at concentrations of 1–40 µg/ml (Fig. 7 b). Comparable results were reported by Bhakya et al. [43] where H₂ O₂ scavenging activity of 93.31% was obtained for AgNPs biosynthesised using root extract of Helicteres isora and Lateef et al. [44], where H₂ O₂ scavenging activity of 77–99.8% was obtained for AgNPs biosynthesised using extracts of cobweb, pod, seeds, and seed shell of Kola nut fruit. H₂ O₂ are produced in vivo due to the activities of some enzymes such as superoxide dismutase; and because they are incorporated into most personal care products such as bleaching agent or disinfectant in a bid to kill germs. Humans also become extremely exposed to them through their regular usage. Exposure to H₂ O₂ promotes the generation of very reactive hydroxyl radicals (•OH) in the living system [59] which thereafter leads to cellular injury and several debilitating diseases [60]. Thus, it is fundamental to limit contact of living cells to H₂ O₂, and this can be achieved by consumption of antioxidant‐rich foods [61] and treatment of environmental sources of H₂ O₂, mainly drinking and wastewater.

3.7 Anticoagulant and thrombolytic potentials of AgNPs

Formation of clots was inhibited by the addition of the biosynthesised nanoparticles to fresh blood of healthy human donor (Fig. 8 a) as observed in the positive control (EDTA), whereas coagulation was noticed in all the negative controls and this was confirmed by the microscopic examinations of blood samples. The result obtained is in conformity with anticoagulant potentials of AgNPs synthesised from diverse biomolecules as previously reported [14, 15, 18, 21, 44, 57, 62]. Although the anticoagulation characteristic is very valuable in nanomedicine to avoid coagulation of blood in patients suffering from blood coagulation disorders, it also indicates the fact that such nanoparticles can be used as drug carriers or as coating agents on medical instruments as the contact with blood shall not lead to commencement of blood coagulation [63].

Fig. 8.

Anticoagulant and thrombolytic potentials of nanoparticles: Anticoagulant

(a) and thrombolytic, (b) activities of biosynthesised AgNPs

Similarly, the nanoparticles generated clot dissolution within 5 min of reaction when reacted with preformed blood clot (Fig. 8 b). The control experiments performed with crude xylanases (NE and TE), and AgNO₃ displayed no thrombolytic activities. The optical micrographs showed the dispersal of cells due to reaction of blood clot with biosynthesised AgNPs which indicated potentials of the AgNPs for suspension of thrombus. A number of biologically synthesised nanoagents and nanoparticles have been developed to manage cardiovascular pathologies and other related health challenges [7, 64]. Though blood clotting is required to prevent bleeding, but its dissolution is equally important in preventing thrombosis and maintenance of haemostasis, of which nanoparticles play important roles in rendering efficient lysis of blood clots [65]. The use of nanoparticles in thrombolysis circumvents the problems of neutralisation by antibodies, short half‐life, and risk of excessive bleeding that are found associated with conventional and traditional anti‐thrombotic treatments such as streptokinase. The anticoagulant and thrombolytic potentials of metallic nanoparticles have been recently reported [14, 15, 18, 21, 44, 57, 58, 62]. The pioneering results of the present investigation on the use of xylanases to produce nanoparticles for potential management of blood coagulation disorder may represent a paradigm shift in the entry of enzyme nanobiotechnology in nanomedicine.

4 Conclusion

Fungal xylanases synthesised by A. niger L3 and T. longibrachiatum L2 biofabricated spherical, cylindrical, and oval‐shaped AgNPs with sizes ranging from 15.21 to 77.49 nm. Both biosynthesised AgNPs displayed noteworthy antibacterial and antifungal activities. The nanoparticles also showed prominent antioxidant potentials to scavenge DPPH and H₂ O₂. Furthermore, the particles were able to effectively degrade malachite green and methylene blue dyes. The nanomedical significance of the nanoparticles in the potential management of blood coagulation disorders and thrombotic diseases was established as both AgNPs efficiently dissolved blood clots and also functioned as excellent blood anticoagulants. Therefore, this study has comprehensively established the potential vast nanobiotechnological applications of fungal xylanases, displaying them as immeasurable utilities in both industrial and biomedical sectors.