Bioactive potential of marine Aspergillus niger AMG31: Metabolite profiling and green synthesis of copper/zinc oxide nanocomposites – An insight into biomedical applications

2.1 Isolation and characterization of marine fungi from the Red Sea

Seawater samples were collected from the Red Sea and diluted with 0.9% saline solution as a preparatory step. The diluted samples were then plated on potato dextrose agar and prepared with seawater using the spread plate technique to disperse the sample evenly. Following inoculation, the plates were incubated and periodically checked for the growth of fungal colonies starting at 14 days post-plating. Additionally, observation of morphological characteristics and microscopy facilitated identifying and enumerating the isolated marine fungal groups.

The 18S rRNA sequencing was performed to determine the genetic classification of the fungal isolates. BLAST analysis compared these sequences with existing entries in the GenBank database to identify matching patterns. Further examination of the aligned sequences between the isolates and database entries revealed the underlying phylogenetic associations [26].

To investigate bioactive compounds produced by the isolated fungi, liquid cultures were initiated by inoculating yeast extract broth and incubating for 14 days at 25°C. After this growth period, filtration through the Whatman filter paper separated the fungal biomass from the surrounding culture supernatant. The recovered culture supernatant was subjected to liquid-liquid extraction with ethyl acetate to partition secreted fungal molecules into the organic phase. Finally, the ethyl acetate fraction was concentrated via rotary evaporation to obtain a concentrated extract enriched in extracellular compounds synthesized by the marine fungi during cultivation [27].

2.2 Chemical characterization of the fungal extract

2.3 Chemical characterization of the mycosynthesized CZ nanocomposite

Chemical bonding and functional group analysis of the biosynthesized CZ nanocomposite were performed using a Cary-660 FT-IR spectrophotometer. The analysis involved preparing KBr pellets by compressing a mixture of 10 mg CZ nanocomposites with potassium bromide powder. The resulting pellets were scanned across a 400–4,000 cm⁻¹ wavenumber range to obtain their vibrational spectra [37]. X-ray diffraction (XRD) patterns of fungal-derived CZ nanocomposites were recorded on a PANalytical-X’Pert-Pro-MRD diffractometer using CuKα radiation (λ = 1.54 Å). Diffraction data collected between 10° and 80° (2θ) at 40 kV and 30 mA revealed the crystalline nature of the synthesized particles [38].

Transmission electron microscopy (TEM) analysis (JEOL Ltd-1010, Tokyo, Japan) determined the size distribution and morphology of the CZ nanocomposite. The powder underwent ultrasonication in water, followed by drop-casting onto carbon-coated TEM grids and air-drying before microscopic examination [39]. Size distribution measurements of CZ nanocomposite in colloidal suspension were obtained through dynamic light scattering analysis using Malvern Nano-ZS (Malvern Ltd., UK) [40]. The nanoparticles dispersed in MilliQ water eliminated signal interference during measurements. Surface charge characteristics were determined using Malvern Nano-ZS Zeta-sizer under identical conditions [41].

2.4 Hemocompatibility evaluation of CZ nanocomposite

Red blood cell (RBC) suspension was prepared and washed with 150 mM NaCl. The samples were centrifuged at 2,500 rpm for 10 min and resuspended in phosphate-buffered saline (pH 7.4). CZ nanocomposite concentrations between 50 and 1,000 μg·mL⁻¹ underwent testing. The RBC solution was mixed with CZ nanocomposite serial dilutions to reach a 1 mL final volume. After 60 min of incubation at 37°C and centrifugation at 2,500 rpm for 15 min, the supernatant was collected for absorbance measurements at 546 nm [42]. Isotonic solutions and full hemolysis served as negative and positive controls.

2.5 Antioxidant assessment of the fungal extract and CZ nanocomposite

2.6 Anti-inflammatory evaluation of the fungal crude extract and CZ nanocomposite

The in vitro ability of the fungal extract and the CZ nanocomposite to inhibit COX-1 and COX-2 isoenzymes was determined using COX-1 (catalogue number k548) and COX-2 (catalogue number k547) inhibitor screening assay kits (Biovision, USA), respectively, following the manufacturer’s instructions [46]. The extract was dissolved in DMSO and tested at concentrations ranging from 1,000 to 0.5 μg·mL⁻¹ in a final volume of 1 mL [47]. Celecoxib was used as a positive control for both COX-1 and COX-2 inhibition assays. The samples were tested in triplicate at 12 different concentrations.

2.7 Antibacterial activity of the fungal extract and CZ nanocomposite

The antibacterial potential of the fungal extract was investigated against various bacterial strains using the agar well diffusion method on Mueller–Hinton agar. The tested bacteria included Gram-positive Bacillus subtilis (ATCC 6633), S. aureus (ATCC 6538), Enterococcus faecalis (ATCC 29212), and Gram-negative Escherichia coli (ATCC 8739), Klebsiella pneumoniae (ATCC 13883), and Salmonella typhi (ATCC 6539). The Candida species tested were Candida albicans (ATCC 10221) and Candida tropicalis (ATCC 66029), and their results were compared to those of fluconazole as the antifungal control drug. The fungal extract and gentamicin (control) were dissolved in DMSO at a 10 mg·mL⁻¹ concentration. Wells were loaded with 10 mg·mL⁻¹ units of fungal extract [44]. After incubation, the diameters of inhibition zones around the wells were measured. Minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) were further evaluated following Clinical and Laboratory Standards Institute (CLSI) guidelines [48].

3.1 Isolation, screening, and molecular classification of Red Sea fungal strains

Fungal sampling from Red Sea sediments yielded eight isolates through dilution methods. Cross-streak tests identified AMG31 as the strain exhibiting superior characteristics to other recovered isolates. Isolate AMG31’s microscopy revealed the hallmark features of A. niger, with a prominent dark brown to black spherical vesicle. The vesicle exhibits dense radiating chains of conidia around its surface, forming a distinctive globose head. The conidiophore appears smooth-walled and unbranched, extending from the base of the vesicle. The conidia appear as small, spherical structures, displaying the characteristic dark pigmentation typical of A. niger (Figure 1a).

Figure 1

(a) Microscopic examination of A. niger AMG31. (b) Phylogenetic association tree of AMG31 with closely related Aspergillus species.

Molecular identification through sequence analysis (accession PP493930) confirmed the morphological identification, positioning isolate AMG31 within a distinct clade of the A. niger complex in the phylogenetic tree. The phylogenetic analysis reveals close evolutionary relationships with A. niger isolate Gharib 11 (PP359563) and isolate NAS-A102 (PQ606657), as indicated by their phylogenetic clustering and bootstrap support values (Figure 1b).

3.2 Chemical characterization of the fungal crude extract

3.2.1 FT-IR

The FT-IR analysis of the fungal extract revealed the presence of several key functional groups, as indicated by firm peaks in the spectrum. A strong, broad peak at 3,300 cm⁻¹ suggests the existence of hydroxyl groups from alcohols or carboxylic acids, while a strong peak at 1,709 cm⁻¹ is attributed to carbonyl groups from carboxylic acids, esters, or ketones. A strong peak at 1,033 cm⁻¹ also supports C–O stretching from alcohols or esters (Figure 2, Table 1). These findings indicate that the fungal extract contains compounds with various functional groups, including alcohols, carboxylic acids, esters, and ketones.

Figure 2

FT-IR spectrum of A. niger AMG31 crude extract.

Table 1

FT-IR analysis of the fungal extract: Observed peak wavelengths, functional groups, and bond types

Wavelength (cm−1)	Functional group	Bond type	Relative intensity
3,300	O–H (alcohol, carboxylic acid)	Stretching	Strong, broad
2,918	C–H (alkyl groups)	Stretching (CH2)	Medium
2,850	C–H (alkyl groups)	Stretching (CH2)	Medium
1,709	C═O (carboxylic acid, ester, ketone)	Stretching	Strong
1,450–1,200	C–H (alkyl groups), C–O (alcohol, ester)	Bending (C–H), Stretching (C–O)	Medium
1,033	C–O (alcohol, ester)	Stretching	Strong
719	−(CH2)n-(long alkyl chain	In-plane bending	Weak

3.2.2 GC–MS

Looking at the GC–MS analysis results of the fungus’s secondary metabolites, several significant compounds stand out. The most abundant compound is p-cresol, 2,2′-methylenebis[6-tert-butyl], comprising 24.17% of the total area. Following this, (Z)-13-docosenamide represents another major component at 16.18%. The analysis reveals a substantial presence of phenolic compounds, with phenol, 2,4-bis(1,1-dimethylethyl)-,1,1′,1″-phosphate accounting for 10.75%. The fungus also produces notable amounts of bioactive compounds like pyrrolo[1,2-a]pyrazine-1,4-dione derivatives (5.00%) and γ-sitosterol (4.26%). The presence of various long-chain hydrocarbons, fatty acid derivatives, and steroid-like structures suggests that A. niger AMG31 has diverse biosynthetic capabilities. The detection of glucosylated compounds like 6,8-DI-C-α-glucosylluteolin (3.05%) and various amines, including dimethyl myristamine (3.12%), indicates complex secondary metabolism pathways (Table 2 and Figure 3).

Table 2

Chemical compounds identified by GC–MS analysis

RT	Compound name	Chemical formula	MW	Area (%)
31.99	2-Allyl-5-t-butylhydroquinone	C13H18O2	206	0.25
32.2	1-Dodecanamine,N,N-dimethyl	C14H31N	213	1.73
36.05	Hexadecane	C16H34	226	0.42
37.96	Docosane	C22H46	310	0.22
38.83	n-Lauryl acrylate	C15H28O2	240	0.58
39.69	Dimethyl myristamine	C16H35N	241	3.12
40.02	2,6,10-Trimethyltetradecane	C17H36	240	0.43
40.65	Cyclo(l-prolyl-l-valine)	C10H16N2O2	196	0.65
43.15	Octadecane	C18H38	254	0.46
43.58	Phytan	C20H42	282	1.01
44.67	Pyrrolo[1,2-a]pyrazine-1,4-dione, hexahydro-3-(2-methylpropyl)-	C11H18N2O2	210	5.00
45.39	7,9-Di-tert-butyl-1-oxaspiro(4,5)deca-6,9-diene-2,8-dione	C17H24O3	276	1.33
46.45	2,2,3,3,4,4 Hexadeutero octadecanal	C18H30D6O	274	0.32
46.77	Methyl 14-methylpentadecanoate	C17H34O2	270	0.42
48.08	Hexadecanoate	C16H32O2	256	1.03
40.31	1-Eicosanol	C20H42O	298	1.17
49.6	14-á-H-Pregna	C21H36	288	0.48
50.27	2-Acetyl-3-(2-cinnamido)ethyl-7-methoxyindole	C22H22N2O3	362	0.19
51.7	E,E,Z-1,3,12-Nonadecatriene-5,14-diol	C19H34O2	294	0.62
52.01	Octadecanal, 2-bromo	C18H35BrO	346	0.26
52.33	N-Benzyl-N-methyl-1-tetradecanamine	C22H39N	317	0.90
52.94	MethyL-9,9,10,10-D4-octadecanoate	C19H34D4O2	302	0.23
55.61	Docosan-1-OL	C22H46O	326	1.73
55.73	Eicosyl acetate	C22H44O2	340	1.08
56.24	17-Pentatriacontene	C35H70	490	0.33
56.65	N,N-Dimethylpalmitamide	C18H37NO	283	1.00
58.11	17-Methoxyandrost-4-en-3-one o-methyloxime	C21H33NO2	331	0.61
58.25	Glycidol oleate	C21H38O3	338	0.85
59.34	Ethyl 2-[(4-methylphenyl)amino]propanoate	C12H17NO2	207	1.30
61.33	p-Cresol, 2,2′-methylenebis(6-tert-butyl)	C23H32O2	340	24.17
62.37	1-Hexacosene	C26H52	364	1.92
64.28	1,25-Dihydroxyvitamin D3, TMS derivative	C30H52O3Si	488	1.31
64.68	2,3-Dihydroxypropyl palmitate	C32H64O3	330	1.31
64.78	6,8-DI-c-á-glucosylluteolin	C27H30O16	610	3.05
65.59	9-(2′,2′-Dimethylpropanoilhydrazono)-3,6-dichloro-2,7-bis-[2-(diethylamino)-ethoxy]fluorene	C30H42Cl2N4O3	576	1.82
66.51	E-10,13,13-Trimethyl-11-tetradecen-1-ol acetate	C19H36O2	296	1.18
69.78	7,8-Epoxylanostan-11-ol, 3-acetoxy	C32H54O4	502	0.92
71.58	13-Docosenamide, (Z)-	C22H43NO	337	16.18
80.13	(R)-2,7,8-Trimethyl-2-((3E,7E)-4,8,12-trimethyltrideca-3,7,11-trien-1-yl)chroman-6-ol	C28H42O2	410	1.89
80.39	Arabinitol, pentaacetate	C15H22O10	362	0.46
83.08	ç-Sitosterol	C29H50O	414	4.26
83.38	(E)-24-Propylidenecholesterol	C30H50O	426	2.66
83.97	3-Hydroxyspirost-8-en-11-one	C27H40O4	428	0.39
91.58	Phenol, 2,4-bis(1,1-dimethylethyl)-,1,1′,1″-phosphate	C42H63O4P	662	10.75

Figure 3

GC–MS analysis of chemical compounds of A. niger’s secondary metabolites.

3.2.3 HPLC

The HPLC analysis of the fungal crude extract revealed Hesperetin as the predominant secondary metabolite, reaching 80,471.56 μg·g⁻¹, followed by Rosmarinic acid at 8,396.08 μg·g⁻¹. Daidzein emerged as the third most concentrated compound at 3,172.03 μg·g⁻¹, while naringenin and gallic acid showed substantial levels at 2,630.41 and 2,415.22 μg·g⁻¹, respectively. Chlorogenic acid maintained a notable presence at 1,751.18 μg·g⁻¹, whereas compounds like methyl gallate and ellagic acid appeared in trace amounts at 33.25 and 77.80 μg·g⁻¹ (Figure 4 and Table 3).

Figure 4

Secondary metabolite profiling of A. niger AMG31 by HPLC.

Table 3

Quantification of bioactive metabolites in A. niger AMG31

A. niger AMG31 crude extract
	Area	Conc. (µg·mL−1)	Conc. (µg·g−1)
Gallic acid	546.10	48.30	2415.22
Chlorogenic acid	269.92	35.02	1751.18
Catechin	0.00	0.00	0.00
Methyl gallate	13.20	0.66	33.25
Coffeic acid	279.16	21.60	1080.12
Syringic acid	94.40	6.90	345.19
Pyro catechol	0.00	0.00	0.00
Rutin	59.91	8.84	441.88
Ellagic acid	15.58	1.56	77.80
Coumaric acid	85.47	3.04	152.08
Vanillin	235.66	8.76	437.90
Ferulic acid	62.60	3.64	181.82
Naringenin	575.56	52.61	2630.41
Rosmarinic acid	1566.17	167.92	8396.08
Daidzein	1131.20	63.44	3172.03
Querectin	33.08	4.47	223.27
Cinnamic acid	715.91	12.82	641.03
Kaempferol	227.42	14.35	717.28
Hesperetin	32735.20	1609.43	80471.56

3.3 Characterization of mycosynthesized CZ nanocomposite

3.3.1 FT-IR

The different groups in fungal metabolites and their role in the biofabrication of CZ nanocomposite were detected using FT-IR (Figure 5). As shown, the peaks of the nanocomposite were shifted, decreased in intensity, or increased after coating with fungal metabolites. Moreover, some peaks have disappeared. The strong peak at a wavenumber of 3,320 cm⁻¹ signifies the NH of the primary amide, which overlaps with the OH group [49]. This peak was shifted at 3,400 and 3,570 cm⁻¹ wavenumbers after CZ formation. The medium peaks at 3,055 and 3,010 cm⁻¹ indicate the stretching CH of alkene, whereas the broad peak at 2,670 cm⁻¹ refers to the stretching OH group of carboxylic acid [50]. The peak at 2,315 cm⁻¹ and shifted to 2,365 cm⁻¹ after CZ formation refers to the presence of carbon dioxide (CO₂) in the air [51]. The strong peak at 2,185 cm⁻¹ corresponds to the stretching S–C≡N of thiocyanate, whereas weak peaks at 1,800 and 1,870 signify the bending C–H of aromatic compounds [52,53]. The peak at 1,625 cm⁻¹ (shifted to 1,640 cm⁻¹ after CZ nanocomposite formation) related to the stretching C═N for amides and C═O overlapped by N–H of amines [54]. Peaks in the ranges of wavenumbers of 1,250–1,550 cm⁻¹ (five peaks) in the fungal metabolites could be related to the bending C–H of alkane or aldehyde, and stretching N–O of nitro compounds [54].

Figure 5

FT-IR analysis for fungal extract vs CZ nanocomposite showing different functional groups.

These peaks decreased to only two peaks at 1,385 and 1,485 cm⁻¹ after fabrication of the CZ nanocomposite. Moreover, the 1,010–1,210 cm⁻¹ peaks correspond to the stretching C–N of amine. After formation of CZ, the peak at 1,135 cm⁻¹ refers to the stretching vibration of O–H of H₂O molecules in the lattice of Cu–Zn–O nanocomposite [55]. The peaks in the ranges of 700–900 cm⁻¹ in fungal extract are related to the bending C═C of alkene [56]. The intensity and number of these peaks are decreased after forming the CZ nanocomposite. The peaks at 695, 620, and 425 cm⁻¹ formed after CZ fabrication are related to the successful formation of Cu–O–Zn [57].

3.3.2 XRD

The nature of the fabricated nanocomposite, whether crystalline or amorphous, was detected using XRD investigation. As shown, the presence of sharp and clearance peaks for Cu and Zn in the XRD analysis indicates the successful doping and formation of a crystallographic Cu/ZnO nanocomposite (Figure 6). Bragg’s peaks at 2 theta degree of 31.8, 34.66, 36.47, 47.71, 56.85, 62.93, 66.64, 68.2, and 69.3 corresponding to (100), (002), (101), (102), (110), (103), (200), (112), and (201) confirmed successful formation of crystallographic hexagonal wurtzite ZnO structure according to JCPDS file 05-0664 [58]. On the other hand, diffraction peaks of (110), (111), (−202), (−113), (310), and (022) at 2 theta degree of 32.9°, 36.47°, 49.2°, 61.22°, 66.64°, and 68.2°, respectively, confirmed formation of crystallographic CuO based on JCPDS card 80-1916 [41]. The results are compatible with different published investigations for green synthesis of crystalline structure of Cu/ZnO nanocomposite [57,59]. The presence of other peaks in XRD analysis could be related to the fungal capping agent, which coated the surface of the nanocomposite and increased its stability.

Figure 6

XRD analysis shows Bragg’s diffraction peaks for CuO and ZnO and confirms the crystalline structure of the nanocomposite.

The average crystallite size for Cu/ZnO nanocomposite was calculated using the Debye–Scherrer equation as follows:

Average crystallite size = 1.54 × 0.9 β cos θ ,

where 1.54 and 0.9 are the wavelength of X-ray and Debye–Scherrer constant, respectively, whereas β and θ are the half maximum intensity and peak Bragg’s angle, respectively.

The overall average CZ crystallite size was calculated to be 23 nm due to the overlapping of the dominant XRD peaks for CuO and ZnO. Similarly, the overall crystallite size of Zn/CuO nanoparticles calculated by the equation of Debye–Scherrer was 26.5 nm [55]. The authors calculated the overall crystallite size due to the overlapping peaks of Cu and Zn in XRD analysis. Also, the average crystallite size of Cu doped with ZnO was in the range of 20.8–28.3 nm as calculated using XRD analysis [57].

3.3.3 Morphological and compositional structure

The shape and composition of the as-formed nanocomposite were investigated using TEM and energy-dispersive X-ray spectroscopy (EDX) analysis. These parameters are vital for analyzing nanocomposites’ biological and environmental applications [60]. TEM images exhibit successful fabrication of spherical shape, well-arranged without agglomeration, and have average sizes of 12–45 nm (Figure 7a and b). Sonkar et al. reported successful formation of Cu doped with varied concentrations of ZnO in the sizes of 19–30 nm and showed that the sizes increased by increasing the amount of ZnO [57]. On the other hand, Cao and coauthors successfully formed the spherical shape of CuO doped with ZnO with sizes in the range of 20–130 nm [21]. The authors returned this wide range of sizes due to the varied ZnO shapes after doping with CuO. Penicillium chrysogenum and Commelina benghalensis facilitated Cu/ZnO nanoparticle formation. The resulting particles mainly assume hexagonal and spherical morphologies, while researchers have also observed rod-like and coral reef-type structures. Nanoparticle dimensions span 9.0–59.7 nm, with ZnO/CuO composite materials measuring 35–50 nm in particular investigations [61]. Another study reported that the sizes of monometallic Cu and Zn are similar and small, whereas they increased with agglomeration percentages after forming bimetallic Cu/Zn nanoparticles due to the interactions of van der Waals [62]. Also, the authors reported that the wide applications of mono and bimetallic NPs are affected by surface characteristics of NPs, such as pore diameter, surface area, and charges, rather than the similarity in morphological properties [62].

Figure 7

Parts (a) and (b) are the TEM analysis of CZ nanocomposites, part (c) is the EDX analysis showing the metal composition of the synthesized sample, and part (d) is the DLS and zeta potential analysis.

The current one-pot synthesis design streamlines the process while maintaining reproducibility, addressing efficiency and cost considerations crucial for potential scale-up applications. While plant extract methods typically require 6–24 h for synthesis completion, bacterial approaches involve lengthy cultivation steps and sterile conditions. Our fungal route completed nanocomposite formation in 2 h at 85°C using a simple sequential addition of metal salts. The fungal extract acted as both a reducing agent and a stabilizer, removing the need for additional chemicals. This approach generated 12–45 nm NPs at lower costs and with fewer processing steps than plant extraction or bacterial fermentation methods.

The green synthesis protocol utilizing 7 g A. niger AMG31 biomass in 20 mL fungal extract combined with 200 mL ZnSO₄·7 H₂O (2 g) and 100 mL CuSO₄·5 H₂O (2 g) solutions demonstrated reproducible nanocomposite formation through consistent visual transitions (white to dark green) occurring within 30 min and 2 h timeframes at standardized conditions (pH 10, 85°C, atmospheric pressure, 150 rpm agitation). Reproducibility of nanocomposite characteristics is confirmed by uniform particle size distribution (12–45 nm), stable crystalline structure through XRD analysis, zeta potential measurements via DLS with minimal standard deviations across hemocompatibility testing (1.7% hemolysis at 1,000 μg·mL⁻¹). The thermal treatment (50°C, 48 h) produced dry, crystalline nanocomposites maintaining structural integrity throughout experimental procedures, while comprehensive characterization using six analytical techniques provides quality checkpoints supporting batch consistency. The combination of readily available fungal biomass, common laboratory chemicals, mild processing conditions, and short reaction durations indicated favorable scalability parameters.

The chemical composition of the synthesized nanocomposite was detected using EDX analysis. As shown, the main components of the sample were Cu, Zn, and O ions, which localized at bending energies of (0.9, 8.1, and 8.9 keV), (1.08 and 8.6 keV), and (0.5 keV), respectively (Figure 7c). The data obtained are matched with the EDX of synthesized Cu/ZnO bimetallic nanoparticles. The weight and atomic percentages of the ions within the synthesized sample were presented as 61, 17, and 22% and 59.5, 15.5, and 25% for Cu, Zn, and O ions, respectively (Figure 7c). Similarly, the weight percentages of Cu and Zn in the synthesized nanocomposite were represented by 34.7 and 36.5%, compared to the weight percentages of monometallic NPs (Cu = 74.42% and Zn = 77.42%). Also, the weight percentages of O, Cu, and Zn in green synthesized bimetallic CuO–ZnO nanostructure using the extract of Annona muricata were represented as 53.8, 14.3, and 31.9%, respectively [63]. The high Cu weight and atomic percentages compared to Zn could be related to the high reduction efficacy of Cu (0.34) compared to Zn reduction potential (−0.76) as reported previously [64]. This finding could be because the synthesis of nanocomposites was achieved through mixing two salts simultaneously in the fungal extract under stirring conditions, leading to the reduction of Cu first by the fungal metabolites, followed by Zn.

3.4 Hemocompatibility evaluation of CZ nanocomposite

The hemocompatibility data show the CZ nanocomposite tested at 25–1,000 μg·mL⁻¹ concentration. At 1,000 μg·mL⁻¹, hemolysis reached 1.7%, while 800 μg·mL⁻¹ showed 0.6% hemolysis. The mid-range concentrations of 600, 400, and 200 μg·mL⁻¹ exhibited minimal hemolysis of 0.3, 0.2, and 0.2% respectively. Lower concentrations of 100 and 50 μg·mL⁻¹ displayed 0.2 and 0.1% hemolysis (Figure S1). The lowest tested concentration of 25 μg·mL⁻¹ resulted in no measurable hemolysis (0%), matching the negative control (Table 5). Compared to the positive control’s 100%, the consistently low hemolysis values indicate the nanoparticles’ exceptional compatibility with red blood cells.

3.5 Antioxidant activity of the fungal extract and CZ nanocomposite

Antioxidant activity evaluation using a DPPH scavenging assay revealed distinct inhibition patterns between the fungal extract from A. niger and the CZ nanocomposite across multiple concentrations. At the lowest tested concentration of 1.9 μg·mL⁻¹, the fungal extract demonstrated 27% inhibition, while the CZ composite showed 12.6% inhibition. As the concentration increased to 3.9 μg·mL⁻¹, the fungal extract showed 31% scavenging activity, while at 7.8 μg·mL⁻¹, it reached 38.4% inhibition. At 15.6 μg·mL⁻¹, the fungal extract’s inhibition increased to 44.4%, whereas the CZ nanocomposite had 37.5% inhibition. This comparison revealed that while the CZ nanocomposite needed a higher concentration, it achieved a greater inhibition percentage at this point. The fungal extract crossed the 50% inhibition threshold at 31.2 μg·mL⁻¹ with 51.6% scavenging activity, corresponding closely with its calculated IC₅₀ of 25.66 μg·mL⁻¹. In contrast, the CZ nanocomposite reached 54.4% inhibition at 62.5 μg·mL⁻¹ and 63.1% at 125 μg·mL⁻¹, placing its IC₅₀ at 42.71 μg·mL⁻¹. During this concentration range, ascorbic acid had already achieved 74% inhibition at 15.6 μg·mL⁻¹.

At medium concentrations, the fungal extract showed 58.6% inhibition at 62.5 μg·mL⁻¹ and 66.5% at 125 μg·mL⁻¹. The CZ nanocomposite performed slightly better, with 63.1% inhibition at 125 μg·mL⁻¹ and 71.5% at 250 μg·mL⁻¹. By this point, ascorbic acid had reached a significantly higher inhibition of 89.1% at 125 μg·mL⁻¹, with IC₅₀ recorded as 3.26 μg·mL⁻¹. At higher concentrations, the fungal extract achieved 72.2% inhibition at 250 μg·mL⁻¹ and 78.2% at 500 μg·mL⁻¹. The CZ nanocomposite showed comparable results with 71.5 and 79.7% inhibition at 250 and 500 μg·mL⁻¹. At their highest tested concentration of 1,000 μg·mL⁻¹, the fungal extract reached 83.2% inhibition, while the CZ nanocomposite achieved 88.6% (Figure 8).

Figure 8

Dose–response DPPH radical scavenging activity by the fungal extract and CZ nanocomposite compared to ascorbic acid as a reference standard. Results expressed as mean ± SD from triplicate experiments with statistical significance determined by one-way analysis of variance (*p < 0.05, **p < 0.01, ***p < 0.001).

Moving to the ABTS˙ ⁺ scavenging assay, at lower concentrations, the fungal extract showed a gradual increase in scavenging activity, starting with 12.1% inhibition at 1.9 μg·mL⁻¹, progressing to 21.6% at 3.9 μg·mL⁻¹, and reaching 29.6% at 7.8 μg·mL⁻¹. The CZ nanocomposite displayed more robust initial activity with 39.9% inhibition at its lowest tested concentration, which progressed to 51.3 and 62.5% at 3.9 μg·mL⁻¹ and 7.8 μg·mL⁻¹. At 15.6 μg·mL⁻¹, the fungal extract demonstrated 40.7% inhibition, while the CZ nanocomposite reached 73.9% inhibition. By 31.2 μg·mL⁻¹, the fungal extract crossed the halfway mark with 51.2% inhibition, corresponding closely with its calculated IC₅₀ of 33.36 μg·mL⁻¹. while CZ nanocomposite IC₅₀ was 47.34 μg·mL⁻¹. Comparatively, gallic acid showed superior activity with 41.8% inhibition at its lowest tested concentration and an IC₅₀ value of merely 2.54 μg·mL⁻¹. The fungal extract continued its progression with 58.4% inhibition at 62.5 μg·mL⁻¹ and 69.9% at 125 μg·mL⁻¹. During this range, the CZ nanocomposite demonstrated 83.2% inhibition and 88.8% at the same tested concentrations. At a maximum concentration tested of 1,000 μg·mL⁻¹, the fungal extract reached 89% inhibition, while the CZ nanocomposite achieved near-complete scavenging with 97.6% inhibition at its highest concentration point, matching exactly the maximum inhibition displayed by gallic acid at 97.6% (Figure 9).

Figure 9

Dose-dependent fungal extract and CZ nanocomposite ABTS˙+ scavenging activity (%) compared to gallic acid standard across the concentration range (1.9–1,000 µg·mL⁻¹). Values shown as mean ± SD (n = 3) with statistical analysis by one-way analysis of variance (*p < 0.05, **p < 0.01, ***p < 0.001).

The TAC assay revealed that the fungal extract possesses a substantial total antioxidant capacity, with a mean value of 334.1 ± 4.3 AAE equivalent µg·mg⁻¹. At the same time, the CZ nanocomposite exhibited significantly higher antioxidant activity at 394.08 ± 4.491 AAE equivalent µg·mg⁻¹, both expressed as AAE. Similarly, the FRAP assay demonstrated the extract’s ability to reduce ferric ions (Fe³⁺) to ferrous ions (Fe²⁺), with a mean value of 252.45 ± 2.57 AAE µg·mg⁻¹. In contrast, the CZ nanocomposite showed markedly superior reducing power at 362.57 ± 4.179 AAE µg·mg⁻¹ (Table 6). These comparative results highlight the enhanced electron-donating capacity of CZ nanocomposite, reinforcing its potential as a more potent antioxidant biomaterial than the fungal extract.

The predominant compound, hesperetin at 80471.56 μg·g⁻¹ concentration, accounts for the robust DPPH (IC₅₀: 25.66 μg·mL⁻¹) and ABTS (IC₅₀: 33.36 μg·mL⁻¹) radical scavenging through electron donation from its hydroxyl groups [70]. While rosmarinic acid (8396.08 μg·g⁻¹) enhances antioxidant capacity by metal chelation and lipid peroxidation prevention through its catechol structure [71]. γ-Sitosterol (4.26% in GC–MS) disrupts bacterial cell walls by altering membrane stability, resulting in antimicrobial effects [72]. The combined phenolic compounds (55.517 mg·g⁻¹) and flavonoids (28.757 mg·g⁻¹) generate the measured TAC (334.1 AAE μg·mg⁻¹) and FRAP (252.45 AAE μg·mg⁻¹) through hydrogen donation and ferric ion reduction mechanisms [73].

Following our data, fungal-derived CuO–ZnO nanocomposites from A. niger exhibit remarkable free radical neutralization capabilities. The biogenic nanocomposite structures efficiently counteract unstable molecules that trigger oxidative stress [74]. A similar composite, but of bacterial origin from Bacillus velezensis spores, enabling rapid oxidation of ABTS into stable ABTS⁺ radicals without requiring H₂O₂ [75]. Similarly, a phyto-derived nanocomposite from Allium sativum (garlic) showed significant free radical scavenging capabilities through multiple DPPH, ABTS, and FRAP assays. The authors attributed the antioxidant activity to the heightened electron donation capacity through metal ion synergy, while their structural integrity and dimensions directly influence their radical scavenging performance. These stable, morphologically optimized particles enable sustained antioxidant defense through cooperative metal interactions and enhanced surface reactivity [76].

Comparatively, the fungal metabolite nigerloxin outperformed curcumin in metal ion reduction capacity while showing impressive radical neutralization across multiple testing platforms [77]. Another study reported that A. niger extracts showed enhanced antioxidant potency when cultivated in glucose-enriched media, with internal cellular compounds outperforming secreted metabolites in radical neutralization tests. The fungus produced specific phytochemicals, particularly flavonoids and phenols, that significantly boost its oxidative defense capabilities, underscoring how both growth conditions and extraction methods critically impact its therapeutic potential [78]. The ethyl acetate extract from A. niger showed exceptional radical-neutralizing properties through DPPH evaluation, reaching 95.12% elimination at 1 mg·mL⁻¹ concentration [79]. Another metabolite from the same fungus demonstrated powerful oxidative defense capabilities, inhibiting DPPH radicals beyond 50% while maintaining elevated thiol groups (>29 nmol·μL⁻¹) and superoxide dismutase (SOD) activity (>0.010 units·mL⁻¹) [80].

3.6 Anti-inflammatory activity

Next, the fungal extract and the biosynthesized composite were evaluated for their anti-inflammatory potential by inhibiting COX enzyme activity at various concentrations. The CZ nanocomposite demonstrated substantially superior COX-1 inhibitory activity compared to the fungal extract across all tested concentrations, and this is clearly reflected in the IC₅₀ values: CZ nanocomposite (22.72 ± 0.84 μg·mL⁻¹) showed approximately ninefold greater potency than the fungal extract (205.54 ± 5.98 μg·mL⁻¹), while celecoxib exhibited the most potent inhibitory effect with an IC₅₀ of 6.32 ± 0.74 μg·mL⁻¹ for COX-1 inhibition.

At the lowest tested concentration (0.5 μg·mL⁻¹), both showed minimal COX-1 inhibition, with the fungal extract exhibiting negligible activity (0.67%) compared to the CZ nanocomposite (6.23%). In contrast, celecoxib already demonstrated substantial inhibition (21.38%) at this concentration. When the concentration increased to 2 μg·mL⁻¹, the differences became more pronounced, with the fungal extract showing 3.08% inhibition vs 18.72% for the CZ nanocomposite, while celecoxib reached 36.87%. At 31.2 μg·mL⁻¹, the CZ nanocomposite achieved 58.46% inhibition, surpassing its IC₅₀ value (22.72 μg·mL⁻¹), whereas the fungal extract reached only 20.52% inhibition at this concentration. The fungal extract required a much higher concentration (250 μg·mL⁻¹) to approach its IC₅₀ of 205.54 ± 5.98 μg·mL⁻¹, reaching 56.73% inhibition at this concentration. Notably, at 125 μg·mL⁻¹, the CZ nanocomposite already demonstrated robust inhibition (80.75%), considerably outperforming the fungal extract (37.81%) at the same concentration.

At the highest tested concentration (1,000 μg·mL⁻¹), all three tested groups showed potent inhibitory activity: the fungal extract reached 80.38%, the CZ nanocomposite achieved 94.03%, and celecoxib exhibited the highest inhibition at 96.03%. Even at 500 μg·mL⁻¹, the CZ nanocomposite (91.27%) almost matched celecoxib’s performance (92.74%), whereas the fungal extract trailed considerably with 72.46% inhibition (Figure 10).

Figure 10

Inhibitory effects of fungal extract and CZ nanocomposite on COX-1 enzyme activity across concentration gradient (0.5–1,000 µg·mL⁻¹). Celecoxib served as the positive control standard. Results presented as mean ± SD from triplicate experiments with significance determined by one-way analysis of variance (*p < 0.05, **p < 0.01, ***p < 0.001).

On the other hand, concerning COX-2 inhibition, the enzyme inhibition percentage pattern followed a clear concentration-dependent relationship for all three groups, with substantial differences in their relative potencies. At the lowest tested concentration (0.5 μg·mL⁻¹), the fungal extract exhibited negligible COX-2 inhibition (0.21%), while the CZ nanocomposite showed minimal activity (2.91%). In contrast, celecoxib already demonstrated substantial inhibition (26.37%) at this concentration. Upon increasing to 2 μg·mL⁻¹, the differences became more pronounced, with the fungal extract showing merely 0.75% inhibition vs 13.58% for the CZ nanocomposite, whereas celecoxib reached 39.59%. By raising the concentration of both tested groups to 3.9 μg·mL⁻¹, celecoxib achieved 51.64% inhibition, surpassing its IC₅₀ value (3.73 ± 0.29 μg·mL⁻¹). At the same time, the fungal extract and CZ nanocomposite showed only 1.49 and 21.42% inhibition, respectively. The CZ nanocomposite reached the 50% inhibition threshold at approximately 31.2 μg·mL⁻¹ (49.12%), closely aligning with its IC₅₀ value of 33.03 ± 1.15 μg·mL⁻¹. Meanwhile, at this same concentration, the fungal extract demonstrated minimal inhibition (10.23%), and celecoxib maintained robust activity (72.38%). At 62.5 μg·mL⁻¹, the CZ nanocomposite exhibited considerable inhibition (64.58%), whereas the fungal extract reached only 14.78% inhibition.

At 250 μg·mL⁻¹, the fungal extract achieved just 33.84% inhibition, still considerably below its IC₅₀ value, while the CZ nanocomposite reached 79.43% and celecoxib exhibited 91.63% inhibition (Figure 11). The fungal extract finally approached its IC₅₀ value at 500 μg·mL⁻¹ with 61.29% inhibition, corresponding well with its measured IC₅₀ of 397.18 μg·mL⁻¹. While at the highest tested concentration (1,000 μg·mL⁻¹), all three substances demonstrated strong inhibitory activity: the fungal extract reached 76.82%, the CZ nanocomposite achieved 92.39%, and celecoxib exhibited the highest inhibition at 97.41%.

Figure 11

Concentration-dependent inhibitory effects of fungal extract and CZ nanocomposite on COX-2 enzyme activity across concentration gradient (0.5–1,000 µg·mL⁻¹). Celecoxib served as the positive control standard. Results presented as mean ± SD from triplicate experiments with significance determined by one-way analysis of variance (*p < 0.05, **p < 0.01, ***p < 0.001).

The extract contains rosmarinic acid at 8396.08 μg·g⁻¹, which blocks inflammatory processes by inhibiting NF-κB and directly reducing COX-2 expression [81]. HPLC analysis revealed quercetin (223.27 μg·g⁻¹) and kaempferol (717.28 μg·g⁻¹), both acting through various pathways that include selective COX-2 inhibition and decreased production of inflammatory mediators [82]. GC–MS analysis identified γ-sitosterol at 4.26%, which maintains cellular membrane integrity and regulates phospholipase A2, thereby limiting arachidonic acid release needed for prostaglandin synthesis [72], explaining our observed COX-1 inhibition (IC₅₀: 205.54 μg·mL⁻¹) and COX-2 downregulation (IC₅₀: 397.18 μg·mL⁻¹).

Following our results, Phytobiogenic CuO–ZnO nanocomposites from Dovyalis caffra leaves showed better antioxidant properties than single metal oxides alone. Their scavenging abilities suggest they may inhibit oxidative enzymes, including COX [83]. Plant extract-based copper nanoparticles from Catharanthus roseus and Abutilon indicum displayed anti-inflammatory effects that increased with concentration. This is related to COX inhibition in inflammatory pathways [84]. Also, l-ascorbic acid-synthesized nanocopper demonstrated anticancer and antimicrobial capabilities. These effects are often linked to inhibiting enzymes such as COX, which function in cancer development and inflammation [85].

Copper–zinc nanoparticles show anti-inflammatory effects against COX enzymes through several mechanisms. Beyond COX-2 inhibition, Copper–zinc nanoparticles influence inflammatory cytokines such as TNF-α, IL-1β, and IL-8 [86]. For example, boswellic acid-coated zinc nanoparticles reduce these cytokines, easing inflammation in ulcerative colitis models [86]. Also, copper ions irreversibly inhibit COX-2 activity, lowering the production of inflammatory prostaglandins like PGE2 [87]. These nanoparticles cause less gastrointestinal harm than standard NSAIDs [62,88]. They work by blocking COX-2 and decreasing reactive oxygen species and inflammatory cytokines, as shown in gouty arthritis studies [89].

Furthermore, copper complexes with zinc function like SOD, decreasing superoxide ions and their byproducts that contribute to inflammation [90]. This antioxidant function helps reduce oxidative stress, a key factor in inflammatory processes. These biogenic nanoparticles also improve beneficial immune cell infiltration, including CD8⁺ T cells, while limiting immunosuppressive cells such as myeloid-derived suppressor cells (MDSCs) [91,92]. Similarly, zinc complexes affect gut microbiota composition, which influences immune responses and decreases the expression of interferon-stimulated response genes, helping resolve inflammation [93].

3.7 Antimicrobial activity

The fungal extract of A. niger and CZ nanocomposite were further evaluated for antimicrobial activity against various G+ve and G−ve bacteria and two different Candida species using the agar well diffusion method. The results were compared to the control antibiotic gentamicin and fluconazole as an antifungal reference drug. MIC and MBC values were determined to evaluate the extract’s potency as well as the nanocomposite’s antimicrobial properties. Overall, the fungal extract exhibited broader spectrum antimicrobial activity than the CZ nanocomposite, demonstrating superior effectiveness against five tested bacterial strains. The CZ nanocomposite, meanwhile, showed its strongest activity specifically against E. faecalis, where it remarkably outperformed both the fungal extract and control gentamicin.

For G+ve bacteria, the fungal extract from A. niger demonstrated remarkable activity against S. aureus with an inhibition zone of 30 mm, surpassing both the CZ nanocomposite (25 mm) and the control gentamicin (25 mm). Similarly, against B. subtilis, the fungal extract produced a substantial inhibition zone of 29 mm, outperforming the CZ nanocomposite (21 mm) yet matching the control gentamicin (29 mm) (Figure 12). For E. faecalis, however, the fungal extract showed moderate effectiveness with an inhibition zone of 22 mm. In contrast, the CZ nanocomposite exhibited superior activity (32 mm), notably exceeding the control gentamicin (27 mm) (Figures S2 and S3).

Figure 12

Comparative antibacterial activity of three treatment agents. (a) Bar graph displaying inhibition zones (mm) (mean ± SD) by fungal extract, CZ nanocomposite, and gentamicin. (b) Representative agar plates showing inhibition zones from treatments where position A is the tested compound (either fungal extract or CZ nanocomposite), position B is blank, and position C is gentamicin.

Turning to G−ve bacteria, the fungal extract exhibited potent activity against E. coli with an inhibition zone of 30 mm, significantly exceeding both the CZ nanocomposite (26 mm) and control gentamicin (24 mm). Against K. pneumoniae, the fungal extract generated an inhibition zone of 22 mm, outperforming the CZ nanocomposite (19 mm) but matching the control gentamicin (22 mm) (Figure 12). Finally, for S. typhi, the fungal extract showed considerable efficacy with an inhibition zone of 25 mm, superior to both the CZ nanocomposite (20 mm) and control gentamicin (21 mm, Figures S2 and S3).

MICs and MBCs were then evaluated for both the fungal extract from A. niger and the CZ nanocomposite against the same bacterial spectrum. The CZ nanocomposite demonstrated superior antibacterial activity, particularly against G+ve bacteria. However, the fungal extract showed better effectiveness against E. faecalis and K. pneumoniae.

For B. subtilis, the CZ nanocomposite exhibited approximately twice the potency of the fungal extract, with an MIC of 15.62 μg·mL⁻¹ compared to the extract’s 31.25 μg·mL⁻¹. Similarly, the MBC values maintain this proportional relationship (31.25 vs 62.50 μg·mL⁻¹). Regarding S. aureus, the nanocomposite again demonstrated enhanced effectiveness with an MIC of 7.8 μg·mL⁻¹, whereas the fungal extract requires 15.62 μg·mL⁻¹. The MBC values followed the same pattern, with the nanocomposite requiring half the concentration (15.62 μg·mL⁻¹) compared to the extract (31.25 μg·mL⁻¹). Interestingly, E. faecalis presents a divergent trend. The fungal extract substantially outperforms the nanocomposite, requiring only 15.62 μg·mL⁻¹ (MIC) compared to the nanocomposite’s 62.5 μg·mL⁻¹. This pattern extends to the MBC values (31.25 vs 125 μg·mL⁻¹) (Table 7).

For the tested G−ve bacterial spectrum, the nanocomposite showed remarkable potency against E. coli with an MIC of 7.8 μg·mL⁻¹, which is notably lower than the fungal extract’s 15.62 μg·mL⁻¹. The corresponding MBC values follow suit (15.62 vs 31.25 μg·mL⁻¹). For K. pneumoniae, the fungal extract performed better, exhibiting an MIC of 31.25 μg·mL⁻¹ compared to the nanocomposite’s 62.5 μg·mL⁻¹. The MBC values reflect this advantage as well (62.50 vs 125 μg·mL⁻¹). With S. typhi, the nanocomposite showed greater effectiveness with an MIC of 15.62 μg·mL⁻¹ compared to the extract’s 31.25 μg·mL⁻¹. The MBC for the nanocomposite (31.25 μg·mL⁻¹) is significantly lower than that of the extract (125 μg·mL⁻¹) (Table 7).

Concerning the anticandidal assessment, the CZ nanocomposite demonstrated superior inhibition activity against C. albicans with a zone of inhibition measuring 35 mm, outperforming both the fungal extract (31 mm) and the standard antifungal fluconazole (30 mm). Conversely, against C. tropicalis, the fungal extract exhibited the strongest inhibitory effect with a 27 mm zone, while the composite produced a 20 mm zone, and fluconazole yielded a 21 mm zone (Figure 13).

Figure 13

Anticandidal activity comparison of three treatment agents. (a) Bar graph displaying inhibition zones (mm) (mean ± SD) by fungal extract, CZ nanocomposite, and fluconazole. (b) Representative agar plates showing inhibition zones from treatments where position A is the tested compound (either fungal extract or CZ nanocomposite), position B is blank, and position C is fluconazole.

Regarding the MIC and MFC values, the CZ nanocomposite demonstrated remarkable potency against C. albicans with an MIC of 7.8 μg·mL⁻¹ and an identical MFC, resulting in an MFC/MIC ratio of 1, whereas the fungal extract showed a higher MIC of 15.62 μg·mL⁻¹ and an MFC of 31.25 μg·mL⁻¹ with an MFC/MIC ratio of 2. Conversely, for C. tropicalis, the fungal extract exhibited greater efficacy with MIC and MFC values both at 15.62 μg·mL⁻¹ and an MFC/MIC ratio of 1, while the CZ nanocomposite required higher concentrations with an MIC of 31.25 μg·mL⁻¹ and MFC of 62.5 μg·mL⁻¹, yielding an MFC/MIC ratio of 2 (Table 8). These results suggest that while the CZ nanocomposite exhibits more vigorous fungicidal activity against C. albicans, the fungal extract proves more effective against C. tropicalis.

Consistent with our data, biogenic Cu–Zn NPs fabricated by Shewanella oneidensis, its antibacterial inhibitory action was quantified through both diffusion testing and concentration-dependent growth suppression, which was found to be effective against S. aureus and E. coli [94]. Green tea-synthesized copper oxide nanoparticles (GT-CuO NPs) demonstrated remarkable antibacterial efficacy against N. gonorrhoeae, killing 50% of bacteria at just 0.625 μg·mL⁻¹ and 100% at 3.125 μg·mL⁻¹ [95]. Similarly, biogenic Cu–Zn NPs were biologically fabricated, and their antimicrobial assessment revealed superior efficacy with lower MIC values (100 μg·mL⁻¹) and faster killing kinetics (3–3.5 h) against multidrug-resistant S. aureus and E. coli [96]. Similar NPs were synthesized using the methanolic extract of Ocimum sanctum, which demonstrated high antibacterial activity against S. aureus as determined through agar well diffusion, MIC, and MBC assays [97]. Actinomycetes fabricated another nanostructure that suppressed the bacterial growth of Bacillus cereus, showing the highest susceptibility (25.3 mm zone of inhibition) [98].

Biosynthesized Cu–Zn NPs demonstrated potent antifungal action as well against C. albicans with an MIC of 35.5 µg·mL⁻¹, inducing ROS production, while ultrastructural analysis revealed significant nanoparticle accumulation within the cytoplasm, cell wall, and extracellular environment of the fungal cells [99]. Similarly, nanoparticles biosynthesized using Salvia officinalis leaf extract showed potent anticandidal activity, particularly against C. tropicalis (MIC: 40 µg·mL⁻¹, MFC: 80 µg·mL⁻¹), while demonstrating significant synergistic effects when combined with conventional antifungals, showing the highest synergy with terbinafine against C. glabrata and fluconazole against C. albicans [100]. Spherical Cu–Zn nanocomposites were synthesized via biogenic and chemical routes with sizes ranging from 6 to 50 nm, demonstrating 100% growth inhibition against both P. aeruginosa and C. albicans, showing excellent antimicrobial efficacy against these clinically significant pathogens [101].

Biogenic Cu–Zn NPs suppress microbes’ growth by breaking cell walls and disrupting vital functions through several distinct mechanisms. These metallic structures penetrate cell walls, destroying membrane integrity and triggering cell rupture [102]. Also, the particles’ surface properties facilitate molecular alterations to bacterial membranes, compromising their structural stability [103]. Simultaneously, these nanoparticles trigger oxidative damage by producing ROS that attack cellular components such as proteins, lipids, and genetic material, creating an inhospitable environment for microbial survival [104].

Furthermore, the gradual liberation of copper and zinc ions (Cu²⁺ and Zn²⁺) from nanoparticle surfaces interrupts essential metabolic sequences within target organisms [105]. Cu–Zn particles excel at dismantling protective biofilms, removing this crucial bacterial shield and heightening susceptibility to antimicrobial treatments [106]. Not only removing bacterial biofilms, but also these nanoparticles’ interaction with microbial membranes creates abnormal permeability conditions, resulting in cytoplasmic leakage and cellular collapse [107].

Following our current findings, the organic extract from A. niger inhibited suppressed Fusarium oxysporum f. sp. lycopersici proliferation by approximately 51.5% at 80 µg·mL⁻¹ [108]. Likewise, examination of the filtrate crude extract unveiled remarkable fungicidal properties when tested against C. albicans, pointing toward its prospective role in combating fungal pathogens [109]. Additionally, researchers identified a unique antimicrobial polypeptide from A. niger that demonstrated effectiveness at minimal concentrations, requiring merely 8 µg·mL⁻¹ to constrain S. aureus growth, though necessitating a fourfold increase (32 µg·mL⁻¹) to achieve comparable results against drug-resistant MRSA strains [110].

The A. niger AMG31 extract’s notable bactericidal efficacy is rooted in its chemically diverse arsenal targeting multiple bacterial vulnerabilities simultaneously. Phenolic compounds (55.517 mg·g⁻¹), notably hesperetin (80471.56 μg·g⁻¹), disrupt bacterial membranes [111] by intercalating between phospholipids [112], while p-Cresol derivatives (24.17%) denature critical microbial proteins [113]. Meanwhile, flavonoids (28.757 mg·g⁻¹), including daidzein and naringenin, chelate essential metals and interrupt nucleic acid synthesis [114,115], whereas organic acids like rosmarinic acid (8396.08 μg·g⁻¹) acidify bacterial surfaces, compromising membrane integrity [116]. Furthermore, Pyrrolo[1,2-a]pyrazine derivatives (5%) subvert quorum-sensing pathways [117], whereas steroidal compounds such as γ-sitosterol (4.26%) disorganize membrane lipids, particularly effective against Gram-positive species [118].

This multi-pronged chemical approach incorporates amides like (Z)-13-docosenamide (16.18%) functioning as membrane-solubilizing surfactants [119], while tannins (18.650 mg·g⁻¹) irreversibly bind proline-rich proteins [120], and block enzymatic substrate access [121]. Consequently, this sophisticated chemical composition explains the extract’s broad-spectrum efficacy, evidenced by equivalent potency against both E. faecalis and E. coli (MIC 15.62 μg·mL⁻¹), often matching or surpassing gentamicin’s performance. Additionally, the consistent MBC/MIC ratios 2 confirm the extract’s bactericidal rather than bacteriostatic action.

Several future research directions emerge to advance our understanding and applications of marine fungal metabolites and their derived NPs. The initial step involves expanding the screening process to encompass diverse marine fungal strains from various Red Sea depths and locations, which could uncover novel bioactive compounds with unique therapeutic properties. This expansion should be complemented by comprehensive in vivo toxicological assessments of fungal metabolites and BCZ nanoalloy, ensuring their safety profiles for potential pharmaceutical applications. Further research should delve into the underlying mechanistic pathways of these compounds’ antimicrobial and anti-inflammatory effects through molecular docking studies and gene expression analyses, providing deeper insights into their therapeutic mechanisms.

Furthermore, developing optimized formulations for drug delivery systems and investigations into scalable nanoparticle production methods would facilitate the transition from laboratory findings to practical applications. Lastly, exploring the potential synergistic effects between fungal extracts and their synthesized NPs could uncover enhanced therapeutic properties, possibly leading to more effective treatment strategies.