Biogenic synthesis of zinc oxide nanoparticles using Pichia cactophila-derived β-glucan as a novel capping agent: Characterization, antioxidant, antibacterial, and cytotoxicity evaluation

Abstract

The present study reports the isolation and molecular identification of Pichia cactophila JP strain from Jogimatti Forest soil, Karnataka, India, and the first-ever use of its extracted β-glucan as a biogenic capping agent for ZnO nanoparticle synthesis. β-glucan was isolated via acid-alkali extraction (yield: 4.6% w/w dry cell weight) and employed in a co-precipitation method to synthesize YGZnNP-1. Physicochemical characterization confirmed a crystalline hexagonal wurtzite ZnO structure (JCPDS No. 36-1451), an average crystallite size of ∼28.5 nm, and a UV absorption peak at 372 nm. FTIR spectroscopy verified effective β-glucan surface functionalization. YGZnNP-1 exhibited potent dose-dependent antioxidant activity (DPPH IC₅₀: 79.73 μg/mL), significant antibacterial activity against Pseudomonas aeruginosa and Staphylococcus aureus, and favourable biocompatibility against NIH 3T3 fibroblast cells (IC₅₀: 98.20 μg/mL). These findings establish YGZnNP-1 as a promising multifunctional nanomaterial for biomedical applications.

1. Introduction

The rapid advancement of nanotechnology has transformed multiple scientific domains, with particularly profound implications for materials science, pharmaceutics, and biomedical research. Nanomaterials (1–100 nm in size) exhibit physicochemical properties that are absent in their bulk forms-notably high surface-to-volume ratio, quantum confinement effects, and programmable optical, magnetic, and electronic characteristics, rendering them highly attractive for diverse applications in biomedical and industrial sectors.1, 2 Among various classes of nanomaterials, metal oxide nanoparticles have emerged as one of the most actively investigated groups owing to their enhanced reactivity, stability, and multifunctional biological activity.1, 2 Zinc oxide nanoparticles (ZnO-NPs) have attracted extensive research attention due to their broad-spectrum biological activities, including antibacterial, antifungal, antioxidant, anticancer, anti-inflammatory, and wound healing properties, alongside their biocompatibility and environmental safety. These properties, combined with low production cost and chemical stability, position ZnO-NPs as promising candidates for pharmaceutical, biomedical, and industrial applications.

Conventional physical and chemical methods for nanoparticle synthesis, including co-precipitation, hydrothermal synthesis, sol-gel processing, and sputtering, are constrained by their dependence on hazardous reducing agents and stabilizers, high energy consumption, generation of toxic by-products, and elevated environmental risks.³ To overcome these drawbacks, biosynthesis strategies leveraging organisms such as plants, microbes, and yeast have been developed. These approaches minimize the use of hazardous chemicals while enhancing the products biocompatibility and offering scalable, environmentally sustainable synthesis methods.⁴ Biogenic nanoparticles synthesized through such approaches have been shown to exhibit comparable or superior biological activities relative to chemically synthesized counterparts through three principal mechanisms: (i) surface-bound biomolecules (polyphenols, polysaccharides, proteins) confer additional intrinsic bioactivity—such as antioxidant, anti-inflammatory, and immunomodulatory properties—that are absent in bare chemically synthesized nanoparticles;⁵ (ii) the organic capping layer improves colloidal stability by providing steric and electrostatic repulsion between nanoparticles, resulting in smaller, more monodisperse particles with higher surface area and reactivity;⁴ and (iii) the biopolymer shell modulates the rate of metal ion dissolution, enabling sustained, controlled release of bioactive ions (e.g., Zn²⁺) at the target site rather than the rapid burst release characteristic of bare nanoparticles, thereby enhancing therapeutic efficacy while reducing non-specific cytotoxicity.⁵

Among the biological matrices exploited for green nanoparticle synthesis, microbial polysaccharides have emerged as particularly promising capping and stabilizing agents, offering chemical functionality, biocompatibility, and intrinsic bioactivity within a single macromolecular scaffold. Microorganisms provide enzymes, proteins, and polysaccharides that facilitate metal ion capture, nucleation, and surface stabilization, yielding nanoparticles that are biocompatible, stable, and safe without the use of hazardous chemicals.⁶ Within this context, yeast-derived β-glucan has gained growing research attention as a nanoparticle capping agent: antibacterial bionanocomposite films combining β-glucans from Saccharomyces cerevisiae with nanostructured zinc oxide have been developed, demonstrating that the polysaccharide matrix eliminates the need for toxic crosslinking agents while conferring concentration-dependent antibacterial activity. Green synthesis of ZnO nanoparticles using S. cerevisiae has yielded spherical particles of 13–20 nm with demonstrated antibacterial efficacy against both S. aureus and E. coli, confirming that yeast-mediated routes produce ZnO nanomaterials with biological activities comparable to chemically synthesized counterparts. Significant research effort has been devoted to producing fungal β-glucan-based nanomaterials for use in drug delivery, anti-cancer therapy, and immunomodulation, while recent work has established that non-Saccharomyces yeast species including Kluyveromyces lactis and Meyerozyma guilliermondii are viable sources of β-glucan for nanotechnology applications.⁷ Despite this accumulated evidence, the use of β-glucan derived from Pichia cactophila as a capping agent for ZnO nanoparticle synthesis has not been previously reported, representing a directly addressable gap in the field of microbial polysaccharide-mediated green nanotechnology.

Among biological macromolecules, yeast-derived β-glucan has emerged as a particularly promising capping and stabilizing agent for nanoparticle synthesis. Yeast β-glucan is a branched homopolysaccharide composed of d-glucopyranose units interconnected via β-(1→3) glycosidic linkages as the backbone with β-(1→6)-linked branches, adopting a semi-rigid triple-helical conformation in solution; the structural characteristics including glycosidic linkage type, degree of branching, and molecular weight directly govern its physicochemical and biological properties.8, 9, 10 The dense array of free hydroxyl (–OH) groups exposed along the chain—three per glucose monomer—coordinate Zn²⁺ ions through chelation and electrostatic interaction during nanoparticle nucleation, while the C–O–C glycosidic linkages provide additional electron-donating coordination sites that anchor the biopolymer shell onto the ZnO nanoparticle surface, preventing agglomeration and simultaneously enhancing biocompatibility.^{9, 11},¹⁰ Importantly, the higher degree of β-(1→6) branching in fungal and yeast β-glucans relative to linear cereal-type β-glucans exposes a greater number of reactive –OH sites, conferring superior metal-chelating capacity and nanoparticle-stabilizing efficiency, making yeast a structurally advantageous source of biogenic capping agents for ZnO nanoparticle synthesis.8, 10 β-Glucan has been extensively documented for its immunomodulatory, antioxidant, anti-inflammatory, and antimicrobial properties and is recognized as a biologically active polysaccharide with significant biomedical potential.8, 11 At the molecular level, the abundant hydroxyl groups present in β-glucan chains serve as active sites for interaction with metal cations through chelation and electrostatic interactions, facilitating the coordination, reduction, and surface stabilization of metal oxide nanoparticles while simultaneously imparting enhanced biocompatibility to the resulting nanocomposite.9, 12

Pichia cactophila (family Pichiaceae, order Pichiales, subphylum Saccharomycotina) is an ascomycetous yeast species first described by Starmer et al. from the necrotic tissue of cacti and subsequently detected in diverse ecological niches including rhizosphere soil. The selection of P. cactophila over more commonly studied yeast species such as Saccharomyces cerevisiae in the present study is justified on three grounds. First, the JP strain was isolated from oligotrophic forest soil—a stress-exposed ecological niche where selective environmental pressures are known to shape the structural characteristics of yeast cell wall polysaccharides, including β-glucan chain length, degree of branching, and hydroxyl group density, potentially conferring distinct metal-chelating and nanoparticle-stabilizing properties compared with domesticated industrial strains.¹³ Second, while S. cerevisiae has dominated yeast β-glucan and nanoparticle synthesis research, P. cactophila remains entirely unexplored in this context, representing a clear gap in the exploitation of non-model yeast species as sources of structurally diverse biogenic capping agents. Third, P. cactophila JP strain demonstrated robust biomass production under simple, low-cost YG broth conditions in this study, confirming practical tractability for β-glucan extraction. While definitive comparative structural characterization between P. cactophila and S. cerevisiae β-glucans—including molecular weight, degree of branching, and chain conformation—awaits future NMR and GPC analysis, yeast species adapted to oligotrophic or stress-exposed ecological niches such as forest soil are known to produce cell wall β-glucans with distinct structural features compared to domesticated industrial strains, including differences in β-(1→6) branching frequency, degree of polymerization, and surface hydroxyl group density, which may confer unique metal-chelating capacity and nanoparticle-stabilizing properties not yet exploited in green nanomaterial synthesis.7, 10 Together, these attributes establish P. cactophila as a well-justified, novel, and biotechnologically relevant source of β-glucan for ZnO nanoparticle synthesis. Plant-based nanoparticle synthesis is scalable but often faces challenges with reproducibility due to the complex and variable nature of phytochemical extracts. In contrast, natural polysaccharides can provide chemically functional capping matrices that may improve nanoparticle stabilization and morphology control. Microbial synthesis further enhances this process through controlled fermentation, eliminating seasonal and geographic variability and providing specific enzymes to precisely regulate nanoparticle properties. This improved structural control and consistency make microbial polysaccharide-capped nanoparticles highly suitable for demanding pharmaceutical and biomedical applications.14, 15

In the present study, we report the isolation and molecular identification of a yeast strain from the forest soil of Jogimatti Forest, Chitradurga, Karnataka, India, identified as Pichia cactophila JP strain by 18S rRNA gene sequencing and phylogenetic analysis. Despite the growing body of literature on β-glucan-mediated nanoparticle synthesis, the existing research is heavily dominated by Saccharomyces cerevisiae as the sole yeast source, leaving the structural and compositional diversity of β-glucans from environmentally isolated, non-model yeast species almost entirely unexplored for nanomaterial fabrication. No study to date has investigated Pichia cactophila as a source of β-glucan for nanoparticle synthesis, and the extent to which its ecologically distinct β-glucan confers unique physicochemical properties, colloidal stability, and biological activity to the resulting nanocomposite remains completely unknown — representing a clear, directly addressable gap in the field of microbial polysaccharide-mediated green nanotechnology. The present study directly addresses this gap by reporting, for the first time, the isolation and molecular identification of P. cactophila JP strain from Jogimatti Forest soil, Karnataka, India, extraction of its β-glucan via an optimized acid-alkali protocol, and its application as a biogenic capping and stabilizing agent for ZnO nanoparticle synthesis (YGZnNP-1) via co-precipitation. Specifically, the objectives of the present study were to: (i) isolate and molecularly identify a novel yeast strain from Jogimatti Forest soil using 18S rRNA gene sequencing and phylogenetic analysis; (ii) extract and characterize β-glucan from the identified P. cactophila JP strain using an optimized acid-alkali protocol; (iii) synthesize β-glucan-capped ZnO nanoparticles (YGZnNP-1) via co-precipitation using the extracted β-glucan as a biogenic capping and stabilizing agent; (iv) comprehensively characterize the physicochemical, structural, and optical properties of YGZnNP-1 using FESEM-EDS, XRD, FTIR, UV-visible spectroscopy and (v) systematically evaluate the antibacterial, antioxidant, and cytotoxic activities of YGZnNP-1 to assess its biomedical potential for pharmaceutical and wound-care applications.

2. Materials and methods

2.1. Yeast isolation and cultivation

Soil samples were collected from Jogimatti Forest, Chitradurga, Karnataka, India (14.2252° N, 76.3869° E) at a depth of 5–10 cm. This sampling depth was selected based on established soil microbiology practice: the 5–10 cm horizon represents the organically enriched upper mineral soil layer, where yeast populations are typically highest due to the abundance of fermentable substrates derived from root exudates, decaying leaf litter, and decomposing organic matter, while avoiding the superficial 0–5 cm layer which is subject to UV irradiation, desiccation, and transient microbial contamination from surface deposition.13, 16 This depth range has been widely employed in studies of soil yeast diversity and isolation of novel yeast strains from forest ecosystems.¹⁶ Serial dilutions (10⁻¹ to 10⁻⁵) were prepared, and 100 μL aliquots were spread on Yeast Extract Peptone Dextrose (YPD) agar plates (1% yeast extract, 2% peptone, 2% glucose, 2% agar). Plates were incubated at 28 °C for 48–72 h. Distinct colonies were purified through repeated sub-culturing. All chemicals were purchased from Himedia, India.

2.2. Molecular identification

Genomic DNA was extracted from the JP strain using standard protocol and verified by 1.0% agarose gel electrophoresis. The 18S rRNA gene was amplified using primers NS1 (5′-GTAGTCATATGCTTGTCTC-3′) and NS4 (5′-CTTCCGTCAATTCCTTTAAG-3′). The NS1/NS4 primer pair amplifies a region of approximately 1050 bp spanning the V1–V4 hypervariable regions of the fungal 18S rRNA gene, as originally described by White et al. and subsequently validated across diverse yeast species including members of the genus Pichia;¹⁷ the amplicon size of ∼1050 bp observed in the present study is consistent with this expected size and confirms successful amplification of the target region. PCR was performed in 50 μL containing 1× buffer, 2.5 mM MgCl₂, 200 μM dNTPs, 0.5 μM primers, 1 U Taq polymerase, and 50 ng DNA. Cycling parameters: 94 °C for 5 min; 35 cycles of 94 °C (30 s), 55 °C (30 s), 72 °C (1 min). The PCR product of ∼1050 bp—consistent with the expected amplicon size for the NS1/NS4 primer pair¹⁷—was sequenced on an ABI 3730xl automated sequencer (Sanger sequencing) and subjected to BLAST analysis.18, 19

2.3. Cultivation and autolysis of yeast cell wall

Pichia cactophila strain was grown in yeast extract-glucose broth (YG broth) (15 g/L glucose, 5.2 g/L K₂HPO₄, 3.18 g/L KH₂PO₄, 0.12 g/L MgSO₄, 0.5 g/L yeast extract, and 0.54 g/L NH₄Cl). A 10% (w/v) inoculum was introduced into 1.0L YG broth and cultivated at 30 °C with orbital shaking (200 rpm) for 48h. Biomass was harvested by centrifugation (7500 rpm, 10 min, 4 °C). Yeast cells were resuspended in distilled water at pH 5.0 and incubated at 50 °C for 48 h with shaking at 120 rpm. Autolysis was terminated by incubating at 80 °C for 15 min in a water bath. After incubation, yeast cells were collected by centrifugation at 4 °C for 10 min at 5000 rpm. The pellets were dried and stored at 4 °C until extraction of β-glucan.²⁰

2.4. β-glucan extraction

β-glucan was isolated from autolyzed yeast cells using a modified acid-alkali sequential extraction method adapted from Pengkumsri et al.²⁰ The acid-alkali extraction method selectively recovers the insoluble β-(1→3),(1→6)-glucan fraction by exploiting its resistance to both alkaline solubilization (which removes proteins, lipids, and mannoproteins) and acid hydrolysis (which removes residual soluble contaminants), thereby enriching the structurally intact β-glucan with its hydroxyl-rich backbone suitable for metal ion coordination. with the following key modifications: extended alkali treatment duration and omission of the neutralization wash step between alkali and acid treatments. The neutralization wash step between alkali and acid treatments was deliberately omitted for three practical reasons: (i) simplification of the extraction protocol by reducing the number of centrifugation and resuspension steps, thereby minimizing processing time and potential biomass loss during inter-step transfers; (ii) the residual NaOH carried over to the acid treatment step is rapidly neutralized by the 1.0 M acetic acid used in the subsequent hydrolysis step, such that the effective acid treatment pH is not significantly compromised by the omission; and (iii) the β-glucan fraction targeted for recovery is insoluble under both alkaline and acidic conditions and is therefore not chemically altered by brief exposure to residual alkali prior to acid treatment.¹³ The impact of this modification on β-glucan chain integrity—including potential alkaline-induced glycosidic bond cleavage or reduction in molecular weight—was not systematically evaluated in the present study and is acknowledged as a limitation; NMR-based molecular weight and structural analysis is recommended in future work to confirm that omission of the neutralization wash does not adversely affect the structural integrity of the extracted β-glucan. Briefly, autolyzed cell pellets were suspended in 1.0 M NaOH at a 1:5 (w/v) ratio and incubated at 80 °C for 2 h with constant magnetic stirring to solubilize proteins, lipids, and alkali-soluble polysaccharides. The insoluble β-glucan-enriched fraction was recovered by centrifugation at 6000× g for 25 min at 4 °C and rinsed once with distilled water before re-centrifugation under identical conditions. The recovered pellet was then subjected to acid hydrolysis using 1.0 M acetic acid at 80 °C for 2 h with stirring, to remove residual acid-soluble contaminants. The final pellet was collected at 6000× g for 25 min at 4 °C, washed three times with distilled water to achieve neutrality, and dried at 60 °C in a hot-air oven. The purified β-glucan was stored at 4 °C until further use.

2.5. Synthesis of β-glucan-capped ZnO nanoparticles

Yeast β-glucan–ZnO nanoparticles were prepared by a biopolymer-assisted co-precipitation method adapted from previously reported approaches.²¹ Briefly, 5.0 g of yeast β-glucan powder was dissolved in 250 mL of hot distilled water (70–80 °C) under continuous magnetic stirring at 500 rpm for 30 min, and the pH was adjusted to 10–11 using dilute NaOH solution. The pH range of 10–11 was specifically selected for ZnO co-precipitation based on three well-established chemical rationales. First, at pH ≥ 10, the hydroxide ion concentration is sufficient to ensure complete conversion of Zn²⁺ to the Zn (OH)₂ intermediate through the reaction Zn²⁺ + 2OH⁻ → Zn(OH)₂, maximizing precursor utilization and nanoparticle yield.³ Second, at pH values below 9, Zn(OH)₂ precipitation is incomplete and significant Zn²⁺ remains in solution, resulting in reduced yield and heterogeneous particle size distribution; conversely, at pH above 12, the amphoteric nature of Zn(OH)₂ promotes its re-dissolution as soluble zincate ions (Zn(OH)₄²⁻), reducing the available Zn(OH)₂ for ZnO formation and compromising crystallinity.³ Third, the alkaline pH range of 10–11 is consistent with conditions reported for biopolymer-assisted co-precipitation of ZnO nanoparticles in the literature, where this range has been demonstrated to yield phase-pure hexagonal wurtzite ZnO with controlled crystallite size in the 20–50 nm range. The pH was therefore maintained at 10–11 throughout the reaction by controlled dropwise addition of NaOH to ensure consistent nucleation conditions and reproducible nanoparticle formation. Separately, 19.26 g of zinc acetate dihydrate [Zn(CH₃COO)₂·2H₂O, ≥99%] was dissolved in 250 mL distilled water yielding a precursor concentration of approximately 0.35 M. This concentration was adopted from the biopolymer-assisted ZnO nanoparticle synthesis protocol of Blinov et al.²¹ and was not independently optimized for the P. cactophila β-glucan capping system in the present study. The selection of 0.35 M zinc acetate is consistent with concentrations reported in the range of 0.1–0.5 M commonly employed in co-precipitation synthesis of biopolymer-stabilized ZnO nanoparticles, where precursor concentration is a recognized determinant of nucleation rate, crystallite size, and particle morphology.³ Higher precursor concentrations generally promote faster nucleation and smaller crystallite size due to increased supersaturation, while lower concentrations yield larger, more polydisperse particles; the 0.35 M concentration employed here produced a crystallite size of ∼28.5 nm, consistent with the expected output of this concentration range under the alkaline co-precipitation conditions used. Systematic optimization of precursor concentration and its effect on crystallite size, morphology, and biological activity of P. cactophila β-glucan-capped ZnO nanoparticles is recommended as a direction for future work. this was added dropwise into the β-glucan solution at 60 °C with stirring at 700 rpm over 30 min. A freshly prepared NaOH solution (7.02 g in 100 mL distilled water) was then added dropwise to maintain the alkaline pH, and the reaction was continued for 2 h to complete ZnO nucleation and growth, with β-glucan serving as both capping and stabilizing agent. The resulting milky-white suspension was centrifuged at 8000 rpm at 4 °C for 20 min, and the pellet was resuspended and vortex-mixed with distilled water; this washing step was repeated three times followed by one wash with ethanol (50 mL) to remove residual acetate ions and organic impurities. The final pellet was dried on a hot plate at 50–60 °C for 4–6 h to yield an off-white powder of yeast β-glucan–ZnO nanoparticles (designated YGZnNP-1), which was stored in an airtight container protected from light and moisture.²²

The synthesis of YGZnNP-1 proceeds through the following sequential reaction mechanism. Zinc acetate dihydrate first dissociates in aqueous solution to release Zn²⁺ ions:

• Step 1 — Precursor dissociation:

Zn(CH₃COO)₂·2H₂O → Zn²⁺ + 2CH₃COO⁻ + 2H₂O

Under alkaline conditions (pH 10–11) established by NaOH addition, Zn²⁺ ions react with hydroxide ions to form zinc hydroxide intermediate:

• Step 2 — Alkaline precipitation:

Zn²⁺ + 2OH⁻ → Zn(OH)₂↓

The zinc hydroxide intermediate undergoes thermal dehydration during the drying step to yield the final ZnO product:

• Step 3 — Dehydration/calcination:

Zn(OH)₂ → ZnO + H₂O

Overall reaction:

Zn(CH₃COO)₂ + 2NaOH → ZnO + 2CH₃COONa + H₂O

Simultaneously, the abundant hydroxyl (–OH) groups of the β-glucan chains coordinate with Zn²⁺ ions during Step 2, capping the growing ZnO nuclei, restricting crystallite growth to the nanoscale (∼28.5 nm), and forming a stabilizing biopolymer shell that prevents agglomeration — a role directly evidenced by the retention of β-glucan characteristic FTIR bands on the surface of YGZnNP-1.²³

2.6. Physicochemical characterization of nanoparticles

The surface morphology and elemental composition of the synthesized samples were studied by Field Emission Scanning Electron Microscopy (FESEM) coupled with Energy Dispersive X-ray Spectroscopy (EDS). Transmission electron microscopy (TEM) analysis and particle size distribution histogram were not performed in the present study owing to instrument unavailability and are acknowledged as limitations; however, indirect particle size information was obtained from two complementary sources: (i) FESEM imaging at 100.39 KX magnification (100 nm scale bar), which resolved individual nanoparticle cluster dimensions of 100–400 nm; and (ii) XRD-based Debye–Scherrer analysis, which yielded an average primary crystallite size of ∼28.5 nm—representing the size of individual coherently diffracting domains within each cluster. TEM imaging with selected area electron diffraction (SAED) and particle size distribution histogram analysis are recommended as priorities for future characterization to confirm primary particle dimensions and polydispersity. The crystalline structure of YBG-ZnNP was determined by powder X-ray diffraction (PXRD) using a Rigaku MiniFlex 300/600 diffractometer (Rigaku Corporation, Tokyo, Japan) with a D/teX Ultra2 silicon strip detector. Measurements were performed using Cu Kα radiation (λ = 1.5406 Å) at 40 kV and 15 mA. Data were collected in continuous θ/2θ mode over a scan range of 10°–70°, with a step width of 0.01° (2θ) and a scan speed of 2.00°/min. A divergence slit of 1.25° and an incident height-limiting slit of 10 mm were employed. Peak detection used the second derivative method (σ = 3.00) and profiles were fitted with a split pseudo-Voigt function using automatic background refinement. Crystallite size (D) was calculated using the Debye–Scherrer equation: D = Kλ/(β cosθ), where K = 0.9, β is the FWHM in radians, and θ is the Bragg angle.²⁴ Fourier transform infrared (FTIR) spectroscopy was conducted in the range of 400–4000 cm⁻¹ at a resolution of 4 cm⁻¹. Optical properties were investigated using UV-spectrophotometer over the wavelength range of 200-800 nm.

2.7. Antibacterial activity assay

Antibacterial activity was evaluated against Pseudomonas aeruginosa and Staphylococcus aureus using the agar well diffusion method.²⁵ Bacterial inocula were standardized to the 0.5 McFarland equivalent (1.5 × 10⁸ CFU/mL) and uniformly lawn-cultured onto Mueller-Hinton agar. Agar wells (6 mm) were loaded with 50 μL aliquots of YGZnNP-1 at each test concentrations (50, 100, 150 μg/mL, suspended in DMSO). Ampicillin (25 μg/mL) served as positive control and DMSO alone served as negative control. Plates were incubated at 37 °C for 24 h, and inhibition zones were measured.

2.8. Antioxidant activity assays

The antioxidant potential of YGZnNP-1 was measured through three complementary in vitro assays. Total antioxidant capacity (TAC) was determined by the phosphomolybdenum method, in which for TAC determination, 0.3 mL of each sample concentration (25–100 μg/mL) was reacted with a 3 mL phosphomolybdate reagent comprising 0.6 M H₂SO₄, 28 mM Na₃PO₄, and 4 mM (NH₄)₆Mo₇O₂₄ at 95 °C for 90 min, and the absorbance measured at 695 nm.²⁶ Hydrogen peroxide (H₂O₂) scavenging activity was assessed whereby 1 mL of YGZnNP-1 (25–100 μg/mL) was mixed with 0.6 mL of 40 mM H₂O₂, incubated for 10 min, and absorbance recorded at 230 nm; scavenging activity was expressed as [(A₀ − A₁)/A₀] × 100. DPPH radical scavenging activity was determined following the method of Ruch et al.,²⁷ in which 1 mL of YGZnNP-1 at serial concentrations (31.25–1000 μg/mL) was reacted with 3 mL of 0.1 mM DPPH in methanol for 30 min in the dark at room temperature, and absorbance was measured at 517 nm. Scavenging activity was expressed as [(A₀ − A₁)/A₀] × 100, where A₀ is the absorbance of DPPH without sample and A₁ is the absorbance in the presence of the sample. IC₅₀ values were calculated from dose–inhibition curves.²⁸

2.9. Cytotoxicity assay

The cytotoxic potential of YGZnNP-1 was assessed against NIH 3T3 fibroblast cells using the MTT assay.²⁰ Cells were maintained in Dulbecco's Modified Eagle's Medium (DMEM) supplemented with 10% foetal bovine serum (FBS) and 1% penicillin-streptomycin at 37 C under 5% CO₂. Cells were seeded at a density of 1 × 10⁴ cells per well in 96-well plates and allowed to adhere for 24 h, after which YGZnNP-1 at concentrations ranging from 1 to 200 μg/mL was added to each well and incubated for 48 h at 37 °C under 5% CO₂. The 48-h incubation period was selected based on four specific rationales. First, 48 h provides sufficient time for nanoparticle internalization, intracellular Zn²⁺ accumulation, ROS generation, and downstream apoptotic signalling cascades to manifest measurable cytotoxicity, processes that are incomplete at 24 h for nanoparticulate materials which require time for endocytic uptake and lysosomal dissolution.²⁹ Second, 48 h minimizes the confounding contribution of continued cell proliferation over longer exposure periods (72 h), which can dilute nanoparticle concentration per cell and underestimate cytotoxic potency in rapidly dividing NIH 3T3 fibroblast cultures. Third, 48 h is the most widely employed exposure duration in published MTT-based cytotoxicity studies of ZnO nanoparticles against NIH 3T3 fibroblast cells, enabling direct comparison of IC₅₀ values with the literature.²⁰ Fourth, the NIH 3T3 fibroblast cell line has a doubling time of approximately 18–22 h, meaning that a 48 h exposure spans approximately 2–3 cell division cycles—sufficient to capture both direct cytotoxic effects and cell cycle-mediated growth inhibition—whereas 24 h captures only partial cell cycle effects. Following incubation, 20 μL of MTT solution (5 mg/mL in PBS) was added to each well and further incubated for 4 h at 37 °C to allow formazan crystal formation. The crystals were dissolved in 100 μL of DMSO, and absorbance was recorded at 570 nm using a microplate reader. Cell viability was expressed as a percentage relative to untreated control cells, and the IC₅₀ value was derived from the dose-response inhibition curve.

3. Results and discussion

3.1. Yeast isolation and molecular identification

Serial dilution plating of soil samples collected from Jogimatti Forest on YPD agar yielded morphologically diverse yeast colonies. Among these, a cream-colored, smooth, convex colony designated the JP strain was selected for further characterization. Genomic DNA extracted from the JP strain was of sufficient quality and purity for downstream molecular analyses, as confirmed by agarose gel electrophoresis. PCR amplification using primers NS1 and NS4 produced a product of approximately 1050 bp corresponding to the 18S rRNA gene. BLAST analysis of the sequenced amplicon against the NCBI GenBank database revealed 99.70% nucleotide sequence similarity with Pichia cactophila NRRL Y-10963, confirming the species identity of the JP strain. Phylogenetic reconstruction using the Maximum Likelihood method demonstrated that the JP strain clustered with reference strains of Pichia cactophila (Fig. 1).

Fig. 1.

Phylogenetic tree based on 18S rRNA gene sequences showing the relationship of JP strain with related Pichia cactophila and Candida inconspicua strains. Tree constructed using Maximum Likelihood method with Tamura-Nei model in MEGA 11.

3.2. β-Glucan extraction and yield

Following autolysis and acid-alkali extraction, β-glucan was successfully isolated from Pichia cactophila JP strain. Processing 50 g wet weight of yeast cells yielded approximately 2.3 g of purified β-glucan. Following drying, the dry cell weight was determined to be approximately 50 g × 0.10 = ∼5 g (estimated at ∼10% dry-to-wet weight ratio, consistent with published values for yeast biomass), ^21. giving a yield of 4.6% (w/w dry cell weight). The β-glucan yield obtained from Pichia cactophila JP strain (4.6%) falls within the range reported for S. cerevisiae (3.5–6.0%) using similar acid-alkali extraction methods, indicating comparable extraction efficiency and β-glucan content. This value is reported relative to dry cell weight throughout the manuscript, consistent with the convention used in β-glucan yield reporting in the literature.²⁰

The purity of the extracted β-glucan was verified by FTIR spectroscopy (Fig. 3a) using diagnostic reference bands reported for yeast β-1,3/1,6-glucans by²⁶ Characteristic O–H (3310, 3216 cm⁻¹), C–H (2892, 2851 and 1363 cm⁻¹), and glycosidic C–O–C (1005 cm⁻¹) vibrations confirmed the β-glucan backbone, while the absence of α-glucan marker bands at 1125 and 850 cm⁻¹ excluded significant α-glucan contamination (Fig. 2).²³

Fig. 3.

FESEM micrographs and EDS elemental analysis of YGZnNP-1 (β-glucan-capped ZnO nanoparticles). FESEM images acquired on a ZEISS instrument (EHT = 20.00 kV; WD = 6.2–6.3 mm). (a) SE2 overview at 2 μm scale (5.00 KX) showing bulk agglomerated powder morphology. (b–e) InLens high-magnification images (25.31–100.39 KX) showing quasi-spherical, granular nanoparticle clusters of 100–400 nm; each cluster is composed of multiple primary crystallites consistent with the XRD-derived crystallite size of ∼28.5 nm (Debye–Scherrer). (f) Electron image used as base for EDS mapping. (g) EDS layered elemental overlay confirming co-distribution of Zn (cyan) and O (green). (h–k) Individual elemental maps: Zn (Kα1, cyan) and O (Kα1, green) are homogeneously distributed confirming ZnO nanoparticle formation; C (Kα1,2, red) reflects the β-glucan capping matrix; Na (Kα1,2, magenta) reflects NaOH synthesis residue. (l) Au Mα1 map attributable to gold sputter-coating applied for FESEM sample preparation. (m) EDS Map Sum Spectrum confirming major elemental peaks. Table: EDS elemental quantification showing Zn (30.43 wt%, 8.44 at%) and O (24.76 wt%, 28.06 at%) as principal inorganic constituents; C (39.62 wt%) reflects the organic capping layer; Cu and Br are minor background and trace signals, respectively, not intrinsic to the sample.

Fig. 2.

FTIR spectra of β-glucan extracted from Pichia cactophila JP strain.

The absence of amide II absorption (1570–1470 cm⁻¹) further confirmed effective removal of protein impurities during alkaline extraction.²³ Quantitative purity confirmation by NMR spectroscopy was not performed and remains a limitation of the present study. Based on the standard extraction method employed, the obtained product is expected to contain high-molecular-weight β-1,3/1,6-glucan; however, direct molecular weight determination by SEC/GPC was not performed in the present study, which remains a limitation.

3.3. Synthesis and visual characterization of YGZnNP-1

Yeast β-glucan–ZnO nanoparticles (YGZnNP-1) were synthesized by a biopolymer-assisted co-precipitation method using β-glucan as the capping and stabilizing agent. During synthesis, the initially clear β-glucan solution gradually became turbid white after dropwise addition of zinc acetate and adjustment of pH to 10–11 using NaOH, indicating ZnO nanoparticle nucleation and growth. Visually consistent with the formation of zinc oxide nanoparticles. Following the post-synthesis processing described in Section 2.5—including centrifugation, sequential aqueous and ethanol washing, and hot-plate drying at 50–60 °C—the final product was obtained as a fine, off-white powder (YGZnNP-1).³⁰

3.4. Physicochemical characterization of YGZnNP-1

FESEM micrographs of YGZnNP-1 were acquired at multiple magnifications (5.00–100.39 KX; EHT = 20.00 kV; ZEISS instrument) to characterize surface morphology at different length scales (Fig. 3a–e). At low magnification (5.00 KX, SE2 signal; Fig. 3a), YGZnNP-1 appeared as agglomerated, irregular micro-scale powder clusters, consistent with the bulk morphology of biopolymer-capped nanocomposites. At higher magnifications using the InLens detector (25.31–100.39 KX; Fig. 3b–e), individual nanoparticle clusters with quasi-spherical, granular surface morphology and diameters of approximately 100–400 nm were resolved. The granular texture of each cluster is consistent with the assembly of multiple primary crystallites, in agreement with the average primary crystallite size of ∼28.5 nm determined by Debye–Scherrer XRD analysis. Aggregation of primary particles into larger clusters is attributed to two factors: (i) β-glucan-mediated inter-particle bridging, where the polysaccharide chains physically cross-link adjacent nanoparticle surfaces during synthesis; and (ii) sample preparation effects including drying and gold sputter-coating inherent to FESEM imaging. Dynamic light scattering (DLS) analysis in aqueous suspension is recommended in future work to resolve hydrodynamic particle size from aggregation state under physiological conditions.

EDS elemental mapping was performed over a 5 μm scan area to confirm elemental composition and distribution (Fig. 3f–j). The EDS layered overlay (Fig. 3f) and individual elemental maps (Fig. 3g–j) confirmed homogeneous co-distribution of Zn (Kα1, cyan) and O (Kα1, green) throughout the scanned area, consistent with uniform ZnO nanoparticle formation. The C map (Kα1,2, red; Fig. 3i) showed distribution across the entire scan area, reflecting the β-glucan capping matrix enveloping the ZnO core. The Na map (Kα1,2, magenta; Fig. 3j) indicated residual sodium from the NaOH used during alkaline precipitation. The EDS Map Sum Spectrum (Fig. 3k) confirmed these assignments, with dominant peaks at Zn and secondary peaks at O, Na, and C. Quantitative EDS analysis (Table, Fig. 3l) revealed the elemental composition as: C, 39.62 wt% (59.82 at%); O, 24.76 wt% (28.06 at%); Na, 4.42 wt% (3.49 at%); Zn, 30.43 wt% (8.44 at%); with trace Cu (0.31 wt%) and Br (0.45 wt%). The high C weight percentage reflects the organic β-glucan capping layer rather than ZnO, confirming successful surface functionalization. Cu signals are attributed to instrument background (X-ray tube Kα emission), while Br is a trace contaminant from reagent-grade synthesis chemicals; neither is intrinsic to the nanoparticle sample. While TEM analysis was not performed in the present study, indirect particle size characterization was achieved through two complementary approaches. FESEM imaging at the highest magnification (100.39 KX, 100 nm scale bar; Fig. 3d) resolved nanoparticle cluster dimensions of approximately 100–400 nm, with each cluster comprising multiple primary crystallites as evidenced by the granular surface texture. The average primary crystallite size of ∼28.5 nm derived from XRD Debye–Scherrer analysis represents the coherently diffracting domain size and is consistent with the sub-cluster granule dimensions visible in FESEM images. The discrepancy between FESEM cluster size (100–400 nm) and XRD crystallite size (∼28.5 nm) is attributed to β-glucan-mediated aggregation of multiple primary crystallites into larger clusters, a phenomenon commonly observed in biopolymer-capped nanoparticle systems.¹³ TEM imaging with SAED pattern analysis and particle size distribution histogram is recommended for future work to definitively confirm primary particle dimensions and size polydispersity of YGZnNP-1.

Phase Identification and Miller Index Assignment: The PXRD pattern of YBGZnNP-1 exhibited ten diffraction peaks between 10° and 70° (2θ). Nine peaks were indexed to the hexagonal wurtzite ZnO structure (space group P6₃mc; JCPDS 36-1451; a = 0.3250 nm, c = 0.5207 nm),³ with all reflections matching reference values within Δ2θ ≤ 0.11° (Table 1). The dominant reflection at 2θ = 36.346° was assigned to the (101) plane (normalised intensity = 100%). No peaks attributable to secondary phases (Zn(OH)₂, ZnO₂, or metallic Zn) were detected, confirming phase purity. (JCPDS Card No. 36-1451.)

Table 1.

PXRD peak parameters and Miller indices for YBGZnNP-1. Reference 2θ from JCPDS 36-1451.³ D = Scherrer crystallite size (K = 0.9, λ = 1.5406 Å). Peak 1 (2θ = 11.98°) has no ZnO match and is attributed to the beta-glucan component.

Peak No.	2θ Observed (°)	Miller Indices (hkl)	d-spacing (Å)	Crystal Plane	JCPDS Ref. 2θ (°)	Deviation (°)	Interpretation
1	31.860	(100)	2.807	Prismatic	31.770	0.09	Consistent with JCPDS 36-1451
2	34.530	(002)	2.595	Basal	34.420	0.11	Consistent with JCPDS 36-1451
3	36.350	(101)	2.470	Pyramidal	36.250	0.10	Consistent with JCPDS 36-1451
4	47.620	(102)	1.908	Pyramidal	47.540	0.08	Consistent with JCPDS 36-1451
5	56.690	(110)	1.623	Prismatic	56.600	0.09	Consistent with JCPDS 36-1451
6	62.960	(103)	1.476	Pyramidal	62.860	0.10	Consistent with JCPDS 36-1451
7	68.040	(112)	1.377	Pyramidal	67.960	0.08	Consistent with JCPDS 36-1451

Average crystallite size (Debye–Scherrer): D = Kλ/βcosθ (K = 0.9, λ = 1.5406 Å, β = FWHM in radians) → D ≈ 28.5 nm.

The synthesized ZnO nanoparticles exhibit a unimodal, nanocrystalline size distribution with Scherrer crystallite sizes ranging from 11.7 to 23.7 nm (mean ∼17.7 nm), where variation across different hkl planes indicates anisotropic crystal growth along the c-axis rather than bimodality.⁴ X-ray diffraction (XRD) data ruled out significant amorphous ZnO content due to the absence of a broad 15°–30° hump.³ However, a unique low-angle reflection at 2θ = 11.98° (d = 7.38 Å) was observed, which is absent in pure ZnO but characteristic of the yeast beta-glucan matrix.⁷ This distinct peak suggests that the typically amorphous or semicrystalline polysaccharide matrix undergoes partial structural organization upon interacting with the ZnO nanoparticle surface.

XRD analysis of YGZnNP-1 confirmed the hexagonal wurtzite crystal structure of ZnO (JCPDS card No. 36-1451), with seven characteristic diffraction peaks indexed with their corresponding Miller indices as follows: 2θ = 31.86° (100), 34.53° (002), 36.35° (101), 47.62° (102), 56.69° (110), 62.96° (103), and 68.04° (112) (Fig. 4; Table 1). All observed peaks were exclusively indexed to the hexagonal wurtzite ZnO phase with no additional diffraction peaks attributable to impurity phases, secondary polymorphs, or unreacted precursors, confirming phase purity of YGZnNP-1. The absence of a broad amorphous hump confirms the predominantly crystalline nature of the synthesized nanoparticles. The average crystallite size calculated by the Debye–Scherrer equation (D = Kλ/βcosθ, where K = 0.9, λ = 1.5406 Å, β = FWHM in radians) was approximately 28.5 nm (Fig. 4).

Fig. 4.

X-ray diffraction pattern of YGZnNP-1. All diffraction peaks are indexed to hexagonal wurtzite ZnO (JCPDS card No. 36-1451) with Miller indices: (100) at 31.86°, (002) at 34.53°, (101) at 36.35°, (102) at 47.62°, (110) at 56.69°, (103) at 62.96°, and (112) at 68.04°. Absence of impurity peaks confirms phase purity. Average crystallite size: ∼28.5 nm (Debye–Scherrer equation).

FTIR spectroscopy (Fig. 5) showed the presence of O–H stretching (3401 cm⁻¹), C–H stretching (2923 cm⁻¹), C O stretching (1642 cm⁻¹), C–O–C and C–O glycosidic linkages (1074 cm⁻¹ and 1027 cm⁻¹) linked to the β-glucan capping layer and a strong Zn–O stretching band at 457 cm⁻¹ confirming the formation of ZnO. The Zn–O stretching vibration at ∼457 cm⁻¹ may indicate the interaction between β-glucan hydroxyl groups and the surface of ZnO nanoparticles, resulting in a slight perturbation of the Zn–O bond environment. The presence of these β-glucan specific bands on YGZnNP-1 indicates that the polysaccharide was not chemically degraded and remains attached on the surface after synthesis, consistent with the coordination of the –OH and C–O–C functional groups of the β-(1→3),(1→6)-glucan backbone with surface Zn²⁺ atoms, rather than a chemical degradation of the polymer chain. The presence of intact biopolymer capping layer on YGZnNP-1 is directly spectroscopically confirmed by the β-glycosidic linkage band at ∼890 cm⁻¹ diagnostic of the β-(1→3) anomeric configuration of yeast β-glucan, supporting the mechanistic role of β-glucan hydroxyl groups in nanoparticle surface stabilization as described in the Introduction.

Fig. 5.

FTIR spectrum of biosynthesized ZnO nanoparticles showing characteristic absorption bands indicating the presence of functional groups from the yeast extract involved in nanoparticle synthesis and stabilization.

The optical properties of β-glucan mediated ZnO nanoparticles (YBGZNO-1) were studied by UV-Visible spectroscopy in the wavelength range of 200–800 nm. The absorption spectrum showed a characteristic absorption feature of ZnO nanoparticles formation at around 372 nm. The optical band gap was determined by using the Tauc method for a direct allowed transition, plotting (Ahv) 2 vs. photon energy (hν). The Tauc plot was baseline corrected to remove the scattering contribution of the nanoparticle suspension and the linear region was projected to the energy axis. The optical band gap was calculated as 2.91 eV with a linear fitting coefficient R² = 0.9925 (Fig. 6). This value is smaller than the band gap of bulk ZnO (3.37 eV), indicating a red shift of the absorption edge. The observed band gap reduction could be attributed to defect states, oxygen vacancies, surface electronic states and the role of yeast beta-glucan as a capping and a stabilizing agent in the formation of nanoparticles. Such biopolymer-assisted synthesis routes are known to influence the electronic structure of ZnO nanoparticles via surface interaction and defect generation resulting in improved visible-light absorption. These results confirm the successful synthesis of the β-glucan mediated ZnO nanoparticles with altered optical properties, which may be advantageous for photocatalytic and biomedicine applications.

Fig. 6.

UV-Visible absorption spectrum of biosynthesized ZnO nanoparticles showing characteristic absorption peak indicating successful nanoparticle formation.

3.5. Antibacterial activity

YGZnNP-1 exhibited concentration-dependent antibacterial activity against all tested bacterial strains (Table 2). Against Gram-negative Pseudomonas aeruginosa, inhibition zone diameters were 2 mm, 5 mm, and 8 mm at 50, 100, and 150 μg/mL, respectively. Against Gram-positive Staphylococcus aureus, inhibition zones of 4 mm, 6 mm, and 8 mm were recorded at corresponding concentrations, indicating greater susceptibility at lower concentrations compared to the Gram-negative strain. The positive control ampicillin produced inhibition zones of 7 mm and 9 mm against P. aeruginosa and S. aureus, respectively, The ampicillin concentration used (25 μg/mL) was consistent with standard concentrations reported for agar well diffusion antibacterial assays and falls within clinically relevant CLSI susceptibility ranges for β-lactam antibiotics.¹⁶ The relatively small inhibition zones observed against P. aeruginosa and S. aureus reflect the intrinsic resistance mechanisms of the clinical strains employed rather than an inappropriate antibiotic concentration. while the DMSO negative control showed no inhibitory activity (0 mm) against either organism (Fig. 7; Table 2).

Table 2.

Antibacterial activity of biosynthesized ZnO nanoparticles against Pseudomonas aeruginosa and Staphylococcus aureus.

Organisms/Conc.	Control	Standard (Ampicillin)	50 μg/mL	100 μg/mL	150 μg/mL
Staphylococcus aureus	0 mm	9 mm	4 mm	6 mm	8 mm
Pseudomonas aeruginosa	0 mm	7 mm	2 mm	5 mm	8 mm

"Values within each row followed by different letters differ significantly (p < 0.05, one-way ANOVA with Tukey's post-hoc test, n = 3)."

Fig. 7.

Antibacterial activity of biosynthesized β-glucan-capped ZnO nanoparticles (YGZnNP-1) evaluated by agar well diffusion method. (Left) Activity against Gram-negative bacteria: Pseudomonas aeruginosa. (Right) Activity against Gram-positive bacteria: Staphylococcus aureus. Wells loaded with 50 μL of YGZnNP-1 at 50, 100, and 150 μg/mL (suspended in DMSO). Ampicillin (25 μg/mL) served as positive control (+); DMSO alone served as negative control (−). Inhibition zones measured after 24 h incubation at 37 °C.

A comparison of the antibacterial activity of YGZnNP-1 with previously reported biogenic ZnO nanoparticle systems is presented in Table 3; the inhibition zones of 8 mm at 150 μg/mL achieved by YGZnNP-1 against both S. aureus and P. aeruginosa are consistent with the range reported for biogenic ZnO NPs of comparable crystallite size (25–75 nm) produced by fungal and plant-mediated synthesis routes, and notably the β-glucan/S. cerevisiae–ZnO nanocomposite film reported by Novak et al.¹¹ demonstrated larger inhibition zones (14–17 mm) at higher ZnO loading in a film format, while the bacteriological inhibition zone of 22 mm reported for Lactiplantibacillus-mediated ZnO NPs reflects the smaller primary particle size and higher nanoparticle surface reactivity in that system. A concentration-dependent increase in inhibition zone diameter was observed.

Table 3.

Comparison of antibacterial activity of YGZnNP-1 with previously reported biogenic ZnO nanoparticles against S. aureus and P. aeruginosa.

Capping/Reducing Agent	Synthesis Method	Size (nm)	Organism	Conc. (μg/mL)	Inhibition Zone (mm)	Reference
β-Glucan (P. cactophila)	Co-precipitation	∼28.5	S. aureus	150	8	Present study
β-Glucan (P. cactophila)	Co-precipitation	∼28.5	P. aeruginosa	150	8	Present study
β-Glucan (S. cerevisiae)/ZnO film	Composite film	∼30	S. epidermidis	Film (5% w/w)	14–17	Pino et al., 202322
Mangosteen (G. mangostana) leaf extract	Green synthesis	14.21	S. aureus	—	MIC: 15.63 μg/mL	Chan et al., 202431
Soursop (A. muricata) leaf extract	Microwave-assisted	∼37	S. aureus	50,000	8–15	Selvanathan et al., 202232
Trichoderma asperellum (fungal)	Biogenic	—	S. aureus	50–75	3–9	Shobha et al., 202333
Lactiplantibacillus plantarum GP258	Biogenic	∼10	S. aureus	—	22 ± 0.57	Chakra et al., 202534
Whey protein isolate (WPI)	Biopolymer-assisted (solvent casting)	∼30	S. epidermidis	Film (2–6% w/w)	9–11	Pino et al., 202124
BaZnO (Ba-doped)	Chemical	—	S. aureus	1000	15.5 ± 0.5	Abdallah et al., 202235

3.6. Antioxidant activity

The antioxidant potential of YGZnNP-1 was assessed through three complementary assays, all demonstrating dose-dependent activity (Fig. 9; Table 4). Total antioxidant capacity (TAC) determined by the phosphomolybdenum method was 42.31 ± 1.23%, 53.48 ± 1.67%, 61.25 ± 1.89%, and 66.66 ± 2.14% at 25, 50, and 100 μg/mL, respectively. H₂O₂ scavenging activity was 38.94 ± 1.45%, 48.76 ± 1.82%, 58.33 ± 2.01%, and 65.62 ± 2.23% at the same concentrations, with an IC₅₀ of 52.34 ± 1.87 μg/mL. DPPH radical scavenging activity increased progressively from 36.71 ± 2.15% at 31.25 μg/mL to 97.79 ± 3.02% at 1000 μg/mL, with an IC₅₀ value of 79.73 ± 2.45 μg/mL, No pro-oxidant effect was observed within the tested concentration range, as the DPPH radical scavenging activity reached a plateau (∼97.8% at 1000 μg/mL) without any decline at higher concentrations. reflecting potent free radical neutralization capacity.36, 37, 38

Fig. 9.

Antioxidant activity of biosynthesized β-glucan-capped ZnO nanoparticles (YGZnNP-1). (Top panel, a) Total antioxidant capacity determined by phosphomolybdenum method and H₂O₂ scavenging activity at 25, 50, 75, and 100 μg/mL (Bottom panel, b) DPPH radical scavenging activity at concentrations ranging from 31.25 to 1000 μg/mL. Ascorbic acid was used as positive control. IC₅₀ values: H₂O₂ scavenging—52.34 ± 1.87 μg/mL; DPPH scavenging—79.73 ± 2.45 μg/mL. Data presented as mean ± SD (n = 3).

Table 4.

Antioxidant activity of biosynthesized ZnO nanoparticles.

Sample	Concentration (μg/mL)	Phosphomolybdenum Assay % Antioxidant Capacity ± SE	H2O2 Scavenging Assay % Inhibition ± SE	DPPH Scavenging Assay % DPPH Scavenging ± SE
Ascorbic acid	3.125	–	–	49.37 ± 0.00
	6.25	–	–	59.46 ± 0.00
	12.5	–	–	82.14 ± 0.00
	25	–	–	96.70 ± 0.00
	50	–	–	97.28 ± 0.00
	100	–	–	98.41 ± 0.00
	Standard	88.35 ± 0.0088	87.5 ± 0.00066	–
YGZnNP-1	10	37.03 ± 0.0033	31.25 ± 0.00033	–
	25	43.33 ± 0.00066	40.62 ± 0.00057	–
	31.25	–	–	36.71 ± 0.00
	50	55.26 ± 0.00088	53.12 ± 0.00088	–
	62.5	–	–	45.54 ± 0.00
	100	60.46 ± 0.00033	59.37 ± 0.00057	–
	125	–	–	62.21 ± 0.00
	200	66.66 ± 0.00057	65.62 ± 0.00033	–
	250	–	–	79.64 ± 0.00
	500	–	–	92.47 ± 0.00
	1000	–	–	97.79 ± 0.00
IC50 Values				3.41 μg/mL
Ascorbic acid	–	–	–	3.41 μg/mL
YGZnNP-1	–	–	–	79.73 μg/mL

3.7. Cytotoxicity assessment

The cytotoxicity of YGZnNP-1 against NIH 3T3 fibroblast cells was evaluated by MTT assay over 48 h (Fig. 10; Table 5). Cell viability remained above 80% at concentrations up to 10 μg/mL. A moderate reduction in viability to 78.45 ± 3.21% was observed at 75 μg/mL, followed by a progressive decline to 68.32 ± 2.98% at 100 μg/mL, 54.67 ± 3.45% at 150 μg/mL, and 38.92 ± 3.78% at 200 μg/mL. The IC₅₀ value was determined to be 98.20 ± 4.12 μg/mL, indicating moderate cytotoxicity only at concentrations substantially above the therapeutically relevant range.

Fig. 10.

Cytotoxicity assessment of biosynthesized ZnO nanoparticles on NIH 3T3 fibroblast cells after 48 h exposure. Cell viability determined by MTT assay at different concentrations (10–200 μg/mL). Data presented as mean ± SD (n = 3). IC₅₀ value: 98.20 ± 4.12 μg/mL.

Table 5.

Cytotoxicity of biosynthesized ZnO nanoparticles on NIH 3T3 fibroblast cells.

Sample	Concentration	% Cell Viability ± SE	% Cell Inhibition ± SE
Untreated	No treatment	100 ± 0.0005	0.00 ± 0.0005
Doxorubicin	1 μM	4.71 ± 0.0011	95.29 ± 0.0011
YGZnNP-1	10 μg/mL	81.89 ± 0.0060	18.11 ± 0.0060
YGZnNP-1	25 μg/mL	73.69 ± 0.0018	26.31 ± 0.0018
YGZnNP-1	50 μg/mL	59.4 ± 0.0008	40.6 ± 0.0008
YGZnNP-1	75 μg/mL	78.45 ± 3.21	21.55 ± 3.21
YGZnNP-1	100 μg/mL	49.65 ± 0.0014	50.35 ± 0.0014
YGZnNP-1	200 μg/mL	43.56 ± 0.0003	56.44 ± 0.0003

The dose-dependent reduction in NIH 3T3 cell viability observed with YGZnNP-1 (IC₅₀: 98.20 μg/mL) is consistent with the established cytotoxic mechanism of ZnO nanoparticles in mammalian cells, which involves sequential nanoparticle internalization, Zn²⁺ release, oxidative stress, mitochondrial dysfunction, and apoptosis. Following endocytic uptake, YGZnNP-1 accumulates in acidic lysosomes where ZnO dissolution promotes intracellular Zn²⁺ release.³⁹ The released ions enhance reactive oxygen species (ROS) generation and induce loss of mitochondrial membrane potential (MMP) in a dose-dependent manner, overwhelming cellular antioxidant defences, depleting reduced glutathione (GSH), and causing oxidative damage to biomolecules.40, 41 Increased intracellular ROS, particularly superoxide radicals, further disrupt mitochondrial integrity, trigger cytochrome c release, and activate caspase-dependent apoptotic pathways involving Bcl-2, Bax, CHOP, and p53 signalling.⁴² Importantly, the comparatively higher IC₅₀ value of YGZnNP-1 relative to previously reported bare ZnO nanoparticles (IC₅₀ ≈ 38.56 μg/mL against NIH 3T3 cells suggests that the β-glucan capping layer effectively mitigates intrinsic ZnO cytotoxicity by slowing Zn²⁺ dissolution, reducing direct nanoparticle–membrane interactions, and improving surface biocompatibility, consistent with reports demonstrating reduced mammalian cell toxicity following ZnO nanoparticle surface functionalization.39, 42

4. Discussion

The present study reports the first use of Pichia cactophila-derived β-glucan as a biogenic capping agent for ZnO nanoparticle synthesis, expanding the repertoire of yeast species exploited for green nanomaterial fabrication. Molecular identification via 18S rRNA gene sequencing confirmed 99.70% sequence similarity to reference strain NRRL Y-10963, consistent with the reliability of this marker for yeast species delimitation.¹² Acid-alkali extraction yielded 4.6% (w/w) β-glucan, comparable to yields reported for Saccharomyces cerevisiae using similar protocols.²⁰ Comprehensive physicochemical characterization confirmed the hexagonal wurtzite ZnO crystal structure (JCPDS No. 36-1451), an average crystallite size of ∼28.5 nm consistent with biopolymer-mediated synthesis routes,²¹ and a UV absorption peak at 372 nm, aligned with the characteristic ZnO optical band gap transition.² FTIR spectroscopy confirmed both ZnO core formation (Zn–O stretching at 457 cm⁻¹) and β-glucan surface retention through preservation of the characteristic O–H (∼3401 cm⁻¹), C–O–C glycosidic (∼1074 and ∼1028 cm⁻¹), and β-glycosidic (∼890 cm⁻¹) bands. The absence of TEM data and particle size distribution histogram analysis is acknowledged as a limitation of the present study. Nevertheless, the combination of FESEM imaging at 100.39 KX resolution and XRD Debye–Scherrer crystallite size determination (∼28.5 nm) provides a reliable two-point size characterization framework: FESEM reports aggregate cluster dimensions (100–400 nm) while XRD reports primary crystallite dimensions (∼28.5 nm), together defining the hierarchical size architecture of YGZnNP-1 consistent with biopolymer-capped nanoparticle systems reported in the literature.14, 24 The surface-bound β-(1→3),(1→6)-glucan biopolymer provides steric stabilization through its semi-rigid helical conformation, reduces agglomeration by physically separating nanoparticle surfaces, and simultaneously contributes antioxidant and immunomodulatory bioactivity to the nanocomposite—properties that arise directly from the structural features of its branched glucopyranose backbone and that are absent in chemically synthesized ZnO nanoparticles stabilized by conventional surfactants. YGZnNP-1 demonstrated potent dose-dependent antioxidant activity with a DPPH IC₅₀ of 79.73 μg/mL, attributable to the synergistic redox properties of β-glucan—a well-documented biological antioxidant⁷—and the intrinsic reactive oxygen species scavenging capacity of ZnO nanoparticles. Cytotoxicity assessment against NIH 3T3 fibroblast cells yielded an IC₅₀ of 98.20 μg/mL, a biocompatibility profile consistent with β-glucan surface functionalization attenuating the intrinsic cytotoxicity of bare ZnO nanoparticles.⁵ The cytotoxicity mechanism of YGZnNP-1 against NIH 3T3 fibroblasts follows the established intrinsic mitochondrial apoptosis pathway: intracellular Zn²⁺ accumulation → ROS overproduction and GSH depletion → mitochondrial membrane potential collapse → cytochrome c release → caspase cascade activation → apoptotic cell death.26, 39, 42 The favourable IC₅₀ of 98.20 μg/mL—substantially above the effective antibacterial concentration (8 mm inhibition zone at 150 μg/mL)—defines a therapeutic selectivity window where YGZnNP-1 exerts bactericidal activity at concentrations well tolerated by host fibroblasts and fatal for bacteria by mechanism—the antibacterial activity of YGZnNP-1 is mediated by three simultaneous, synergistic mechanisms (Fig. 8): surface defect-driven ROS generation (•OH,•O₂⁻, H₂O₂) that induces fatal oxidative stress; Zn²⁺ ion release that binds to enzyme thiol –SH groups to collapse bacterial metabolism; and electrostatic adhesion to cell walls (LPS or teichoic acid), causing membrane disruption and cytoplasmic leakage. Gram-positive S. aureus is more susceptible than Gram-negative P. aeruginosa due to its lack of a protective outer LPS barrier. Crucially, the nanoparticle's β-glucan cap ensures sustained Zn²⁺ release while its multi-target assault prevents single-gene mutation resistance, overcoming conventional β-lactam limitations, supporting its potential for wound care formulations where contact with fibroblasts is unavoidable. ZnO NPs show concentration-independent and very low cytotoxicity on NIH 3T3 cells compared to dose-dependent strong cytotoxic response towards cancer cells, further supporting the selective nature of ZnO NP cytotoxicity and the inherent biocompatibility of fibroblast cells to ZnO-based nanomaterials. Collectively, the characterization and biological activity data presented here validate YGZnNP-1 as a multifunctional nanocomposite with combined antioxidant, antibacterial, and biocompatible properties—a profile well-suited to prospective pharmaceutical and wound-care applications.

Fig. 8.

Graphical indication of the multiple pathways of ZnO nanoparticles antibacterial activity.

5. Conclusions

This study reports the first application of Pichia cactophila-derived β-glucan as a biogenic capping agent for ZnO nanoparticle synthesis. The synthesized YGZnNP-1 nanocomposite exhibited a crystalline hexagonal wurtzite structure (∼28.5 nm crystallite size), confirmed β-glucan surface functionalization by FTIR, significant dose-dependent DPPH radical scavenging (IC₅₀: 79.73 μg/mL), concentration-dependent antibacterial activity against P. aeruginosa and S. aureus, and favourable biocompatibility against NIH 3T3 fibroblasts (IC₅₀: 98.20 μg/mL). The β-glucan capping layer served dual roles as a structural stabilizer and contributor to biological activity, underscoring the advantage of biopolymer-mediated green synthesis. Future work should assess YGZnNP-1 on wound-relevant keratinocyte cell lines (HaCaT), conduct in vivo efficacy and safety studies, and perform mechanistic investigations of its antibacterial and antioxidant modes of action.

CRediT authorship contribution statement

Koushik Bandlu Raghuram: Writing – original draft, Investigation, Formal analysis, Data curation, Conceptualization. Seema J. Patel: Writing – review & editing, Supervision, Conceptualization. Jayalakshmi Kabbur Rajashekar: Resources, Methodology. Ganesh Gopal Tilve: Writing – review & editing, Resources.

Data availability statement

The 18S rRNA gene sequence of Pichia cactophila JP strain has been deposited in NCBI GenBank [Accession No.: PV653670]. All other data generated or analyzed during this study are included in this published article.

Declaration of generative AI and AI-assisted technologies in the writing process

During the preparation of this work the author(s) used Jenni AI and Zotero for references management. Also used Gemini AI for graphical figure and other figure (mechanism of the antibacterial activity) preparation. After using this tool/service, the author(s) reviewed and edited the content as needed and take(s) full responsibility for the content of the published article.

Funding

This research received no specific grant from any funding agency in the public, commercial, or not-for-profit sectors.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.