Physical and chemical methods of synthesizing metal nanoparticles are expensive, have negative impacts, and are hazardous. Simpicilliumlanosoniveum, an endophytic fungus found in the seaweed Sargassumwightii, was used in an attempt to synthesize silver nanoparticles (brown seaweed). Ultraviolet-Visible Spectroscopy, Field-Emission Scanning Electron Microscopy, Atomic Force Microscopy, X-ray Diffraction Spectroscopy, Fourier Transform Infrared Spectroscopy, and Dynamic Light Scattering were used to characterize silver nanoparticles. Antioxidant, antimicrobial, Chick chorioallantoic membrane, and cytotoxic experiments were performed on the nanoparticles. The spherical nanoparticles had an average particle size of 20–25 nm, as determined by field-emission scanning electron microscopy. The nanoparticles' size and crystalline composition was also confirmedby X-ray Diffraction Spectroscopy.The nanoparticles demonstrated potent antioxidant, antibacterial, antiangiogenic, and cytotoxic activity. Hence, we believe that the green silver nanoparticles might be usefulin managing microbial infections and cancers.
Research Article
Biosynthesis of silver nanoparticle using endophytic fungus (Simpicillium lanosoniveum) isolated from Sargassum wightii and its biological activity
https://doi.org/10.21203/rs.3.rs-3456948/v1
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Silver nanoparticle
Endophytic fungus
Simpicilliumlanosoniveum
Sargassumwightii. Antiangiogenic activity
Antimicrobial activity
Molecules around 1–100 nanometers in size are considered nanomaterials[1]. They area remarkable class of toolsin great demand for various practical uses. Of late, theyare exploited in biomedicine, pharmaceuticals, biosensors, cosmetics, food technology, electronics, optical devices, dye degradation, and wastewater treatment [2].Metal nanoparticles (MNPs) havedistinctive physicochemical properties due to their enormous specific surface area and high surface atom density.They are involved in catalysis, optical, magnetic, and electrical properties, and have antimicrobial activity[3]. Different metal salts, such as silver, gold, platinum, zinc, copper, and iron, are utilized to synthesize various nano-molecules[4]. However, silver nanoparticles (AgNPs) are preferred over other nanomaterialsowingto their biocompatible nature. The favorable attributes of AgNPs include their optical property, high electrical conductivity, and biological characteristics[5]. AgNPs are increasingly being used in antibacterial, biosensor, food-related, and chemo-preventive applications. The AgNPs are produced using a variety of physical, chemical, and biological methods. Although physical and chemical approaches to nanomaterial synthesis are rapid, they have significant drawbacks, including the need for expensive chemicals and equipment and have adverse side effects too [6]. These limitations of the orthodox approach forced scientists to explore a safe, eco-friendly, cost-effective process forsynthesizing nano-metals. One such method is creating nanomaterials utilizing reducing and capping agents from biological sources[7]. AgNPs produced biologically exhibit high yields, excellent solubility, stability, and antibacterial characteristics[8]. "Green synthesis" is the use of many biological sources, such as plants, microorganisms, animals, and marine materials.Because they are simpler to handle than other natural products, microorganisms are preferred[9]. Diverse microorganisms from terrestrial and marine environments have been used in the synthesis of an array of AgNPs[10].
The process of creating metal nanoparticles (MNPs) utilizing fungus biomass and products is known as myco-nanotechnology. A large surface area for interactions is provided mainly through the cell wall of fungal mycelia, which facilitates the absorption and reduction of metallic ions to generate MNPs[11]. Nanocrystals can be produced using fungi's internal or extracellular components. Proteins, enzymes, and cellular metabolites all contribute to intracellular synthesis[12]. Proteins containing cysteine are involved when NADH/NADPH-dependent reductase enzymes produce metal ions for extracellular use[13].
Additionally, it has been discovered that fungi allow significant polydispersity, stable structures, and diversity in the AgNPs dimension. Filamentous fungi are preferred for producingAgNPs,asthey release more enzymes and secondary metabolites[14].So, the employment of fungi in the creation of nonentities is becoming more popular.
All marine plants, including algae, seagrass, driftwood, mangrove plants, marine vertebrates, mainly fish, and marine invertebrates, sponges, and coral, have been found to harbor marine endophytic fungus that donot show any outward symptoms of disease[15]. Seaweeds have been shown to be a superior source of endophytic fungus. Due to their ability to thrive in unique biological niches, endophytic fungi connected with seaweed have distinctive secondary metabolites[16]. The scientific community was inspired to look for new compounds due to the bioactive secondary metabolites from seaweed endophytes exhibiting anti-algal, anticancer, antimicrobial, anti-plasmodial, antioxidant, insecticidal, andAChE modulation activities[17].
To our knowledge, very little research has used marine endophytic fungi to produce gold, silver, and copper nanoparticles[11]. Recently, we synthesized the bioactive gold nanoparticles and isolated Penicilliumcitrinum and Cladosporiumcladosporiodes from Sargassumwightii[18]. The current study's objective was to create AgNPs using the endophytic fungus S. lanosoniveum from the seaweed S. wighttii (brown seaweed). These myco-nanoparticles demonstrated potential cytotoxic, antioxidant, and antibacterial properties.
2.1. Cultivation of Microorganisms
The endophytic fungus was isolated from S. wightiifollowingAriffinet al. S. wightii was carefully cleansed in distilled water, tap water, and seawater to flush out salts and other materials to get rid of epiphytic germs. The seaweed was surface-sterilized for one minute using 70% ethyl alcohol following three minutes of submersion in a 4% sodium hypochlorite solution. It was rinsed in distilled water before drying on sterile tissue paper,after which itwas cleaned with sterile distilled water and aseptically cut into 5cm segments. The sterile seaweed fragments were deposited on potato dextrose agar (PDA) containing chloramphenicol.The plates were maintainedat 28°C during mycelial growth. A pure endophytic fungal culture was prepared, sub-cultured on PDA (without antibiotics), and then extracted after 30 days at 28°C [19].
2.2. Molecular Identification
The genomic DNA was isolated by freezing 100 mg of fungal mycelia in liquid nitrogen, followed by mechanical disruption. Using the NucleoSpin® Plant II Kit and manufacturer's instructions (Macherey-Nagel), the genomic DNA was extracted. The ITS region of the 18S rRNA gene was amplified using fungal domain-specific primers:ITS-1F "GGAAGTAAAAGTCGTAACAAGG" (forward primer) and ITS-4R "TCCTCCGCTTATTGATATGC" (reverse primer). Amplification was carried out in a PCR thermal cycler (GeneAmp PCR System 9700, Applied Biosystems) using a 5µL reaction mixture with 1µL template DNA. The reaction mixture was initially denatured at 98°C for 30 s, followed by 40 cycles at 98°C for 5 s, 58°C for 10 s, 72°C for 15 s, and a final extension at 72°C for 1 min. The amplification was assessed by electrophoresis of the amplified sample in a 1.2 percent w/v agarose gel. The amplified DNA was extracted and purifiedusing a gel extraction kit[20]. The sequence was examined with BLASTn and NCBI; then, it was classified using a homology search for closely identical sequences. MEGAX software created a phylogenetic tree using the neighbour-joining technique, and multiple sequences were aligned using ClustalW2.
2.3. Preparation of Fungal Submerged Fermentation Broth
The S. lanosoniveumendophyte was cultured in a 1000 ml flask containing 500 ml of potato dextrose broth (PDB), which was maintained at 28°C for 7 days to establish the fungus biomass. Following incubation, filtering was used to separate the fungal biomass. The fungal biomass pellets were gathered and rinsed three-four times in water to get rid of any connected medium components. This mycelial biomass was cultured for 3 days at 28°C, while still being shaken in a suspension of 10 g in 100 ml of sterile water. The generation of nanoparticles was improved by using the cell filtrate[21].
2.4. Synthesis of Silver Nanoparticles
In a 250 mL Erlenmeyer flask, 50 mL of silver nitrate (AgNO3) and 10 mL of S. lanosoniveum filtrate were combined and incubated for 3 h at room temperature. A control flask containing 50 ml of AgNO3 solution but with no biomass was likewise incubated at the predetermined temperature for 3 h. After incubation, the solution was spun up at 10,000x g for 30 min to extract the AgNPs. Following three rinses with deionized distilled water, the extracted AgNPs were vacuum-processed at 50°C[22].
2.5. Optimization Studies for Silver Nanoparticle Production
Using a UV-Visible spectrophotometer UV-1800 (Shimadzu, Kyoto, Japan), the reduction of pure silver ions in the reaction medium was optimized for several factors, including AgNO3 concentration, duration, and pH.
2.5.1. Effect of Silver Nitrate on Silver Nanoparticles Synthesis
The quantity ofthe substrate affects the quality of AgNPs. S. lanosoniveum filtrate was mixed with a range of AgNO3 (1–5 mM) at pH 8 to determine the impact of metal salt on the synthesis of AgNPs at 30°C for 3 h.
2.5.2. Effects of Reaction Time on Silver Nanoparticles Synthesis
Studies on optimization have been done using different time intervals between 0 and 5 h. AgNO3 was kept at a 5 mM concentration. The pH and temperature of the reaction mixture were maintained at 8 and 30°C, respectively.
2.5.3. Effect of pHon Silver Nanoparticles Synthesis
The synthesis of AgNPs is impacted by the pH of the medium, which also affects the proliferation of endophytic fungus and the production of fungal enzymes. Therefore, the pH of the reaction mixture was chosen in the range of 2–12 to explore the impact of pH on AgNP production. Freshly made buffers (pH 2–12) were used to change the pH without changing the AgNO3 concentration. The pH of the reaction mixture was checked using a pH meter. AgNO3 was maintainedat a 5 mM concentration for the duration of the 3h incubation at 30⁰C.
2.6. Characterization of silver nanoparticles
2.6.1. Ultraviolet-visible (UV–vis) spectrophotometry
The formation of AgNPs was confirmed by subjecting them to a UV-Visible spectral analysis. An absorption spectrum was monitored over the range of 300–800 nm to monitor the biosynthesis of AgNPs[23].
2.6.2. Field-emission scanning electron microscopy (FE-SEM)
FE-SEM (Hitachi S-4 500) was employed to see the particulate size and surface topography.TheAgNPssample was dropped onto a copper grid coated with carbon to create thin films, and any leftover solution was blotted away using a tissue paper. The film was dried on the SEM grid for 5 min using a mercury lamp[24].
2.6.3. Atomic force microscopy (AFM)
The AgNPs solution was spin-coated onto the glass slide to get the samples ready for atomic force microscopy. The slides were examined by AFM after being air-dried for 12 h in a chamber at 28°C. AgNPs' surface morphology, size, and shapes were all analysed using AFM[25].
2.6.4. Dynamic Light Scattering (DLS)
The dynamic light scattering apparatus measured the size of AgNPs (Zetasizer Nano-2590, Malvern Instruments Ltd, Worcestershire, UK). To investigate DLS, 3 ml of the material was placed in a polystyrene cuvette[26].
2.6.5. Energy-dispersive X-Ray (EDX)
By focusing the electron beam on the spot on the silicon wafer-coated surface, the EDX spectrum was captured in spot-profile mode. Synthesized materials were submitted to an EDS microanalysis system, which automatically recognizes and labels the elements responsible for the peaks in this energy distribution to confirm their elemental composition[27].
2.6.6. X-ray diffraction (XRD)
The crystalline nature of the AgNPs was studied in an X-ray diffractometer, as explained by Mehta et al. "The sample was coated onto an XRD grid, and the spectra were recorded, operated at a voltage of 40 kV and a current of 30 mA with Cu Ka radiation. The domain size was calculated from the width of the XRD peaks". In order to determine the nature of the AgNPs, Scherrer’s formula was used as follows[28],
$$D=\frac{K\lambda }{\beta Cos\theta }$$
Where,
“D” is the average crystallite domain size perpendicular to the reflecting planes,
“λ” is the wavelength,
β = full width at half maximum (FWHM),
θ = diffraction angle.
2.6.7. Fourier-Transform Infrared Spectroscopy
The absorption bands in the fungal extracts responsible for AgNP reduction, capping, and stability were detected using an FTIR spectrophotometer (Thermo Nicolet Avatar). The S.lanosoniveum filtrate extracted with ethyl acetate and AgNPs dried at room temperature were analyzed using KBr optics in the range of 500–4000 cm− 1[29].
2.7. Antioxidant Activity
A modified version of the Brand-Williams et al.’s method[30] was used to evaluate the S. lanosoniveumfiltrate, AgNPs, and standard ascorbic acid's in-vitro antioxidant activities. The stable DPPH-free radical's propensity to react with hydrogen donors is the fundamental characteristic of the DPPH test. Ascorbic acid was used as the reference material, and samples were combined with 0.5 ml of a 0.15 mM DPPH solution at various concentrations (20, 40, 60, 80, and 100 g/mL),following which the mixture was incubated for 15 min. Methanol was employed as the blank solution, whereas DPPH served as the control. In a microplate reader, the absorbance was measured at 517 nm (iMark, Bio-Rad). The capacity of the samples to scavenge DPPH radicals was calculated using the following formula.
$$\% \text{S}\text{c}\text{a}\text{v}\text{e}\text{n}\text{g}\text{i}\text{n}\text{g} \text{P}\text{o}\text{t}\text{e}\text{n}\text{t}\text{i}\text{a}\text{l}=\frac{\text{X}\text{c}-\text{X}\text{s}}{\text{X}\text{c}}$$
Where Xs = Absorbance of the sample,andXc = Absorbance of the control.
2.8. Antimicrobial Activity
Disc diffusion method was used to test the antimicrobial activity ofEscherichia coli (MTCC 118), Staphylococcus aureus (MTCC 7443), Bacillus subtilis (MTCC 441), Pseudomonas aeruginosa (MTCC 424), and Aspergillusniger (MTCC 281). The fungus was maintained on Czapek yeast extract agar, while the bacterial cultures were cultured on nutrient agar (NA) media. The wells created in the petri plates containing the microorganisms were filled with 50 L each of the S. lanosoniveum filtrate, AgNPs, and mycelium-free extract. Ampicillin (1 mg/mL) and fluconazole (1 mg/mL) were the corresponding positive controls for the bacterial and fungal strains [31].
2.9. Chick Chorioallantoic Membrane (CAM assay)
The CAM assay was carried out in accordance with Muslim et al. The eggsfertilized athatcheries were randomly divided into two groups and incubated for 96 h at 37°C with 40–60% humidity. After 7 days, a window (1.5 cm2) was made in the shell to display a section of the CAM. A 100 g of fungal extract was placed on sterile Whatman's filter paper No. 1 discs before being applied to the CAM. After 24 h of incubation, the number of new vessels in the CAM around the paper disc was counted. CAM samples were fixed in 10% formalin, embedded in paraffin, cut into sections, and finally stained with hematoxylin and eosin for histo-pathological investigation.
2.10. Cytotoxicity
To ascertain the harmful effects of AgNPs, the standard 3-(4, 5-dimethyl-thiazolyl-2)-2, 5-diphenyltetrazolium bromide (MTT) assay was employedin the cell viability test. In 96-well plates with 100 mL of cell culture media (DMEM supplemented with 10% FBS and 1% penicillin-streptomycin solution), HeLa and MCF-7 cells were planted at a density of 5,000 cells per well. In a CO2 incubator, cells were cultured for 24 h at 37°C and 5% CO2. Fresh media containing AgNPs (25–250 g/mL) was added to the wells after 24 h, which werethen incubated at 37°C for 24–48 h. After replacing the old medium with 100 µL of fresh medium, 10 µL of MTT solution (5 mg/mL in DMSO) was added to each well.The plates had further two hours of incubation to assess cell viability. After discarding the MTT solution, 100 mL of DMSO was added to each well, and the wells were then allowed to incubate for 40 min in the dark. A microplate reader measured the solution's absorbance at 492 nm [32].
2.11. Statistical Analysis
Threemean and standard deviations of the independent datawere used to express all experimental outcomes. Analysis of variance (ANOVA) was used in the statistical analysis of the data, which was carried out using Origin 8.0 program.
Nanotechnology prefers the biosynthesis of inorganic nanomaterials employing physiologically active extracellular agents, such as proteins, enzymes, or other organic molecules[33]. Recently, extra- or intracellular components of microorganisms have been used to synthesize inorganic nanoparticles. The nanoparticles produced by the fungi are easily distinguishable from the microbial biomass and are ready for use right away with no additional purification. Due to a large number of biologically active secretory enzymes present in the extracellular components of the filamentous fungi, these components are widely employed in the creation of inorganic nanomaterials[34]. We have recently synthesized the silver nanoparticles using Penicilliumcitrinum[21] and Cladosporiumcladosporides[23]. In this study, we used the endophytic fungus S. lanosoniveum isolated from S. wightiito examine the extracellular production of AgNPs and their antioxidant, antimicrobial and cytotoxic activity.
Sequence analysis of the ITSrDNA region was used to identify the fungi, andthe genomic DNA sequence was entered into the NCBI Gen Bank database (KY075854). The isolated fungus was recognized as S. lanosoniveum based on the characteristics of the colony, microscopic findings, and sequence similarity analyses. MEGA X software was used to generate phylogenetic relationships (Fig. 1) illustrating the taxonomic link between the endophytes known from diverse sources.
3.1. Biosynthesis,Optimization, and Characterization of silver nanoparticles
3.1.1. Ultraviolet-Visible Spectroscopy
With the addition 4 ml of S. lanosoniveum filtrate to an aqueous AgNO3 solution, the color of the mixture shifted from yellowish brown to black, indicating the synthesis of AgNPs.Surface Plasma Resonance (SPR)gave the reaction mixture (Fig. 2) its dark color. A characteristic SPR band in the 430 nm region was visible in AgNPs.The effect of pH, time, and AgNO3 concentration on AgNPs formation was assessed. The maximum AgNPs yield was obtained at 5mM AgNO3. Figure 3a shows that high concentration ofAgNO3leads to increased absorption peak intensity. The synthesis of AgNPs was also found to depend on the reaction time.
The highest amount of AgNPswas produced at 3h and 30 min of incubation, as illustrated in Fig. 3b, and incubation beyond 3h and 30 minresulted in a reduction inthe absorption peak. AgNPs production was boosted as pH increased, and maximum synthesis was seen at pH 9 (Fig. 3c). The pH of the reaction system has been hypothesized to impact AgNP size and form[35].The results of our investigation showed that acidic conditions are unsuitable for the formation of AgNPs.The peak's strengthwasreducedwhenthe acidity of the medium increased. The acidic medium suppresses theformation of the AgNP nuclei, whichmay lead to a decrease in the amount of AgNPs[36].
In contrast, the reaction medium's alkaline pH promotes the production of AgNPs[37]. Silver oxides (Ag2O) are produced when unbound Ag+ reacts with basic sodium hydroxide, increasing the peak of both AgNPs and Ag2O, as demonstrated by Liu et al.[38]. The alkaline state of the reaction medium in this investigation may have contributed to the highest AgNPs production.
3.1.2. Field-emission scanning electron microscopic (FE-SEM)
The surface morphology of AgNPs was uniform in shape and spherical, as observed in theFE-SEM images (Fig. 4). The range of the particle size histogram in the current investigation is 20 to 25 nm. This finding firmly established the role of S. lanosoniveum filtrate as a reduction and capping mediatorduring the synthesis of AgNPs.Similar outcomes were reportedon AgNPssynthesized using beetroot extract[39].
3.1.3. Atomic force microscopy (AFM)
The advantages of atomic force microscopy include high resolution,andthe samples do not need to be conductive nor need high-pressure vacuum conditions. It is a tool to analyse the size, shape, and agglomeration pattern, and provides visualizations of three-dimensional views of the nanoparticles[40]. AgNPs (Fig. 5) were studied with AFM at 4 m resolution, and the particles were found to be polydispersed, spherical, and had a surface roughness of 40 nm. They did not aggregate together.
3.1.4. Dynamic Light Scattering (DLS)
The DLS size distribution image of AgNPs is shown in Fig. 6. AgNPs were hydrodynamic sized between 40 and 90 nm, andhad an average size of 70 nm. AgNPs' particle size was smaller in the DLS analyser spectrum than in the UV-vis spectrum.
3.1.5. Energy-dispersive X-Ray (EDX)
Due to SPR for metallic AgNPs, the energy-dispersive spectra of the nanoparticles displayed (Fig. 7) a conspicuous peak at 3 keV.The quantitative data of AgNPsdepicted other elements, such as O and C.The presence of O and C elements in EDX is attributed to the proteins in the extract, which handled the reduction and capping of silver ions. Similar observations were reported by Hulikere et al[18].
3.1.6. X-ray diffraction (XRD)
AgNPs' XRD pattern, shown in Fig. 8, supports their crystalline nature. The (1 1 1), (2 0 0), (2 2 0), and (3 1 1) reflection planes of the face-cantered cubic structure of silver can be indexed to the four distinctive diffraction peaks at 2θ values of 38.30, 44.29, 64.46, and 77.54, respectively[41].
3.1.7. Fourier-Transform Infrared Spectroscopic (FTIR)
The S. lanosoniveumfiltrate and AgNPs were subjected to Fourier transform infrared studies in order to pinpoint the potential functional groups.FTIR spectra of dried aqueous S. lanosoniveum filtrate, AgNO3, and AgNPs are shown in Fig. 9. The phytochemical analysis of S. lanosoniveum filtrate revealed the presence of flavonoids, alkaloids, steroids, and tannins. In S. lanosoniveum filtrate, the peaks were observed at 1075, 1631, 1958, 2320, and 3338cm− 1,and in AgNO3, they were observed at 1643, 2046, 2131, and 3348cm− 1. After the reaction of S. lanosoniveum filtrate with AgNO3, the peaks were shifted to a higher wave number side, such as 1079, 1620, 1946, 2133, 2343, and 3330cm− 1.The O-H or N-H stretching vibration of alcohol and phenolic groups caused the broad peak at 3330cm− 1[42]. The O-H stretching of carboxylic acids was suggested as the cause of the band at 2343cm− 1.Absorbance peaks at 1643cm− 1 were due to the symmetric stretching and vibrations of C = O groups of amino acid residues in proteins and the amide I group [43].The phenol group's O-H stretching was observed at 1079 cm− 1[44]. According to the current data, flavonoids and proteins may be responsible for the rapid decrease and capping of AgNPs. Strong reducing agents found in the flavonoids in the S. lanosoniveum filtrate may indicate that AgNPs were formed by reducing AgNO3.
3.2. Antioxidant activity
Figure 10 displays the antioxidant activity of AgNPs, S. lanosoniveum filtrate, and ascorbic acid. The antioxidant potential of AgNPswas found to be dependent on concentration. However, the silver nanoparticlesshowed greater DPPH radical scavenging activities (63.20% ±0.12%) than the S. lanosoniveum filtrate (46.50% ±0.15%), indicating that the AgNPs had better proton-donating ability and may act as good antioxidants[45].
3.3. Antimicrobial activity
Five distinct human pathogenic bacteria were used to study the antimicrobial impact of AgNPs (Table 1). The average diameter of the zone of inhibition for S. aureus, P. aeruginosa, B. subtilis, E. coli, and A. niger was 13, 12, 10, 11, 8.5, and 9.3 mm, respectively (50 g/mL). AgNPs were reported to have the strongest antibacterial efficacy against P. aeruginosa. These results are consistent with earlier research[46] that looked at the antibacterial action of AgNPs against P. aeruginosa. According to reports, the antibacterial activity of AgNPs is caused by the bacterium being penetrated; the cell membrane is damaged, and the cell contents are released. Another hypothesis puts out the release of silver ions from the nanoparticles, which may help explain how AgNPs are bactericidal[47].Interestingly, the AgNPs synthesized using S. lanosoniveumextractexhibited significant antimicrobial activity against Gram-positive and negative, and the fungi highlighted the opportunity to use these AgNPs as broad-spectrum antimicrobial agents.
Table 1: Antibacterial activity of silver nanoparticle synthesized using Simpicilliumlanosoniveumfungal filtrate against the pathogenic bacteria and fungi.
3.4. Chick chorioallantoic membrane (CAM assay)
Using a chorioallantoic membrane (CAM) experiment, the fungal extract's in-vivo antiangiogenic efficacy was assessed. CAM treated with S. lanosoniveum filtrate had (10.0 ± 0.37) fewer blood vessels than the control (18.0 ± 0.90). The AgNPs' ability to inhibit angiogenesis was demonstrated by the reduction in the number of blood vessel branches on CAM (Fig. 11).
3.5. Cytotoxicity
AgNPs were tested for toxicity against the cancer cell lines HeLa and MCF-7 in the cytotoxic investigation. HeLa and MCF-7 cancer cell lines' proliferation was found to be decreased by AgNPs, and the extent of cytotoxicity was dependent on the concentration. AgNPs' IC50 against HeLa cells was shown to be 200 g/mL (Fig. 12).
Using the marine endophytic fungus S. lanosoniveum from S. wightii, we successfully produced silver nanoparticles. UV-vis, FTIR, AFM, FESEM, XRD, DLS, and EDX analyses were employed to characterize the AgNPs produced by the green synthesis method. The produced AgNPs had a size of less than 100 nm and significantly increased antioxidant, antibacterial, and cytotoxic properties. It has been established that the proteins and phenolic chemicals in the extracts create these nanostructures. The current study's findings can be further investigated in order to develop novel broad-spectrum antibacterial agents and formulations to treat diseases caused by radicals, including infections, inflammatory conditions, and cancer.
ACKNOWLEDGMENT
The authors acknowledge Mangalore University, India, for facilities, and Dr. A.P. Lipton, Former Head-In-Charge, Central Marine Fisheries Research Institute, Vizhinjam, Thiruvananthapuram, India, for help in the collection and identification of seaweeds.
AUTHOR CONTRIBUTIONS
ManjunathHulikere M: Investigation, formal analysis, data curation, writing- review and editing, project administration and funding acquisition. B Dinesh: Writing original draft, software, validation, formal analysis, data curation, and writing-review and editing. Chandrashekhar G. Joshi: Conceptualization, methodology, investigation, validation, formal analysis, data curation, resources, writing-review and editing, supervision and funding acquisition. All authors have read and approved the manuscript.
DATA AVAILABILITY STATEMENT
The data relevant to the findings of the study are available from the corresponding author upon reasonable request.
Ethical Approval–Not applicable
Funding – Not applicable
Informed Consent– None.
Consent for Publication– All the authors agreed to publish the data in this journal.
Research Involving Humans and Animals Statement– None.
Competing Interests– The authors declare no competing interests.
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No competing interests reported.
