INTRODUCTION 
Antibiotic resistance is considered one of the most critical worldwide threats to humanity in the 21st century. Reducing the effectiveness of antimicrobial treatments has created a need for novel designs and the development of antibacterial agents [1] . However, the improper use and excessive intake of antibiotics increase the development of antibacterial resistance which allows pathogenic bacteria to create drug-resistant [2]. One of the key factors leading to antimicrobial resistance (AMR) is bacterial biofilm formation, which supplies the pathogens with a protective shield against antibiotics. In reality, Biofilm development is another method that bacteria might use to survive when antibiotics are present [3]. According to reports from the National Institutes of Health (NIH), microbiological biofilm is responsible for about 65 % of microbiological infections and 80 % of chronic infections, respectively [4].
Although scientists, pharmacists, and doctors search for safe, broad-spectrum medications, resistant and biofilm-forming bacteria remain difficult to treat. Due to these issues, nanotechnology and nanoparticles as antibiotic alternatives seem appealing. Silver nanoparticles (AgNPs) are intriguing nanomaterials because of their distinctive properties and several nanomedicine applications. AgNPs release and quantity determine silver’s antibacterial properties. Non-ionized silver binds to tissue proteins and damages bacterial cell walls and nuclear membranes. Nanoparticles of silver denature and limit bacterial growth by binding DNA and RNA [5,6]. Consequently, AgNPs can be synthesized through different methods, including biological, physical, and chemical. Among these, biological synthesis is regarded as a sustainable and environmentally friendly approach, as it reduces the reliance on toxic chemicals [7]. Moreover, fungal AgNPs can be efficiently synthesized using species from the genus Fusarium, which are favored due to their filamentous morphology and ease of isolation from plant and soil sources. This well-characterized genus grows on simple media at moderate temperatures, making it ideal for nanoparticle biosynthesis [8].
Recent developments in nanoparticle functionalization have brought attention to the advantages of peptide-conjugated AgNPs, which extend beyond fungal-mediated production. Both nisin-loaded silver nanoparticles (N-AgNPs) and tryasine-conjugated AgNPs exhibited strong activity against both Gram-positive and Gram-negative bacteria, including Staphylococcus aureus, methicillin-resistant Staphylococcus aureus (MRSA), and Escherichia coli. This suggests that conjugation enhances targeting efficiency by increasing the local concentration of the active agent at the infection site [8-10].
 Building on these advances and enhancing the robustness of AgNPs against multidrug-resistant pathogens, incorporating safe antimicrobial agents such as the peptide nisin onto AgNPs represents a promising strategy to improve therapeutic efficacy. Nisin is a polycyclic peptide that functions as an antimicrobial agent and contains 34 amino acid residues with five lanthionine rings [11]. Nisin is a cationic bacteriocin that comes from Lactococcus lactis and is safe to use (GRAS). It was initially employed to keep food fresh, but its ability to kill Gram-positive microorganisms has made it useful in medicine as well [12,13]. The mechanism of the action of the antibacterial nisin is mainly due to the interaction with the negatively charged cytoplasmic membrane and the formation of pores on cell membranes, which results in the alteration of the permeability of the cells [14].To our knowledge, no studies have reported the use of Fusarium species to synthesize nisin-functionalized AgNPs. While Fusarium-derived AgNPs are known for their antimicrobial activity, their conjugation with peptides like nisin remains unexplored, representing a novel and promising research direction. This study aimed to biosynthesize, optimize, and characterize AgNPs. AgNPs using the filamentous fungus Fusarium solani. The resulting AgNPs were then conjugated with nisin to produce N-AgNPs. These nanomaterials were subsequently evaluated for their synergistic antibacterial efficacy and antibiofilm activity.

MATERIALS AND METHODS
Materials
Bacterial growth was done on Nutrient Broth (NB) and Luria-Bertani (LB) broth, while Mueller-Hinton Broth (MHB) was used for testing for antibacterial sensitivity. Fungi were cultivated in Potato Dextrose Broth (PDB). Media were sterilized by autoclaving at 121°C for 15 min and made up according to standard methods. The nisin was obtained from Smart Kimya (Izmir/ Turkey). All the reagents used were analytical grade.

Optimization of silver nanoparticles
The optimization of AgNPs production was achieved by methodically assessing four critical elements influencing synthesis yield. Their parameters were utilized to optimize nanoparticle creation and enhance production efficiency. This variable includes the AgNO3 salt concentration (0.5, 1, and 1.5 mM), the reduction time (24, 48, and 72 h), the pH value (5.0, 7.0, and 9.0), and the volume ratio of cell-free supernatant (CS-F) to AgNO3 (1:1, 1:2, and 2:1 V/V). Every experiment was carried out in triplicate to confirm the dependability and reproducibility.

Optimization of nisin-loaded biosynthesized AgNPs
To enhance stability and antibacterial efficacy, the nisin was loaded onto synthesized AgNPs by adjusting critical formulation conditions. Different concentrations (0.5, 1, and 3 mg/ml) were introduced to the biosynthesized AgNPs, after which the solution was subjected to a shaker incubator for an additional hour at 120 rpm to enhance homogeneous binding to the AgNPs.

Characterization of nanoparticles
Optical absorption spectra were obtained with a UV-line 9400 (SI Analytics) system. The samples were scanned from 300 to 600 nm to preliminarily characterize reduced AgNPs and N-AgNPs. Transmission electron microscopy (TEM) Ultrahigh-resolution TEM can visualize NPs at the nanoscale and their components. This method helps to explain membrane construction, development, and fouling [15]. The morphology of nano-structural materials is typically examined using scanning electron microscopy (SEM), as well as analysis of size, shape, purity, surface morphology, and dispersion for biosynthesized nanoparticles [16]. Under high vacuum mode, 4.83 mm working distance images were taken with numerous magnifications. The elemental compositions of NPs were investigated by energy dispersive X-ray spectroscopy (EDS) (1024 channels, 20 kV, 10 nA, 12,500× magnification). The crystallographic structure of materials is characterized using X-ray diffraction (XRD), the chemical composition and physical features of the material, and the NP thin film [17]. The average crystalline size of NPs was estimated using Debye-Scherrer’s equation based on the full width at half maximum (FWHM) value of the peaks [18]. Nanoscale surface analysis is safe with high-resolution atomic force microscopy (AFM). We produce air, liquid, or vacuum 3D topography. Sharp-tip, flexible-cantilever AFM probe. Interatomic cantilever deflections are measured by lasers. Movements offer nanometer-resolved 3D Surface Images [19]. the functional groups of materials were identified by analyzing certain peaks using the Fourier transform infrared spectroscopy (FTIR) technique. KBr-ground powdered samples were scanned for the spectrum between 400 to 4000 cm-1 and resolution to 4 cm-1 [20]. The Zeta potential assay was used for measuring the charge and stability of NPs. The particles above ±30 mV exhibit high electrostatic stability, while lower values may stabilize sterically [21]. The zeta potential of samples was assessed at 25°C using standard settings in a dispersion medium. 

Isolation and molecular identification of pathogenic bacteria
Isolation of pathogenic bacteria
Urine, ear, and wound swabs were used to isolate pathogenic bacteria, including Escherichia coli, Acinetobacter baumannii, and Staphylococcus aureus. Identification was performed using molecular techniques and the BD PhoenixTM technology. One single colony for each isolate was chosen and kept in glycerol stock at –20°C for additional study.

Identification of bacteria by BD Phoenix™
BD Phoenix is an automated microbiology system (BD Diagnostic Systems) used for the identification (ID) and antimicrobial susceptibility testing (AST) [22]. Results led isolates to be categorized as sensitive, resistant, or multidrug-resistant (MDR).

Molecular identification of pathogenic bacteria
The polymerase chain reaction (PCR) technique was employed for the molecular identification of isolated Bacteria, which is a technique used to amplify a specific sequence of DNA through denaturation, annealing, and extension steps. The bacterial total genome was extracted using a (for better BIO-TECH) following the manufacturer’s guidelines. The PCR reaction was carried out in a total volume of 20 μl, including 10 μl master mix (BIO-Rada), 0.5 μl of each forward and reverse primers, 1μl of DNA template, and 8μl of DDH2O. The 16S rRNA gene was targeted using universal primers (7F:5′-AGAGTTTGATYMTGGCTCAG-3′), (1510R:5′ACGGYTACCTTGTTACGACTT-3′) [23,24]. The PCR conditions were set to 30 cycles, with initial denaturation at 94°C for 5 minutes. Followed by denaturation at 94°C for 30 seconds, annealing at 58°C for 30 seconds, and extension at 72°C for 1 minute, with the final extension at 72°C for 5 minutes. Sanger DNA sequencing at Macrogen-Republic of Korea sequenced the ~1.5 kbp PCR product. Chromas software version 2.6.6 was used for quality and sequencing edition (Technelysium). 

Construction of the phylogenetic trees
Based on its 16S rRNA gene sequence, as previously reported, the isolate was taxonomically characterized by building a phylogenetic tree [25]. ClustalX 2.1 helped the sequence match closely related NCBI species [26]. Using MEGA 7, the phylogenetic tree was generated via the Neighbor-joining method, applying (Kimura 2-parameter model, with 1000 bootstrap repetitions. [27].

Antibacterial activity assay 
A disc diffusion assay was done according to [28]. with some modifications. The bacteria were activated in Müller-Hinton broth (MHB) at 37°C for 22–24 h. Then, 100 μl culture broth (0.5 McFarland) was uniformly distributed on Müller-Hinton agar plates. Different concentrations (4, 3, 2, and 1, 2, 3, and 4 mg/ml) of AgNPs and N-AgNPs were prepared, a sterile filter paper discs (5mm diameter) were loaded with 50 μl of each prepared distinct concentration. The plates were incubated at 37°C for 18–22 hours to allow interaction between the bacteria and the nanoparticles. Additionally, nisin alone(0.5mg/ml) was added to a sterile disc under the same conditions to evaluate individual antibacterial activity. The inhibition zones were daily measured in triplicate.
To calculate the minimum inhibitory concentration (MIC) for AgNPs and Nisin-AgNPs, Pathogenic bacterial isolates were adjusted to (108cells/mL), equivalent to 0.5 McFarland, in fresh overnight cultures. A 120 μl bacterial culture was suspended in 96-well sterile polystyrene microtiter plates. Following this, 80 μl of AgNPs and N-AgNPs at the appropriate concentrations (0.06, 0.125, 0.25, 0.75, 0.5, and 1 mg/ml) were introduced. A 120 μl of bacteria, 80 μl of media used as a positive control, AgNPs, N-AgNPs, and medium as a blank were added. incubated overnight at a shaker incubator, and a microplate reader assessed each well’s absorbance at 600 nm (BioTek (ELx800) / USA) [20].

Biofilm Inhibition Assay
Bacteria were cultured in LB medium for 24 h to inhibit biofilm formation by AgNPs and N-AgNPs. After adjusting to (108cells/mL), equivalent to 0.5 McFarland, each 96-well plate was filled with 120 μl of bacterial suspension and 80 μl of AgNPs and N-AgNPs at various concentrations (0.06, 0.125, 0.25, 0.5, 1mg/ml). After incubation for 24 h, all wells were discarded and dried for 15 min. Each well was then filled with 200 μl of deionized water, left for 2 min, discarded, and allowed to dry in the room for 30 min. Then,  each well was treated with 200 μl of 0.1% crystal violet and left at room temperature for 15–20 min. the stains were then removed, and each well was washed three times with deionized water, and air-dried for an hour. followed by adding 200 μl of absolute ethanol and incubated for 15 min.  A microplate reader is used to measure the absorbance at 595 nm [29]. 

Statistical analysis 
All statistical analyses were conducted with IBM SPSS Statistics 27.0.1. All results are presented as the mean ± standard deviation (SD) and were conducted in triplicate. Using the Duncan post hoc test and one-way ANOVA, the significance of differences between the control and treated groups was ascertained as follows. Significance was determined by a p-value of <0.05.

RESULTS AND DISCUSSION
Biosynthesis and optimization of silver nanoparticles
The extracellular synthesis of NPs and the mechanism are still in question; the supernatant and its content of reductase enzymes are mainly responsible for the biosynthesis of NPs. In this study, during biosynthesis of AgNPs, precursor salt (AgNO3) was reduced into AgNPs by F. solani supernatant. The preliminary characterization of AgNPs was done by their changing color after 1 h. of mixing the CF-S and AgNO3 by UV–vis spectroscopy. The reduction of Ag+ into Ag0 was indicated by the color intensity, which transitioned from mild yellow to brown and ultimately to dark brown after 72 hours of incubation. The broad and durable surface plasmon resonance peak (SPR) at 414 nm suggests that the reaction state has undergone a change. A previous study has examined the production of AgNPs using F. solani, and the peak at 415 nm is nearly identical [30]. AgNO3 concentration, pH, time, and ratio directly affect AgNP generation, as measured by UV-vis spectroscopy. The optimal conditions for synthesizing AgNPs using CS-F are pH 9, AgNO3 concentration 1.5 mM, time after 72 h, ratio 1:2, as shown in Fig. 1. The optimal pH for AgNPs synthesis can vary depending on the type of microbe used. In this study, pH 9.0 was found to be the most favorable, showing the highest absorbance and a sharp peak at 414 nm, and the color started to change within a few hours. The bioactive metabolites present in the fungal supernatant appear to be more stable and exhibit greater catalytic activity under alkaline conditions. In contrast, acidic pH levels led to aggregation of the AgNPs, while at pH 7.0 the synthesis was significantly reduced. The other study documented that the AgNPs were synthesized using Fusarium exosporium, which has the greatest production at pH 9.0 and pH 11 [31]. AgNO3 concentration plays another key role in nanoparticle synthesis. In this study, 1.5 mM AgNO3 was found effective for forming stable AgNPs. Higher concentrations can increase yield but may lead to particle aggregation, while lower levels may reduce efficiency. A similar result was reported by [32]. The authors showed that 1–2 mM AgNO₃ was the optimal concentration of salt for the formation of nanoparticles by fungal extracts. Time is a critical factor influencing the efficiency and yield of AgNPs synthesis. In this study, the reaction was monitored over a period of 24 to 72 h. A noticeable color change from pale yellow to dark brown indicated ongoing reduction of silver ions by fungal metabolites. UV-Vis spectroscopic analysis revealed that absorbance intensity gradually increased with time, reaching its peak at 72 h. suggesting continuous and efficient formation of nanoparticles. This implies that extending the reaction time allows for a more complete reduction of Ag⁺ to Ag⁰, leading to higher nanoparticle yield and stability [33]. These findings are consistent with previous reports, where prolonged incubation enhanced nanoparticle formation due to sustained activity of the reducing agents in the biological extract. A similar result was also reported in [34]. where maximum absorbance and nanoparticle formation were observed after 72 h. supporting the outcomes of this study.
The ratio between fungal supernatant and AgNO3 plays a key role in nanoparticle formation. An appropriate ratio ensures efficient reduction of Ag⁺ ions and stable nanoparticle formation. In this study, a 1:2 ratio (supernatant to AgNO₃) produced the best results, leading to strong color change and maximum absorbance. Too little supernatant may limit reduction, while too much may cause aggregation or unstable particles. The ratio of fungal supernatant to silver nitrate is crucial for efficient AgNP synthesis.  In this study, a 1:2 ratio gave optimal results. This ratio has also been suggested in previous studies. For instance, F. solani has been found to synthesize AgNPs stably with an optimum ratio of 1:1 or 1:2, and the resulting SPR peaks were about 414 - 420 nm, revealing the formation of NP efficiently [35,36].

Nisin loaded onto biosynthesized AgNPs 
The synthesis of N-AgNPs was performed using different concentrations of nisin, and the color change was monitored over time. Nisin was effectively loaded into AgNPs to create and improve N-AgNPs by using various amounts and incubation times. Nisin was mixed into AgNPs at a concentration of 0.5 mg/ml (as the optimum concentration) after 72 h. of production, and then the mixture was shaken for one more hour. Fig. 2 shows the formation of N-AgNPs. Loading nisin onto AgNPs resulted in a slight red shift of the UV-Vis absorption peak from 414 nm to 417 nm, indicating successful interaction or binding between nisin and the nanoparticle surface. This shift suggests a change in the SPR due to nisin coating. N-AgNPs also showed enhanced antimicrobial activity compared to AgNPs alone, due to the synergistic effect of silver and nisin. Similar findings were reported by [37]. where nisin-functionalized nanoparticles exhibited efficacy against Pseudomonas aeruginosa.

Characterization of nanoparticles 
The conversion of AgNO₃ and fungal cell-free supernatant into AgNPs was evidenced by UV-Vis spectroscopy, as indicated previously in (Fig. 1) and a noticeable color change from light yellow to dark brown. This visual change is often attributed to the SPR of AgNPs, as shown previously in (Fig. 2b), which has been extensively reported in biosynthesis studies [38,39]. reported that a color change is initial evidence of nanoparticle formation. Similarly [40]. reported the formation of brown color during AgNP formation by Fusarium oxysporum and the reduction of silver metal to Ag0 [41] highlighted this as a common confounding parameter in plant vs. micro pathogen-mediated NP synthesis. Such results corroborate the present observation and establish the color change as a reliable indicator of AgNP synthesis.
TEM results show that the AgNPs had mostly spherical to quasi-spherical shapes for AgNPs and quasi-spherical shapes for N-AgNPs. The particle sizes of AgNPs range from around 6.4 nm to 17 nm (Fig. 3A-a). The average particle size computed was 10.7 nm. Such shapes and nanoscale sizes match those of usual biosynthesis-produced AgNPs [42,43]. With a rather limited distribution, as shown in the size distribution histogram (Fig. 3A-b). The histogram of AgNPs shows high homogeneity and dispersion since most of the particles lie in the 6.4 –17.0 nm region. previous study [44] demonstrated that C. acidovorans yielded AgNPs with a smooth surface and spherical, oval, or irregular forms ranging from 6 to 53 nm.
 TEM illustration (Fig. 3B-a) of  N-AgNPs indicated a similar shape but with some higher particle size. Nisin loading expanded the distribution by surface adsorption and molecular interaction with AgNPs and raised the average particle size to 12.8 nm. As before reported, bio-conjugated nanoparticles with surface functionalization experience small size and heterogeneity changes [45,46]. whereas the sizes ranged from 7.9 nm to 18.1 nm (Fig. 3B-b). The histogram indicates a wider distribution than in simple AgNPs, which could be owing to the interaction between nisin and the nanoparticle surface, hence somewhat increasing the size [47].
SEM was used to investigate the morphology of NPs. SEM analysis was used to show that AgNPs and N-AgNPs were produced. An SEM study was done after the nanoparticle sample had been dried. A thin layer of the sample was made and examined. The microscopy images indicated that the nanoparticles were spread out evenly across the thin layer. However, a notable agglomeration of nanoparticles was observed, as shown in (Fig. 4A). Our approach produced a dense distribution of AgNPs and N-AgNPs with notable agglomeration using SEM examination. This observation is consistent with the results of [48]. who verified comparable nanoparticle characteristics using green banana peel extract. more monodispersed nanoparticles were observed [49]. which reduced the aggregation of the components in banana peel extract and therefore provided a lowered effective capping effect. Variations in aggregation degrees among studies could result from changes in synthesis conditions, including pH, temperature, and extract content, which influence the stabilization and dispersion of nanoparticles.
EDS was employed to analyze the chemical composition of the samples. EDS analysis of AgNPs indicated the presence of silver (Ag), shown by its characteristic peak at 3 keV, the peak position of the silver element. peaks at around 0.5 keV were also seen, suggesting the existence of oxygen and carbon, as reported in similar studies [50]. (Fig. 4B-a). Also, EDS analysis of N-AgNPs (Fig. 4B-b) approved the occurrence of silver and the organic elements incorporated with nitrogen and reveals the successful loading of nisin into the nanoparticles, the enhanced antibacterial action [51].
The crystalline structure of the synthesized NPs was analyzed using XRD . The XRD pattern exhibited characteristic diffraction peaks at 2θ values of 28.37°, 32.83°, 38.80°, 46.89°, 55.52°, 58.33°, and 77.70°. which can be index to plane (210), (122), (111), (200), (142), (241), and (311) planes of face-centered cubic (FCC) silver (Ag) (JCPDS card no. 04-0783) consistent with past studies on fungal-mediated synthesis [52-55]. As shown in Fig. 5A-a, this confirms the formation of crystalline AgNPs. The crystallite size is 9.23 nm, This value is calculated by employing the Debye-Scherrer equation to determine the (FWHM) of the peak that corresponds to 200 planes. Fig. 5A-b) shows the XRD pattern of the structure of N-AgNPs. Only three distinct peaks were shown at 2θ values 25.810, 30.450, and 50.780, with a crystalline size of the particles increased to 14.29 nm upon loading with nisin, suggesting successful surface modification. This increase is following prior research, which demonstrated that the conjugation of peptides to AgNPs led to a quantifiable increase in particle size as a consequence of surface coating [56]. and a similar result was reported by [57]. showed that silver-nisin nanoparticles maintained their typical face-centered cubic silver crystalline structure, as seen by distinct XRD peaks at 2θ values of 38°, 44°, 64°, and 77°. This proves that nisin loading was effective without losing crystallinity.
 AgNPs and N-AgNPs were confirmed in surface topography and shape using AFM imaging. It was ascertained from the three-dimensional horizontal cross-section of both samples that the surface topography of the produced AgNPs and N-AgNPs were somewhat round in form (Fig. 5B.a-b). The N-AgNPs’ surface seemed to be rather more textured, likely due to the presence of nisin on the surface of the nanoparticles [58] The functionalizing procedure involved in their synthesis [59].
FTIR spectroscopy is an effective technique to determine the bioactive groups of nanomaterials. In this study the FTIR spectra (Fig. 6A) shows the characteristic peaks that verify the existence of functional groups on the surface of AgNPs and N-AgNPs and free nisin. The O-H and N-H stretching vibrations were observed as a broad band around 3435–3422 cm⁻¹ and were indicative of hydroxyl groups and amines .The bands at 2957-2924 cm⁻¹ were assigned to C–H stretch, and are related to an aliphatic chain [60]. The proteins or peptides have an intense peak at approximately 1632 cm⁻¹ corresponding to C=O stretching (amide I band). The peaks at 1465cm−1 belong to C-N stretching so indicating the presence of proteins or peptides involved in nanoparticle stabilization, respectively like what was found in consistent with past fungal-mediated synthesis studies [61,62]. , respectively, whereas those at 1112-1114 cm⁻¹ are due to C-O stretching. The Ag-O bond vibrations for peaks under 700 cm⁻¹ that form for the Ag nanoparticles can also be seen. As a whole, the FTIR spectra of nisin, AgNPs, and nisin-stabilized AgNPs were similar. (Fig. 6A) illustrated that nisin was successfully loaded onto AgNPs. Comparatively the FTIR spectra show the common peaks of relevant to AgNPs , N-AgNPs , and free nisin , especially those for O-H/N-H stretching 3430 cm⁻¹ , C-H stretching 2957–2924 cm⁻¹ , and the amid I band 1632 cm⁻¹indicating the coexistence of biomolecules on the nanoparticle surface. Peaks attributed to C-O stretching at 1112–1114 cm⁻¹ and Ag–O vibrations below 700 cm⁻¹ were also observed . these spectral feature, specially amid I and O-H/N-H bands are commonly reported in previous study in biosynthesized AgNPs and reflected by capping peptide and protein . similarities between the spectra of nisin and nanoparticles indicated the effective stabilization and interaction of AgNPs by nisin. 
The zeta potential value confirms that AgNP is moderately stable even further. The AgNPs were negatively charged in the dispersed media, hence, the resulting value of -3.7 mV (Fig. 6B-a). The results showed that the particles repelled one another, thereby preventing aggregation. While nisin was loaded onto AgNPs, enhanced stability was observed. while free nisin was added dropwise at a low concentration to the NPs solution, more nanoparticles with the same charges were wrapped by negatively charged molecules as confirmed by the zeta potential of –38.4 mV this number shows a stable colloidal solution of N-AgNPs. the value reveals that the generated nanoparticles have well-dispersed electrostatic stability in suspension (Fig. 6B-b). This improvement corresponds with those [63]. who attributed the higher stability to the capping effect of nisin’s amine and amide groups and found that nisin-capped AgNPs had zeta potential values of –23.2 mV and –27.0 mV. Similarly, [64]. found that nisin added to AgNPs raised the zeta potential values of the bioconjugates, enhancing stability and antibacterial action. 

Molecular identification of bacteria 
This study identified bacterial isolates using BD Phoenix and 16S rRNA gene sequencing. The automated phenotypic identification system BD Phoenix identified the clinical isolates as E. coli, A. baumannii, and S. aureus, common hospital organisms. The PCR products were sequenced, and the sequences were aligned to sequences representing known bacteria available on the National Center for Biotechnology Information (NCBI) database. The bacterial isolates were identified as E. coli, A. baumannii, and   S. aureus, and the DNA sequence of the 16s rRNA gene of each of the isolates was submitted to the BANKIT, and gene bank accession numbers were assigned to all of the 16s rRNA sequences. Accession numbers were assigned for each sequence as follows: E. coli strain AD2 (PV596248.1), A. baumannii strain AD3 (PV596249.1), and S. aureus strain AD1 (PV596247.1). The phylogenetic tree of isolates was constructed based on the 16S rRNA gene sequence. (Fig. 7) A newly isolated of E. coli strain AD2 formed a monophyletic group with other E. coli strains such as strain T18-6-1 (accession numbers: PQ097308.1), which was isolated from Japan, and strain DSM 112117 (accession numbers: OM658577.1) isolated from Germany, which shows a common ancestral lineage. Similarly, a new A. baumannii isolated (AD3) clustered withing strongly supported monophyletic clade with strain C9 (accession numbers: MK070057.1) isolated from Washington DC, strain M.pstv.34.1(accession numbers: KM108526.1) isolated from France, and strain GPUL16(accession numbers: MF398381.1) isolated from India. In contrast, the newly isolated S. aureus strain (AD1) formed only a monophyletic group with strain AHL21(accession numbers: MN067694.1), which was previously isolated from Iraq.

Antibacterial activity assay
Disc diffusion was used to assess the antibacterial activity of AgNPs and N-AgNPs against clinically isolated bacterial strains such as E. coli, A. baumannii, and  S.  aureus. We depicted that N-AgNPs have a larger zone of inhibition compared to AgNPs alone. At 4 mg/ml concentration, AgNPs produced an 11 mm inhibition zone for Gram-negative bacteria, including E. coli , but this zone expanded to 14 mm when N-AgNPs were used, which demonstrated a 3 mm improvement. A. baumannii. AgNPs produced a 9.66 mm inhibitory zone, N-AgNPs raised this value to 12.33 mm, so improving by 2.67 mm. Regarding Gram-positive bacteria like S. aureus, the inhibitory zone grew from 8 mm with AgNPs to 9.66 mm with N-AgNPs, increasing the difference by 1.66 mm, as shown in (Table 1). The inhibition zone measured for nisin alone was 6.1mm for E. coli, 6.6mm for A. Baumannii and, 7 mm for S. Aureus. Indicates a synergistic effect when combined with AgNPs.  A naturally occurring antimicrobial peptide, nisin may increase the bactericidal effect of AgNPs when combined. AgNPs and antimicrobials boost antibacterial efficacy. according to other studies, AgNPs alone did not raise P. aeruginosa inhibition zones [65,66] discovered that they have a greater effect when nisin is loaded onto AgNPs against the studied bacteria. Nisin-conjugated zinc oxide nanoparticles have enhanced antibacterial activity against E. coli and S. aureus [13]. This enhanced impact results from nisin’s binding to lipid II, a necessary component of bacterial cell wall formation, and disturbance of membrane integrity by creating holes, therefore promoting the increased antibacterial activity of AgNPs [67]. Published similar results showing enhanced activity of biogenic AgNPs mixed with nisin; Elsherif and his friends [68]. revealed that nisin nanoparticles had higher effects against S. aureus and E. coli O157:H7. 
Table 2 confirms that N-AgNPs reveal greater antibacterial activity than AgNPs alone. As a lower concentration is required to inhibit the bacterial growth for Gram-negative bacteria, Results showed that E. coli showed a two-fold reduction in MIC test (0.25mg/ml for N-AgNPs vs. 0.50 mg/ml for AgNPs ), At the same time, bioconjugation of AgNPs with nisin resulted in a significant reduction of MIC values. respectively, for Gram-positive bacteria S. aureus. These findings are consistent with what other research have shown. For example,  [69] found that silver-nisin conjugates had lower MICs (4 µg/ml for S. aureus and 2 µg/ml for E. coli). but A. baumannii, showed no difference in MIC (at 0.5 mg/ml for AgNPs and N-AgNPs) (Table 2). regardless of how nisin is conjugated. Some research [70]. on the other hand, say that N-AgNP conjugates work better against  A. baumannii, with MICs in the 52–125 µg/ml range. N-AgNPs showed considerable reductions in MIC values for E. coli and S. aureus after bioconjugation with nisin (p < 0.05), suggesting improved antibacterial action compared to AgNPs alone. Nisin conjugation did not improve antibacterial activity against A. baumannii.

Antibiofilm Inhibition Assay
Various concentrations (1, 0.5, 0.25, 0.125, and 0.06 mg/ml) of biosynthesized AgNPs and N-AgNPs were utilized to examine their effectiveness in preventing biofilm formation in clinically isolated bacteria this study, AgNPs and N-AgNPs greatly inhibited the formation of biofilm. The synergistic activity of nisin and AgNPs is responsible for the increased effectiveness of N-AgNPs, especially at lower doses. AgNPs reduced biofilm at 0.5 mg/ml in E. coli, but N-AgNPs were completely inhibited at 0.25 mg/ml, thus demonstrating the capacity of nisin to increase nanoparticle penetration Similar findings were reported by [71]. who observed that nisin-functionalized nanoparticles exhibited greater antimicrobial effects against Gram-negative pathogens due to improved delivery and stability. Likewise, at 0.5 mg/ml  AgNPs and N-AgNPs, reduced biofilm in A. baumannii, suggesting a cooperative action against this pathogen [72]. Notably, N-AgNPs were strongly efficacious against S. aureus, with complete biofilm removal noted at 0.125 mg/ml. These results reinforce the greater susceptibility of Gram-positive bacteria to nisin–based treatments, possibly because of its thick peptidoglycan layer, which could be efficiently targeted by the pore–formation capability of nisin [73]. In contrast, the same AgNPs alone at higher concentrations result in only partial inhibition. A similar phenomenon was reported by [74]. which observed that the bioactivity of NPs after being modified with antimicrobial agents is also improved, leading to a greater substance efficacy of anti-biofilm activity against S. aureus. The difference between the two statements was statistically significant (P<0.05) at all concentrations. The results suggest that loading nisin onto AgNPs increases their biofilm-disrupting potential against clinically isolated bacteria (Fig. 8). 

CONCLUSION
This study successfully highlights the bio-synthesized AgNPs using fungal supernatant and nisin loaded onto AgNPs. Characterization techniques include UV-vis spectroscopy, TEM, SEM, EDS, XRD, AFM, FTIR, and Zeta potential, verified structural integrity, production, and effective conjugation of nisin onto AgNPs, with improved colloidal stability and greater particle sizes. The results of antibacterial and antibiofilm experiments demonstrated that N-AgNPs exhibited significantly enhanced antimicrobial and antibiofilm activity against clinically isolated Gram-negative and Gram-positive bacteria. compare to AgNPs alone. These findings highlight the efficacy of nisin-functionalized AgNPs as a viable antibacterial and antibiofilm agent against clinically isolated pathogenic bacteria.

ACKNOWLEDGMENTS
The authors would like to thank the research center of Charmo University for providing laboratory equipment and support throughout this study, special thanks extended to professor Dr. Haider Mousa Hamzah for valuable assistance and guidance during this research, and miss Payman M. Mohamed salih for her helpful support during the laboratory work. Also We also thank the head of the Medical Laboratory Science Department for continuous support.

CONFLICT OF INTEREST 
The authors declare that there is no conflict of interest regarding the publication of this manuscript.