Instrumentation.

Instrumentation for the structural, morphologicaland elemental composition of the Chromolaena odorata leaf biochar (COLB) and the nanocomposites was done using standard analytical procedures. Genesis 10S UV-Vis spectrophotometer was used to determine the UV-Vis spectra. The Bruker^® D8 Discover x-ray diffractometer, equipped with a Lynx Eye detector, under Cu-Ka radiation (λ = 1.5406 Å at 40 KV and at 30 mA) was used for the determination of the crystalline nature of the COLB and nanocomposite and diffraction recorded with 2θ = 10 to 70. Phenom Prox by Phenom World, Eindhoven, Netherlands, was applied for the determination of the structural morphology of the COLB and nanocomposite. The Cary 630 Agilent Technologies, USA, Fourier transform infrared spectrophotometer (Shimadzu, Japan) in the range of 4000–400 cm^-1 in KBr using nujol mull as the mulling agent was used for functional group determination. The TEM analysis for determining the average crystallite size was done using a JEM ARM 200F. Physicochemical analysis involving temperature, salinity, pH, total hardness, dissolved oxygen, electrical conductivity was effected using Smart 6-in-1 pH multimeter. All drying and equilibrations were effected using oven (model DHG) and rotary shaker (RF-12 Remi equipment) respectively. The Erlenmeyer flask, Autoclave (Alpha Laboratory, Ltd) model NL280-A, Pressure Cooker (Crown Star Atsago, India Instruments Pvt Ltd., India and water bath (helmreasinn multipurpose) model DK420, incubator, and microscope were used for antimicrobial experiments. All microbiological weighing was executed using an analytical balance, Mettler Toledo PL, 303.

Reagents.

Analytical grade reagents were sourced from Merck Germany, Biolbs M0270, England, Zymo Research, Promega Madison, WI, USA, BioRad Universal Hood II Philadelphia, PA, USA, NZYtech Lisbon, Portugal, and K.J. Pharmaceutical and chemical and used without further modification: The following reagents were used for the synthesis of the nanocomposite and for well water tests. Iron(III) trioxonitrate(V), Silver trioxonitrate(V), hydrochloric acid, potassium heptaoxodichromate(VI), ethanol, tetramethyl-p-phenylenediamine, crystal violet, lugols Iodine, acetone, safranin, hydrogen peroxide, dimethylsulphoxide, ethanol, barium chloride, concentrated sulphuric acid, alpha-naphthol (Barrit’s solution A), phenol disulphonic acid, tetraoxosulphate(VI) acid, barium chloride, Potassium hydroxide, (Barrit’s solution B), Zr Fungal/Bacterial DNA Miniprep, Agarose powder, Nuclease free water, Taq 2× Master Mix, UV transilluminator, molecular weight ladder, Eosin methylene blue agar, Salmonella and Shigella agar, MacConkey agar, Bushnel-Hans agar, Sabraud Dextrose agar and Simmon citrate agar.

Chromolaena odorata leaf biochar (COLB) was prepared from the dried leaves of Chromolaena odorata by carbonization. Consequently, 600 g of dried leaves were loaded into a carbonizer and the temperature was set at 400°C for 30 min. The formed COLB was collected, ground into fine powder, and kept in an airtight container.

The synthesis of the nanocomposites involves the addition of 0.3 g each of silver trioxonitrate(V) or iron(III) trioxonitrate(V) for silver oxide and iron(III) oxide Chromolaena odorata leaf biochar nanocomposite respectively to two separate conical flasks. Then, 10 g of COLB and 100 cm³ of Chromolaena odorata leaf extract were added separately to a 1000 cm³ distilled water in each flask. The mixture was heated with continuous stirring for 30 min and then cooled and centrifuged at 4000 rpm. The formed nanocomposites (iron(III) oxide Chromolaena odorata leaf biochar nanocomposite (Fe₂O₃-COLBN) and silver oxide Chromolaena odorata leaf biochar nanocomposite (Ag₂O-COLBN)) were collected, washed with ethanol and distilled water to pH 7, dried at a temperature of 110°C for 2 h, and then kept for further use in an airtight container.

Standard analytical protocols were used to sample well water samples early in the morning (between 7 and 10 a.m.) from different locations (Figure 1) using plastic bottlesprewashed with dilute trioxonitrate(V) acid. Two containers were used for each sampling point, with one container treated with two drops of acid to prevent oxidation before heavy metal determination [18]. The well was allowed to run for at least ten min before assaying the samples. Physicochemical parameters were taken in situ at the sampling site and in the laboratory using standard analytical instruments. The water samples were then transported to the chemistry and microbiology laboratory in sterile bottles for further analysis (heavy metal, physicochemical, and antimicrobial tests).

In the treatment of the well water samples, 200 cm³ of each sample was measured and added to a sterile, labeled, separate container foreach nanocomposite. Then 0.5 g of Fe₂O₃-COLBN or Ag₂O-COLBN was added, and the mixture was equilibrated for 30 min at 27°C. The mixture was filtered as described elsewhere [18], and the filtrate was kept for further analysis.

The nanocomposites were desorbed with HCl, washed with distilled water, dried, and reused three times to evaluate the stability.

The concentrations of heavy metals in the well water before and after treatment with the nanocomposites were determined using the atomic absorption spectrophotometric method [19]. The concentrations of the anions (Cl^-, SO₄^2- and NO₃^-) present in the water samples before and after treatment with nanocomposites were also determined using standard procedures as described elsewhere [20].

The agar well diffusion method as described elsewhere [21] was used to determine the antimicrobial efficacy of the COLB, Fe₂O₃-COLBN, and Ag₂O-COLBN. The test Organisms; Staphylococcus aureus, Escherichia coli, Klebsiella, Salmonella spp., Candida albicans, Aspergillus and proteus were tested for. The isolates were accessed from Alex Ekwueme Federal Teaching Hospital, Abakaliki, Nigeria, and then carefully transported in nutrient broth to the Ultra Modern Laboratory of the Department of Applied Microbiology, Ebonyi State University, Abakaliki. The antimicrobial inhibitory test was performed on different concentrations (0.001, 0.01 and 0.1 g) of the COLB, Fe₂O₃-COLBN, and Ag₂O-COLBN against the test organisms by dissolving the COLB, Fe₂O₃-COLBN, and Ag₂O-COLBN in dimethylsulfoxide [22]. During the test, the cultures of the test organisms were made in nutrient broth, incubated for 24 h, and adjusted to the McFarland turbidity standard. The blank control (gentamycin for bacteria and amphotericin β for fungi) were prepared and aseptically deposited on the Mueller-Hinton agar plates. Then, the cultures of the test organisms and controls were incubated at 37°C for 18-24 h, and the inhibition zone diameters were measured according to standard protocols.

Different media such as Cetrimide agar, Salmonella/Shigella agar, Eosin Methylene Blue agar, Macconkey agar, Nutrient agar, Sabraud Dextrose agar and Bushnel-Hans agar were made for the antimicrobial evaluation of the water samples and were prepared according to the manufacturers instruction. Sterile containers of 500 cm³ was used to transport the water samples at a temperature of 8°C to Ultra-Modern Laboratory of Ebonyi State University, Abakaliki, Nigeria and all were processed within 6 h after collection. All the tests were executed as described elsewhere without modification [6].

The isolation and identification of bacterial strains (Gram staining, cell arrangement, indole test, oxidase test, catalase test, coagulase test, citrate test, and methyl red test) were done using standard procedures as described elsewhere [23,24]. The coliform count per unit (CFU/mL) was determined using Equation 1.

The influence of nanocomposite dosage, pH, and contact time on the percentage removal of heavy metals from borehole water is presented in Figure 10 The removal efficiency increased progressively with increasing adsorbent dosage up to 0.5 g. Beyond this optimum dosage, only marginal improvements were observed, suggesting possible active-site saturation and/or particle aggregation that limit further adsorption efficiency [7].

The effect of pH (Figure 10) showed a gradual increase in removal efficiency from acidic to near-neutral conditions, with maximum performance observed at neutral to slightly alkaline pH. This behavior indicates favorable surface charge interactions under these conditions and reduced competition between hydrogen ions (H⁺) and metal cations for adsorption sites [30].

Similarly, contact time significantly influenced adsorption performance. The removal efficiency increased rapidly with increasing contact time, reaching equilibrium at approximately 30 minutes. Beyond this point, a slight decline was observed, indicating possible desorption or redistribution of adsorbed species on the nanocomposite surface.

The adsorption capacity (q_e) of the nanocomposites for heavy metal removal was determined using the mass balance equation:

where q_e represents the adsorption capacity (mg/g), C_i is the initial metal concentration (mg/L), C_f is the equilibrium concentration after treatment (mg/L), V is the volume of the solution (L), and M is the mass of the adsorbent (g) [6].

In this study, 0.5 g of Fe₂O₃-COLBN or Ag₂O-COLBN nanocomposite was introduced into 200 cm³ (0.2 L) of borehole water and equilibrated for 30 minutes at 27°C. Following filtration, the residual metal concentrations were measured and used to compute q_e, thereby quantifying the adsorption performance under the defined experimental conditions.

The adsorption system exhibited a typical dosage-dependent behavior, where increasing adsorbent mass enhanced removal efficiency due to the greater availability of active surface sites. However, adsorption capacity (mg/g) decreased with increasing dosage, reflecting reduced adsorbate loading per unit mass of adsorbent. The maximum adsorption capacity of 650 mg g^-1 observed at 0.1 g highlights highly efficient adsorbate-adsorbent interactions at low dosage, consistent with adsorption behavior commonly reported for nanostructured materials [30].

The reusability of the nanocomposites was evaluated over three consecutive adsorption-desorption cycles to assess operational stability and regeneration efficiency. After each cycle, the spent adsorbent (0.5 g) was regenerated using 0.01 M HCl, thoroughly washed with deionized water until neutral pH was attained, dried, and reused under identical experimental conditions to treat200 cm³ of borehole water.

As shown in Figure 11, the removal efficiency declined slightly across successive cycles. The efficiencies recorded were 95.0 ± 0.8%, 94.0 ± 1.0%, and 93.0 ± 1.2% for the first, second, and third cycles, respectively. Despite this marginal decrease, the nanocomposites retained approximately 97% of their initial performance after three cycles, corresponding to a total efficiency loss of about 3%. Importantly, no statistically significant variation in decontamination efficiency across cycles was observed, indicating consistent adsorption performance. The narrow error margins further confirm the reproducibility and structural stability of the nanocomposites during repeated use [6]. Such stability is essential for practical water treatment applications, where adsorbent durability directly influences process economics and sustainability [7].

The observed high reusability is attributed to the structural integrity of the nanocomposite matrix and the reversible nature of adsorption processes. Acid regeneration with dilute HCl protonates surface functional groups (-OH, -COOH, and -NH₂), facilitating desorption of bound metal ions without disrupting the adsorbent framework.

The minimal performance loss suggests strong immobilization of nanoparticles within the biochar or supporting matrix, thereby reducing nanoparticle leaching and preventing agglomeration during repeated cycles [30]. Furthermore, the persistence of high removal efficiency indicates that key adsorption mechanisms—surface complexation, ion exchange, and electrostatic attraction remain largely unaffected by regeneration [30].

The slight decline in performance after multiple cycles may be attributed to partial pore blockage, minor loss of active functional groups, or incomplete desorption of strongly bound metal species.

From Table 1, all nine water samples analyzed showed significant microbial contamination before treatment, with total colony counts ranging from 1.60 × 10⁴ to 1.90 × 10⁵ CFU/mL. The presence of bacteria such as Escherichia coli, Staphylococcus spp., Shigella spp., Klebsiella spp., Proteus spp., and Pseudomonas spp. indicates fecal contamination, poor sanitation, and high risk of waterborne diseases [31,32].

Treatment of the well water samples with Fe₂O₃-COLBN showed complete inhibition of microbial growth in seven of nine samples. Only two samples (Nwiboko compound and NO. 1, Orifite street) recorded residual bacterial counts of 2.0 × 10² and 2.4 × 10² CFU/mL, respectively, indicating more than 99% reduction in microbial load. It is believed that increased equilibration to 60 to 120 min could lead to complete removal of the microbial load. The generation of reactive oxygen species (ROS) by Fe₂O₃ nanoparticles, which disrupt microbial membranes and the adsorptive capacity of biochar, which helps trap and deactivate bacteria could be responsible for the antimicrobial activities [32].

The Ag₂O-COLBN was more effective in bacterial decimation, achieving 100% microbial inhibition in all samples except two (Agbaja Azumili and Kpirikpiri market), where microbial counts dropped to 1.4 × 10² and 1.2 × 10² CFU/mL, respectively. The effectiveness could be attributed to the broad-spectrum antimicrobial activity of silver oxide, which causes disruption of bacterial DNA replication and protein synthesis, penetration and destruction of microbial cell walls and membranes, and release of Ag⁺ ions, which are toxic to bacterial cells even at low concentrations [33].

To further establish the nature of the microbes in the water samples, the antimicrobial treatment of well water using Fe₂O₃-COLBN and Ag₂O-COLBN was evaluated based on the presence and biochemical characteristics of bacterial isolates from different well water samples as shown in Table 2.

The untreated water samples from six different wells revealed the presence of various pathogenic bacterial species, identified through biochemical tests such as Gram staining, indole, oxidase, catalase, citrate, methyl red, and coagulase tests, and their characteristics are shown in Table 3.

These isolates are known to be opportunistic or enteric pathogens, indicating fecal contamination and poor water hygiene. Their presence in well water poses significant public health risks, particularly Klebsiella, Shigella, and Proteus, which are commonly implicated in urinary tract infections, diarrhea, and other gastrointestinal disorders [32]. The microbial response based on treatment is shown in Table 4 and illustrates the plausible mechanism underlying microbial deactivation and denaturation.

The Fourier Transform Infrared (FTIR) spectroscopy analysis of Chromolaena odorata biochar (COB), iron(III) oxide-Chromolaena odorata leaf biochar nanocomposite (Fe₂O₃-COLBN), and silver-Chromolaena odorata leaf biochar nanocomposite (Ag-COLBN) is shown in Figure 12. It gives the functional groups and molecular interactions operating in the structures of the biochar and nanocomposites. The observed bands and band shifts along with their respective assignments reveal the structural modifications that occur upon metal oxide incorporation into the biochar matrix.

The presence of broad bands around 3205.5 cm^-1 (COB), 3224.1 cm^-1 (Fe₂O₃-COLBN), and 3380.2 cm^-1 (Ag-COLBN) indicates O-H stretching vibrations, attributed to hydroxyl groups from adsorbed water, alcohols, or phenolic compounds. The C≡C stretching vibration at 2129.9–2120.9 cm^-1 is common across all samples, indicating the presence of alkyne groups, possibly from lignocellulosic precursors of the biomass [22].

The C=O stretching bands shift from 1878.6 cm^-1 (COB) to 1990.4 cm^-1 in Fe₂O₃-COLBN and 2009.0 cm^-1 in Ag-COLBN. This shift suggests interaction between the carbonyl groups and the embedded metal oxides, which may alter electron density and bonding environment [34]. The band at 1408.9 cm^-1 in COB and Fe₂O₃-COLBN, and 1420.1 cm^-1 in Ag-COLBN corresponds to C-H bending vibrations, indicating minor structural rearrangements due to silver nanoparticle loading.

A notable change was observed in the C-O stretching region, which shifts from 1244.9 cm^-1 in COB to 1021.3 cm^-1 in both nanocomposites, suggesting that C-O bonds in alcohols, phenols, or ethers were modified during metal nanoparticle intercalation, possibly due to surface oxidation or complexation [35]. The bands at 872.6–872.2 cm^-1 across all samples correspond to C-H bending, confirming aromatic character in the biochar matrix. The Fe-O stretching vibration at 766.6 cm^-1 in Ag-COLBN and 693.3 cm^-1 in Fe₂O₃-COLBN confirms the successful incorporation of Ag₂O and Fe₂O₃ respectively into the biochar matrix [34].

Scanning Electron Microscopy (SEM) of COLB (C), Silver Oxide-Chromolaena odorata Leaf Biochar Nanocomposite (A), and iron(III) Oxide-Chromolaena odorata Leaf Biochar Nanocomposite (B), as shown in Figure 13, provides detailed information about the surface morphology and texture of the studied materials and is crucial in evaluating how modifications or dopants affect matrix structures. The COB exhibits a relatively porous and less compact surface structure, which reflects the carbonaceous framework derived from the pyrolyzed plant material. This base structure is crucial as it provides the physical and chemical scaffold for nanocomposite formation and provides pore architecture for adsorption capacity but lacks the additional functionalities provided by metal oxide nanoparticles [35].

The SEM micrograph of Ag-COLBN likely reveals a rough and irregular surface with granular deposits, which are indicative of silver oxide nanoparticles well dispersed on the biochar matrix. These particles often appear as bright spots due to their higher electron density compared to the carbon matrix. The presence of Ag₂O nanoparticles enhances surface roughness and may increase the number of reactive sites, thereby improving photocatalytic and antimicrobial properties [36].

The SEM image of Fe-COLBN typically shows aggregated iron oxide particles embedded or anchored onto the porous biochar surface. The surface appears denser and less porous than Ag-COLB, possibly due to the sintering or agglomeration behavior of iron oxide nanoparticles. The iron particles contribute to magnetic and redox activity that are important in catalytic and adsorption applications [35]. The EDX elemental composition of the biochar and nanocomposites is presented in Table 5, Table 6, and Table 7. The presence of Fe and Ag in the nanocomposites of Table 6 and Table 7 respectively indicates that the actual incorporation of the metals into the biochar matrices.

X-ray diffraction (XRD) analysis is vital for determining the crystallinity and phase composition of materials. The XRD patterns of the COLB (C), Fe₂O₃-COLBN (A) and Ag₂O-COLBN (B) are shown in Figure 14. The XRD pattern of COLB shows broad peaks centered around 2θ = 20-30°, indicating the amorphous or poorly crystalline structure of the biochar. This is characteristic of carbonaceous materials, with diffraction peaks attributed to the (002) plane of graphitic carbon [8,13]. The broadness of these peaks confirms a disordered carbon structure, as expected given the heterogeneous composition of plant-derived biochar.

The incorporation of silver oxide nanoparticles onto COLB results in distinct diffraction peaks at 2θ values around 32.3°, 38.1°, 44.3°, 64.4°, and 77.4°, corresponding to the (111), (200), (220), (311), and (222) planes of Ag₂O and metallic Ag (JCPDS Card No. 04-0783 and 65-2871). The sharp peaks confirm the crystalline nature of the silver species and their successful deposition onto the biochar matrix [35].

The XRD pattern of the Fe₂O₃-COLB nanocomposite displays prominent peaks at 2θ ≈ 24.1°, 33.2°, 35.6°, 40.8°, 49.5°, 54.1°, and 62.5°, which are indexed to the (012), (104), (110), (113), (024), (116), and (214) planes of hematite α-Fe₂O₃ (JCPDS Card No. 33-0664). These sharp, well-defined peaks are characteristic of highly crystalline hematite nanoparticles embedded within the biochar matrix [10].

The UV-Vis absorption spectra of COLB (C), Fe₂O₃-COLBN (A) and Ag₂O-COLBN (B) is shown in Figure 15 and provide defined insight into the optical properties and surface plasmon resonance (SPR) characteristics of synthesized nanocomposites. There is no absorption peak in the UV-Vis spectrum of COLB, which could be attributed to its highly carbonized nature, a lack of chromophoric groups, or metal-based electronic transitions, and is in line with other studies [34]. From Figure 15, the absorption peak at 757.27 nm is characteristic of Ag₂O nanomaterials and suggests that the Ag₂O-Chromolaena odorata biochar nanocomposite has strong optical activity in the visible to NIR range [35].

The absorption peak at 271.00 nm suggests the presence of Fe-O charge transfer transitions, commonly observed in iron-based nanocomposites [12]. This peak indicates that incorporating iron(III) oxide into Chromolaena odorata biochar enhances its optical response.

Transmission Electron Microscopy (TEM) micrographs of COLB (C), Fe₂O₃-COLBN (A) and Ag₂O-COLBN (B) are shown in Figure 16 and give detailed information on the morphology, size, and dispersion of nanoparticles in a composite. The histogram showing the particle size distribution of the biochar and nanocomposites is shown in Figure 17.

The TEM image of COLB reveals an amorphous and porous carbonaceous structurecharacteristic of biomass-derived biochar. The porous nature indicates a high surface area suitable for adsorption and nanocomposite loading [8]. The absence of distinct crystalline phases suggests that the biochar is mainly carbon-based and has low crystallinity. The TEM micrograph of the Ag-COLBN nanocomposite (A) shows well-dispersed, spherical-to-quasi-spherical silver oxide nanoparticles distributed on the biochar surface. These nanoparticles appear as darker-contrast regions due to their higher electron density compared to carbon. The particle size appears to range from approximately 10-30 nm, which aligns with typical dimensions reported in similar studies [8]. This uniform dispersion indicates effective incorporation of silver oxide onto the biochar matrix. In the case of Fe-COLBN (B), the TEM micrograph displays clusters of iron(III) oxide nanoparticles embedded within the biochar matrix. These nanoparticles are typically smaller (5-20 nm) and exhibit a more aggregated morphology than their silver counterpart. This aggregation could be due to the magnetic nature of iron oxide nanoparticles, which promotes particle-particle interactions [35].

The antimicrobial evaluation inhibitory effects of Chromolaena odorata leaf biochar (COLB), iron(III) oxide Chromolaena odorata leaf biochar nanocomposite (Fe-COLBN), and silver Chromolaena odorata leaf biochar nanocomposite (Ag-COLBN) against seven microorganisms: Staphylococcus aureus, Escherichia coli, Klebsiella, Salmonella spp., Candida albicans, Aspergillus spp., and Proteus are presented in Figure 18 and Plate 1. The COLB and Fe-COLBN both recorded a 12 mm zone of inhibition, while Ag-COLBN showed slightly lower activity (10 mm). This suggests that Fe-COLBN retains strong bactericidalactivity against Gram-positive bacteria, potentially due to oxidative stress induction and membrane disruption [23]. The Ag-COLBN demonstrated the highest inhibition (14 mm), followed by Fe-COLBN (12 mm) and COLB (10 mm). This reflects silver nanoparticles’ well-documented efficacy against Gram-negative bacteria through silver ion release and reactive oxygen species (ROS) generation [23]. The Fe-COLBN and Ag-COLBN both exhibited 14 mm zones, surpassing COLB (12 mm). The equal performance of the two nanocomposites may be attributed to synergistic metal-biochar effects that enhance antimicrobial penetration and sustained ion release [22]. The Fe-COLBN achieved the highest inhibition (14 mm), with Ag-COLBN at 12 mm and COLB at 10 mm. The superior Fe-COLBN performance here could result from iron oxide nanoparticles’ ability to generate hydroxyl radicals through Fenton-like reactions [21]. All three materials (COLB, Fe-COLBN, and Ag-COLBN) showed similar inhibition (10 mm), suggesting moderate antifungal potential. This may be due to the inherent resistance of yeasts compared to bacteria [23]. The COLB showed the highest inhibition (10 mm), followed by Ag-COLBN (9 mm) and Fe-COLBN (8 mm). This indicates that metal nanoparticle incorporation did not significantly enhance antifungal activity against filamentous fungi in this case. All treatments showed identical inhibition (10 mm), implying that nanoparticle loading did not substantially improve efficacy against Proteus, possibly due to its intrinsic resistance mechanisms [22]. From Table 8, the MIC values revealed greater susceptibility of most microorganisms at 0.001 g/mL. In contrast, Aspergillus spp. and Staphylococcus aureus (for Fe-NC) required a higher concentration (0.010 g/mL), suggesting relatively greater resistance. Nevertheless, the inhibition zones produced by the nanocomposites remained significantly lower (p < 0.05) than those of the positive controls, indicating moderate but concentration-dependent antimicrobial efficacy.