1 Introduction

In recent years, increased anthropogenic activities including incessant use of chemicals in agriculture, production of batteries, leathers [1], iron, paint pigments, photographic materials, coatings [2], automobiles, aeronautical, glasses and steel, mining actions, and the use of leaded petrol for transportation [3], etc., have resulted into pollution of the environment with toxic metals, especially lead (Pb). Lead, an ancient element on the terrestrial planet and a non-essential toxic metal could occur naturally in the ecosystem but becomes an ecological and human health concern when present at levels beyond the permissible limit [4]. It affects biochemical and neurobehavioural functions and vital body systems such as hematologic, neurologic, gastrointestinal, renal, and cardiovascular systems [5].

According to Obeng-Gyasi et al. [6], Nigeria has the highest number of reports on Pb exposure in West Africa. About two decades ago, a major outbreak of lead poisoning occurred in two communities of Zamfara State from unsafe artisanal mining activities and processing of gold from lead-rich ore which led to an estimated death of four hundred children in 4 months [7, 8]. The soils and air of affected communities were heavily contaminated with Pb, and children were exposed through inhalation and ingestion of lead fine particles from lead ore grinding [7, 8]. Literature also revealed high levels of Pb in roadside dust particles, surface soils, and water of major cities of the country including Awka, Enugu, Onitsha, Aba, Port Harcourt, Nnewi, Warri [9], Abakaliki [10], Lagos, and Ibadan [11].

Several efforts are being adopted to alleviate the impact of heavy metals on the environment. Photocatalytic degradation [12, 13], chemical precipitation, hydroxide precipitation, sulfide precipitation, electrochemical methods, ion exchange, reverse osmosis, and adsorption methods are either too expensive to maintain, do not efficiently remove metal ions or generate waste [14, 15]. Thus, there is a rising interest in developing cost-effective and environment-friendly materials for cleaning up Pb from the environment. The use of biological technology to bioremediate heavy metals has proven to be a cheap, readily available, efficient, and eco-friendly alternative method [16, 17]. Biosorption involves the use of biological organisms such as plants, fungi, bacteria, algae, and some polysaccharide materials to remove heavy metals. They exhibit good biosorption capacities towards heavy metal, cover a large surface area, use an effective ion exchange, and operate over a wide range of environmental conditions [18].

Fungi are ubiquitous and possess a large cell wall material that enables excellent metal-binding pronounced tolerance and sorption properties towards metals [19]. Previous studies demonstrated the tolerance and biosorption capability of filamentous fungi from soil [18], lead–zinc mine tailing [20], and earthworm gut [21] to copper, chromium, nickel, lead, zinc, nickel, and cadmium. In the terrestrial environment, soils are the ultimate sink for heavy metals discharged into the environment via waste materials [22]. Consequently, microorganisms inhabiting heavy metal-polluted soils exhibit tolerance that may be important in the remediation (bioaugmentation) process [23]. While literature exists on the subject matter [18, 20, 21], there is a paucity of information on lead tolerance and biosorption capacity of fungi from dumpsite soils in Nigeria. Previous studies isolated fungi from dumpsite soils [24,25,26], however, their tolerance and biosorption capacity were not investigated. Therefore, the present study aims to screen mould isolates from dumpsite soils for tolerance to Pb2+ and to study their biosorption characteristics.

2 Materials and methods

2.1 Description of study areas

The selected areas of study are five major dumpsites in the Osogbo metropolis, Southwest, Nigeria (Fig. 1). Gbonmi, Ilesa Garage, Omobolanle, and Oke-Baale dumpsites are located within or close to residential areas inside the Osogbo metropolis while Egbedi is situated in a developing area outskirt Osogbo township. Egbedi dumpsite is surrounded by largely arable farmland and receives a large volume of waste from many communities within Osogbo, Ede, and environs. The dumpsites hosted different waste including kitchen waste, plastics, electronic and metal scraps, paper, broken bottles and ceramics, batteries, textiles, wood, leaves, leather, rubber, ashes, etc. Goats, dogs, chickens, birds, rodents, and lizards were seen on and around the dumpsites.

Fig. 1
figure 1

Map showing the location of the five dumpsites

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2.2 Collection of dumpsite soil samples

This was done following Dusengemungu et al. [23] with slight modifications. Debris at ten different locations on each site was cleared, and topsoil samples (0–10 cm deep) were randomly collected into ziplock bags and immediately conveyed to the laboratory on an icepack for further processing. The soils collected from each location were mixed to make a composite sample and used for microbiological analysis.

2.3 Isolation and morphological characterization of fungi

A composite soil sample (1 g) was weighed and dispensed into a well-labeled glass test tube filled with 9 mL of sterile distilled water, and serial dilution was done to the appropriate factor. Fungi were isolated using the standard spread plate technique on potato dextrose agar (PDA) media (Oxoid, England, UK) in duplicates. An uninoculated sterile PDA plate served as a control. The Petri dishes were incubated at 28 ± 2 °C for 5–7 days, and distinctive colonies were sub-cultured on sterile PDA plates to obtain pure cultures. Macroscopic identification of the isolates was done by observing the growth, colour, and texture of the colonies. For microscopic identification, 3–5-day-old pure cultures were stained in lactophenol blue on a clean slide and examined under a microscope (X40). Hyphae, conidiophore, and conidia appearance were observed and recorded. The fungal isolates were presumptively identified by comparing the colony and microscopic characteristics with [27].

2.4 Metal solutions preparation

Precisely, 10000 mg/L stock solution was prepared by weighing and transferring 15.98 g of lead nitrate (Pb(NO3)2) salt (Molychem, India) in a well graduated beaker, and made up to 100 ml mark with deionized water. Working solutions were obtained through serial dilution. All chemicals used in the present study were of analytical grade.

2.5 Minimum inhibitory concentration (MIC) and tolerance index of the fungal isolates

Firstly, the MIC of the fungal isolates to 5 concentrations of Pb2+ salt (200, 400, 600, 800, and 1000 mg/L) on amended potato dextrose agar was estimated. Thereafter, tolerance at increased Pb concentrations of 1500, 5000, and 10000 mg/L was determined. A fungal mycelial plug (10 mm diameter) was placed at the centre of the PDA plates amended with 1500, 5000, and 10000 mg/L, respectively, incubated at 28 ± 2 °C for 5–7 days, and observed for colonial growth. Radial growth was measured in millimetres using a measuring ruler. PDA plates without Pb salt served as control plates. The tolerance index (Tί) was calculated using the Eq. 1 below.

$${\text{T}}_{{\text{i}}} = {\text{ D}}_{{\text{t}}} /{\text{D}}_{{\text{u}}}$$
(1)

where, Tί = Tolerance index; Dt = fungal growth area in the presence of metal (mm); and Du = fungal growth area without metal exposure (mm).

2.6 Molecular characterization of the selected fungal isolates

Fungal DNA was extracted from a 5-day-old broth culture using the ZR Fungal/Bacterial DNA kit™ (Zymo Research, USA) according to the manufacturer’s instructions [28]. Quantification and purity of DNA were done using NanoDrop™ One spectrophotometer (Thermo Scientific, UK). The nuclear ribosomal internal transcribed spacer (ITS) region was amplified by PCR with previously described universal primers ITS4 (5′-TCC TCC GCT TAT TGA TATGC-3) and ITS5 (5′-GGA AGT AAA AGT CGT AAC AAGG -3′) [29]. PCR amplicons were analysed using 1.5% (w/v) agarose gel electrophoresis. The gel image was visualized with a Vilber Lourmat E-Box gel documentation imaging to confirm the size of PCR products. PCR products were cleaned up using an enzymatic method (ExoSAP-IT™ PCR Product Cleanup Reagent (Thermofisher, UK)) according to the manufacturer’s instructions. Sanger sequencing was performed using the Nimagen, Brilliant Dye™ Terminator Cycle Sequencing Kit V3.1, according to the manufacturer’s instructions. Pairwise sequence similarities of the generated consensus ITS regions were calculated using the Basic Local Alignment Search Tool (BLAST), for homology to identify the fungal isolates [30]. Sequences were deposited in the GenBank® and for accession numbers Assignment. The phylogenetic analysis employed partial ITS region sequences to establish relationships among our fungal isolates and some reference nucleotide sequences available in the NCBI GenBank® database. Multiple alignments of nucleotide sequences and phylogenetic tree reconstruction were carried out in MEGA 6, using the Neighbour-Joining method All positions containing gaps and missing data were eliminated [31, 32].

2.7 Biosorption capability of the selected fungal isolates

Exactly 1 ml of standardized spore suspension (106 spore/ml) was inoculated in 100 ml of sterile potato dextrose broth (PDB) containing 10000 mg/L Pb2+ salt and incubated at 28 ± 2 °C. Broth, without Pb2+ salt, served as control. The broth was filtered using Whatman filter paper No. 1 into well-labeled universal sterile bottles at days 5, 10, and 15 days. The harvested mycelium was dried at 60 °C for 20 min to obtain dry weight. Exactly 50 mL of filtrate was drawn, diluted, and digested to determine the residual Pb2+ concentration using an air-acetylene flame of atomic absorption spectrometer at 283.3 nm. All experiments were done in duplicate. In calculating the experimental data, the dilution factor(s) was considered and expressed as arithmetic average ± standard deviation.

Biosorption capacity was determined using Eqs. 2 and 3.

$${\text{q}}\,{ = }\,\left\{ {\frac{{{\text{C}}_{{\text{i}}} \, - \,{\text{C}}_{{\text{f}}} }}{{\text{m}}}} \right\}{\text{V}}$$
(2)
$$ {\text{R\% }}\,{ = }\,\left\{ {\frac{{{\text{C}}_{{\text{i}}} \, - \,{\text{C}}_{{\text{f}}} }}{{{\text{C}}_{{\text{i}}} }}} \right\}\,{*100} $$
(3)

where, q = sorption per gram biomass (mg/g), Ci = initial concentration of the metallic ions (mg/L); Cf = final concentration of metallic ions (mg/L); m = dried mass of the biosorbent in the reaction mixture (g); V = volume of the reaction mixture (mL); and R% = percentage removal of Pb2+.

2.8 Fourier-transformed infrared (FTIR) spectral analysis

The functional groups responsible for Pb2+ uptake on the isolates were examined by FTIR. Fungal biomass from the metal-loaded (10000 mg/L) and metal-free (control) samples were collected and lyophilized. Precisely 1 mg of lyophilized and crushed biomass was mixed with 0.1 g potassium bromide (KBr) [33]. Infrared spectra were analyzed on a Nicolet™ iS50 (Thermo Fisher Scientific, USA) FTIR spectrometer.

2.9 Scanning electron microscopy (SEM) analysis

The effect of Pb2+ on the surface morphology of the fungal mycelia was performed with SEM (Hitachi SU 3500 scanning microscope, Tokyo, Japan). The strain with the highest biosorption capability on the 5th day was selected. Fungal biomass of lead-free and lead-loaded samples was collected and washed repeatedly with phosphate buffer (pH 7.0) and fixed in 5% glutaraldehyde using the same buffer of pH 7.0 for 4 h. The sample was rewashed with the same buffer and dried in graded alcohol solution ranging from 30 to 100% twice with an interval of 30 min each. The dried sample was mounted on carbon tape and coated with platinum for 45 s prior to observation on SEM [34].

3 Results

3.1 Morphological characterization of fungal isolates

A total of 17 fungal isolates belonging to 10 species including Aspergillus flavus (1), A. niger (2), A. awamori (2), A. terreus (1), A. ochraceus (1), Alternaria sp. (1), Chrysosporium sp. (1), Penicillium chrysogenum (4), P. purpurogenum (1), Penicillium sp. (3). Were presumptively identified (Table 1).

Table 1 Morphological characterization of fungal isolates
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3.2 Frequency distribution of fungal isolates from the dumpsite soils

Five different fungal species were observed at Gbonmi, whereas it was 3 each at Ilesa Garage, Omobolanle, Egbedi, and Oke-Baale. Species of Aspergillus and Penicillium were noted in each study site (Table 2).

Table 2 Fungal species in the dumpsite soil samples
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3.3 Minimum inhibitory concentration and tolerance index of the fungal strains to the different concentrations of Pb2+

All the presumptively identified fungal isolates were resistant to Pb2+ concentrations at 200, 400, 600, 800, and 1000 mg/L (results not shown). At increased Pb2+ concentrations, Chrysosporium sp. (Gbonmi) and Alternaria sp. (Ilesa Garage) were susceptible at 5000 and 10000, 1500, 5000 and 10000 mg/L, respectively. The tolerance index varies depending on the isolate and Pb2+ concentration. Except in Oke-Baale, one fungus each from the sampling sites showed very high Pb2+ tolerance at 10000 mg/L (> 1.00), and showed a clear zone around colonial growth (Table 3; Fig. 2). These four isolates, Aspergillus awamori (Gbonmi and Ilesa Garage) and Aspergillus niger (Egbedi and Omobolanle) were selected for further studies.

Table 3 Tolerance index of fungal isolates at increased Pb2+ concentration
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Fig. 2
figure 2

Aspergillus species exhibiting tolerance on 10000 mg/L Pb amended plate and clearance zone around radial growth. a Aspergillus awamori (Gbonmi) (b) Aspergillus awamori (Ilesa Garage); (c) Aspergillus niger (Egbedi); and (d) Aspergillus niger (Omobolanle)

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3.4 Molecular identification

3.4.1 Quantity and quality of DNA obtained

The quantity of DNA obtained from the fungal isolates ranged from 10.5 ng/µL (Aspergillus niger—Egbedi) to 19.0 ng/µL (Aspergillus awamori—Ilesa Garage) (Table 4).

Table 4 DNA yield and purity
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3.4.2 PCR amplification, DNA sequence analysis, and evolutionary relationships among fungal isolates

The PCR product yielded about 650 bp after amplification of the ITS regions of the fungal strains (Fig. 3). DNA BLASTn analysis (Table 5) shows that A. awamori OP341445 and OP341446 were closest to strain ON054315 and ON054316 correspondingly with 100% similarity. Likewise, A. niger OP342447 and OP341448 recorded a 100% resemblance with MT316340 and MK450668, respectively. All our fungal isolates belong to a single clade (Fig. 4).

Fig. 3
figure 3

Gel image of the four fungal strains showing amplicon size for the ITS regions. Well 1– DNA marker; Well 2—Aspergillus awamori (Gbonmi); Well 3—Aspergillus awamori (Ilesa Garage); Well 4—Aspergillus niger (Omobolanle); Well 5—Aspergillus niger (Egbedi)

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Table 5 BLASTn analysis summary of the four fungal strains
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Fig. 4
figure 4

Neighbour-joining tree showing the phylogenetic position of OP341445, OP341446, OP341447 and OP341448, based on the ITS 4 and 5 gene sequence

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3.5 Fungal biomass, biosorption potential, and Pb2+ uptake

Generally, the yield of biomass for the selected fungi increased with the days of incubation. The least yield was obtained on day 5 (ranging between 0.65 and 0.86 g dry cell L−1), and the maximum on day 15 (ranging from 0.90 to 1.12 g dry cell L−1) (Table 6). Conversely, biosorption capacity was at the peak on day 5 with a gradual decrease on days 10 and 15 in all the isolates except OP341448 whose highest biosorption capacity was recorded on day 10 (97.82 mg/ g) and lowest on day 5 (77.91 mg/g) (Table 6). Interestingly, the Pb2+ removal rate for the fungi was approximately 86% on the 3 days except for OP41447 and OP41448 with ≈87% on day 15 (Fig. 5).

Table 6 Biosorption capacity of the selected fungal isolates
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Fig. 5
figure 5

Lead ion uptake by the selected fungal strains (Error bar represents means ± SD, n = 2)

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3.6 FTIR analysis of functional groups on Pb2+ untreated and treated fungal biomasses

The characteristic infrared peaks of Pb2+ unloaded fungal biomass (c) and Pb-loaded fungal biomass (t) are presented in Table 7. The results obtained revealed the involvement of functional groups O-H, C-H, N-H, O=C=O, C≡C, N=C=S, C=C, N-O, C-N, and S=O in the biosorption process. Loading of the biomasses with Pb gave rise to unchanged peak intensity (OP341445), decreased peak intensity (OP341447), the appearance of a new peak (OP341446t), and disappearance of peak (OP341448t) corresponding to O-H stretching alcohol. Interestingly, the appearance of peaks identified as C-H stretching of alkyne was observed in OP341446(u) and OP341448(t). Decreased peaks assigned to N-H stretching amine salt were observed in all the loaded fungal biomasses. An additional peak was discovered at approximately 2344.5 cm−1 (O=C=O stretching carbondioxide) in OP341448(u). After Pb2+ treatment, there was either an upward or downward peak shift assigned to C≡C stretching alkyne (OP341446, OP341337 and OP341448); and C=C stretching aromatic compound (OP341445, OP341446 and OP341448); N–O stretching nitro compound (OP341446 between 1401.5 and 1420.1, and OP341445, OP341447 and OP341448 between 1345.6 and 1375.4 in OP341445 and OP341445); C-O stretching alkyl aryl ether (OP341446 and OP341448) and S=O stretching sulfoxide (OP341448). Interestingly, an unchanged peak intensity of 1148.0 (O-H stretching tertiary alcohol), 1625.1 (C=C stretching aromatic compound), 1036.2 (S=O stretching sulfoxide), and 1375.4 (O-H bending alcohol) were also noted in the four fungal biomasses, OP341448, and OP341445, respectively.

Table 7 The frequencies (cm−1) and assignments of FTIR bands for the Pb untreated and treated fungal mycelia
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New peaks 2158.1 (S-C≡N stretching thiocyanate), 2027.7 (N=C=S stretching isothiocyanate), 1285.9 (C-O stretching aromatic ether), and 1025.0 (C-N stretching amine) were observed in OP341445(t). Similarly, in OP341446, OP341447 and OP341448 treated biomasses, new peaks of 1375.4 and 1408.9 (corresponding to O-H bending alcohol) were observed. Additional peaks 1077.2 (C-O stretching primary alcohol) and 1021.3 (C-N stretching amine) were noted in OP341446(t). Two new peaks were also added in OP341448(t) i.e., 1543.1 (N-O stretching nitro compound) and 1304.6 (C-N stretching amine).

Peak disappearances were noted in the treated biomasses, except in OP341448. In OP341445(t), peaks corresponding to C-N stretching amine, C-O stretching alkyl aryl ether, C-O stretching ester, C-O stretching primary alcohol and S=O stretching sulfoxide were missing. In OP341446(t), it was peaks assigned to C-H bending aromatic compound, O-H bending carboxylic, C-N stretching amine, and S=O stretching sulfoxide. Similarly, peaks corresponding to C=C stretching aromatic, C-O stretching alkyl aryl ether, and S=O stretching sulfoxide were not found in OP341447(t).

3.7 Scanning electron microscopy (SEM) analysis

A scanning electron microscope was used to assess morphological changes in the Aspergillus strains before and after Pb2+ biosorption. The control fungal cells had normal conidia, and were intact, smooth, and closely connected with each other (Fig. 6a). After loading Pb2+, the cells appeared to shrink, wrinkle, and deform with some bright areas (Fig. 6b).

Fig. 6
figure 6

Representative SEM images of the selected fungal strains (Aspergillus awamori -OP341445) before (a) and after (b) adsorbing Pb2+

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4 Discussion

A foremost environmental concern in many developing countries is the indiscriminate release and disposal of wastes especially those containing toxic elements [35]. Fortunately, dumpsite soils harbour microorganisms capable of degrading complex organic materials [24], and those able to withstand the levels of heavy metal persist in the soil environment. The present study isolated 10 different fungal species belonging to the genera Aspergillus, Alternaria, Chrysosporium, and Penicillium. Previous studies reported the dominance of different species of Aspergillus, Fusarium, Mucor, Penicillium, Saccharomyces, Trichoderma, and Rhizopus in municipal solid waste dumpsite soils [25, 26]. Aside from the Chrysosporium species, other isolates obtained in our work had been reported elsewhere [36, 37]. The distribution of fungal isolates (Table 2) is similar to results obtained from major dumpsites in the South-South region of Nigeria including Port Harcourt [25, 38] and Benin [24, 26].

Contaminated sites are prime sources of heavy metal-tolerant species with native fungi exhibiting notable tolerance to toxic elements [39]. In this study, all the isolated fungi survived the test concentrations of Pb2+ except, Chrysosporium sp. (Gbonmi) and Alternaria sp. (Ilesa Garage) at 5000 and 10000, and 1500, 5000, and 10000 mg/L, respectively (Table 3). According to Ruta et al. [40], metal tolerance by moulds depends on sample location, the toxicity of the element, heavy metal level in the medium, and importantly, the isolate’s competence. The remarkable tolerance demonstrated by A. awamori (OP41445 and OP41446) and A. niger (OP41447 and OP41448) in this study (Table 3) may be associated with metal binding to cell walls, production of intracellular/extracellular enzymes, intracellular sequestration, extracellular metal sequestration and precipitation, suppressed influx, enhanced metal efflux, and complexation [19, 36]. Previous studies reported notable metal tolerance of Aspergillus species including A. aculeatus [41], A. sclerotiorum, and A. niger [42]. Other fungi including, Verticillium insectorum J3 [20], Komagataella phaffii, Trichoderma harzianum [31], Fusarium solani [42], and Trichoderma brevicompactum QYCD-6 [21] were also noted for their resistance to heavy metals.

In this work, biomass yield increased with days of incubation, and the highest biosorption capacity was noticed on day 5 in all the isolates except OP341448 (day 10) (Table 6). Feng et al. [20] stated that biosorption was not determined by the biomass but was related to the metal concentrations. Generally, metal ion uptake involves a primary swift phase (passive uptake), followed by a slower secondary stage (active phase). The initial stage is physical adsorption or ion exchange at the surface of the biomass and accounted for a major part of total metal uptake, while the second one contributed to a minute part [39, 43]. Additionally, biosorption reduces when the surface-active sites on the fungal cell wall are completely covered with metal ions [44]. Thus, the removal of a metal ion is strongly dependent upon the availability of a free active binding site on the fungal cell wall. The Pb2+ removal rate for the fungi was approximately 86% on days 5, 10, and 15 except OP41447 and OP41448 with ≈87% on day 15 (Fig. 5). Feng et al. [20] reported adsorption equilibrium on the 8th day with approximately 90% Pb2+ removal. These suggest that the absorption principally occurs during the logarithmic growth phase.

FTIR spectra give details on the probable cell-metal ion interactions in metal untreated and treated biomass [44]. The main functional groups i.e., O-H, C-H, N-H, O=C=O, C≡C, N=C=S, C=C, N-O, C-N, and S=O identified to be involved in the biosorption in this work were earlier reported [20, 45, 46]. The change in vibration frequency or the intensities of peaks upon loading with Pb2+ reveals the interaction of fungal biomasses treated with Pb2+. According to [47], band intensity is expected to change upon reaction with metal ions because of new links among functional groups on the biomass surface. In this work, the observed consistency in band wavelength 3261.4 and 1375.4 (OP341445), 1625.1 (OP341448), 1148.0 (OP341445, OP341446, OP341447, and OP341448), and 1036.2 (OP341448) after absorbing Pb2+ in suggest possible ion exchange and/or complexation with Pb2+. Additionally, the appearance and disappearance of peaks indicate that Pb2+ forms stable bonds after chelation `between oxygen-, nitrogen-, sulphur-, or phosphorus-containing ligands of biomass [48]. Thus, functional group assignments participate not only in metal biosorption but also in possible ion exchange and/or complexation processes [49].

In biosorption, morphological change is important for Pb2+ removal [50, 51]. In the current study, the appearance of the control cells (Fig. 6a) provided a large surface area for metal uptake while the structure of the treated cells was due to the accumulation of lead (Fig. 6b). Lethal heavy metals hinder growth, causing morphological changes and affect the reproduction of organisms especially spore formation and conidia production [44, 52].

5 Conclusion

The study isolated 17 fungi belonging to 10 different species from Gbonmi, Ilesa Garage, Omobolanle, Egbedi, and Oke-Baale dumpsite soils in the Osogbo metropolis and environ, Southwest, Nigeria. All the isolates tolerated different levels of Pb2+ concentration, however, four i.e. A. awamori OP341445 and OP341446 and A. niger OP342447 and OP341448 demonstrated outstanding resistance at 10000 mg/L. Although the biomass yield for the strains was highest at day 15, the highest biosorption capacity was observed on day 5 except in OP341448 (day 10) and the equilibrium removal rate was noted on day 5. FTIR analysis revealed that these functional groups: O-H, C-H, N-H, O=C=O, C≡C, N=C=S, C=C, N-O, C-N, and S=O contributed towards Pb2+ uptake. SEM demonstrated the cellular changes induced by the heavy metal. To the best of our knowledge, there are no studies on the Pb2+ tolerance and biosorption capacity of dumpsite soil fungi, and hence the strains A. awamori OP341445 and OP341446 and A. niger OP342447 and OP341448 are promising mycoremediation agents. Further studies would include simulation and field experimental studies to establish the efficiency of these fungi in bioremediating lead.