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Biodegradation of azo dyes by Aspergillus flavus and its bioremediation potential using seed germination efficiency

BMC Microbiology volume 25, Article number: 7 (2025) Cite this article

Abstract

The worldwide textile industry extensively uses azo dyes, which pose serious health and environmental risks. Effective cleanup is necessary but challenging. Developing bioremediation methods for textile effluents will improve color removal efficiency. The recent attention to effectively utilizing microbes to convert toxic industrial azo dyes into non-hazardous compounds has garnered significant attention. In the present study, four fungal strains—Aspergillus flavus, Aspergillus terreus, Aspergillus niger, and Fusarium oxysporium—were employed to screen for the degradation and detoxification of azo dyes including congo red, crystal violet, bromophenol blue, and malachite green. After eight days, A. flavus had degraded azo dyes at the maximum proportion. The maximum decolorization (%) was achieved at 50 mg/L of dye concentration, 8 days of incubation, pH 6, 30 °C temperature, sucrose as a carbon source, NaNO3 as a nitrogen source, Ca+2 as minerals, and using static culture. The efficient production of laccases, lignin peroxidase, and manganese peroxidase enzymes by A. flavus proved that the enzyme played a crucial role in decolorizing the harmful azo dyes. The Fourier Transform Infrared spectrometer (FT-IR) data validated the decolorization and degradation process brought on by absorption and biodegradation. Compared to control plants, the results of the phytotoxicity assay showed that the degraded product was less harmful to maize and common bean plant's growth and germination rates. As a result, the findings indicate that A. flavus is a viable option for remediating azo dyes. This aids in the biodegradation of azo dyes found in wastewater.

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Introduction

Chemicals are being used and released into the environment due to the world's rapid industrialization to meet the demands of the expanding population [1]. Many different types of dyes are utilized in the industrial processes of the food, plastic, textile sectors, and pharmaceutical industries [2]. An estimated 7 × 105 tons of dye is produced annually, yielding 100,000 commercial dyes; nevertheless, a larger portion of this dye is released into water sources as waste during industrial processes [3]. There are various types of textile dyes, including basic, acidic, reactive, and direct dyes, all of which are essentially water-soluble. These types are naturally persistent and can cause significant harm to marine life by blocking the passage of light, slowing down photosynthesis, reducing oxygen levels, and ultimately leading to the death of organisms [4]. Azo dyes are resistant to light, moisture, and oxidants due to their intricate aromatic structures. This resistance comes from the presence of azo (-N = = N-) bonds connected to aromatic amines and benzene or naphthalene rings, which make them resistant to cleavage [5]. This is an attribute that the industry desires, but it poses a risk to the environment.

Synthetic dyes offer numerous benefits over natural dyes, including greater stability, a wider range of colors, lower production costs, and easier application [6]. These benefits have led to their use in various industries, including food, paper, plastic, textiles, rubber, cosmetics, pharmaceuticals, petroleum additives, and photography [7, 8]. Synthetic dyes are mostly used by the textile industry to color its fabrics. Due to the inefficiency of the dyeing process, which results in 200,000 tons of colored fluids flowing to water bodies annually, 10–15% of colored waters are lost to effluents in these procedures [9]. As previously mentioned, synthetic dyes are highly stable and pose a significant environmental risk because of their extended environmental persistence and poor biodegradability [10, 11]. Water containing artificial dyes reduces the amount of aquatic biodiversity by obstructing sunlight and causing issues for photosynthetic aquatic plants and algae [12]. Numerous artificial dyes are poisonous, carcinogenic, and mutagenic [13]. Furthermore, dyes have the potential to build up in sediments, particularly in areas where wastewater is discharged, and to disrupt the aquatic system's natural equilibrium. Contaminant leaching may have an impact on the groundwater system [14].

Various physical and chemical methods are available for removing dyes from wastewater [15]. The principal chemical techniques are photocatalysis, electrochemistry, Fenton's reaction, ozone, and hydrogen peroxide oxidation [16, 17]. In addition to their higher costs and operational challenges, chemical and physical treatments exhibit low efficacy, limited adaptability, susceptibility to interference from other wastewater constituents, and difficulties in managing created residues [18].

In recent decades, there have been suggestions to use biological techniques as a practical and efficient alternative to synthetic dyes in addressing the dye problem. These techniques rely on bioremediation, which involves using microorganisms to catalyze the complete breakdown or conversion of toxic substances into less harmful or environmentally friendly forms [19]. Numerous studies have confirmed the significant potential of various bacterial, fungal, and algal species in dye removal [20,21,22]. Dye removal can be achieved through biomass adsorption, biodegradation, or both methods. Dye biotransformation can lead to complete mineralization or the production of less harmful byproducts [23].

Fungi can transform aromatic compounds such as pesticides, lignin, and polycyclic aromatic hydrocarbons (PAH) through an extracellular enzymatic system. Currently, procedures for fungus decolorization are gaining significant attention [24]. Biodegradation and biotransformation enzymes can be obtained from fungal biomass and used as a sorbent. The biosorption process is characterized by its speed, effectiveness, and adaptability to various types of textile effluents [25,26,27]. Aspergillus niger and Fusarium oxysporium are two examples of non-ligninolytic fungi that exhibit biosorption; their dead biomass can be utilized as an adsorbent [28, 29]. White rot fungus produces enzymes such as lignin peroxidase (LiP) and Mn-dependent peroxidase (MnP), and laccase (Lac), and is the best studied. Its non-specific enzymatic activity enables it to break down a wide range of aromatic molecules and mediated dye decolorization [30,31,32]. Additionally, it was mentioned that non-ligninolytic enzymes might be involved in the breakdown of dyes like triphenylmethane crystal violet [30]. Dye was eliminated from tested strains' living biomass more successfully than their dead biomass [33]. The overall mechanism by which ligninolytic enzymes work involves the generation of free radicals, which set off a sequence of intricate events that ultimately cause the pigments to break [34]. The efficiency of decolorization is mostly dependent on the strain employed in the procedure, and the functional groups in the cell wall [35,36,37,38,39,40]. The sources and concentrations of carbon and nitrogen are crucial variables because they impact the ligninolytic enzyme synthesis. Numerous studies have been conducted on the impact of various carbon and nitrogen sources on the efficiency of decolorization [36, 37].

To determine the most effective fungus, we conducted a screening of the biodegradation capabilities of Aspergillus flavus AUMC8653, Aspergillus niger AUMC8669, Aspergillus terreus AUMC8625 and Fusarium oxysporum MT032355 on various azo dyes. Our goal was to maximize decolorization under different incubation conditions such as pH, temperature, carbon sources, nitrogen sources, and metal salts. We also aimed to study the enzymatic processes involved in breaking down azo dyes and the resulting degraded products. After treatment, we analyzed the fungal biomass using FT-IR to assess surface sorption. Finally, we conducted a phytotoxicity test using Zea mays and common bean seeds as the last step in the environmental risk assessment process.

Materials and methods

Chemicals and reagents

All chemical and azo dyes (congo red, crystal violet, bromophenol blue and malachite green) were purchased from Sigma-Aldrich Pvt. Ltd. Stock solutions of azo dyes (200 mg/L) were made by dissolving azo dyes into sterilized and distilled water, then filtered with a Whatman filter paper (No. 1) then kept in a refrigerator at 4 ± 1 °C until further used.

Microorganism

A. flavus AUMC8653, A. niger AUMC8669, A. terreus AUMC8625 and F. oxysporum MT032355 were used in this study.

Screening of microorganisms' efficiency

For the experiments, Czapek's Dox broth medium was used for the growth of A. flavus, A. niger, A. terreus, and F. oxysporium. Distilled water was used to dissolve the medium and 1% stock dye solution was added, and the medium was autoclaved for 20 min at 121 °C to ensure sterilization. An orbital agitator (120 rpm) at 30 ± 2 °C was used to inoculate 10 ml of spore suspension of each fungal strain with 50 mg/L of congo red, crystal violet, bromophenol blue, and malachite green. The degradation of dye was evaluated after seven days by measuring the absorbance of each dye at its maximum using a UV–Vis spectrophotometer. Every experiment was repeated three times, and the results were compared to the control. The following formula was used to calculate the percentage of decolorization according to Shah et al. [41]:

$$\text{Percentage of decolorized dye }=\frac{\text{Reading of initial absorbance}-\text{ Reading of final absorbance}}{\text{Reading of initial absorbance}}\times 100$$

Determination of growth

The fungal mycelium, the control (untreated), and the treated samples were centrifuged for 10 min at 10,000 × g. The pellets were obtained and then washed three times with sterile distilled water. Then, the fresh weight (FW) was recorded. The biomass was then dried at 60°C until it reached a consistent weight to measure the dry weight (DW).

Optimization of the decolorizing ability of the most potent fungal isolate

The most potent fungal isolate (A. flavus) was used to study the effects of various culture conditions, including incubation periods, pH, temperature, different carbon, nitrogen sources, metals, and static and shaking conditions on the decolorization of congo red, crystal violet, bromophenol blue, and malachite green.

Effect of different dye concentrations on decolorization percentage (%)

A. flavus AUMC8653 was inoculated into Czapek's Dox broth medium after different azo dyes were individually added at various doses (50, 100, 150, and 200 mg/L) as the sole carbon source. After an 8-day incubation period, the highest λ max of each dye was used to calculate the decolorization percentage (%).

Effect of different incubation periods, pH values, and temperature on decolorization percentage (%)

A. flavus AUMC8653 was inoculated into 250 mL conical flasks containing 50 mL of Czapek's Dox medium with the addition of 50 mg/L of each dye. The flasks were placed in an incubator at temperature of 30 ± 2 °C for 2, 4, 6, 8, and 10 days. The percentage of dye decolorization (%) was recorded. Another experiment was carried out using different pH values (5, 6, 7, and 8). After eight days of incubation at 30°C, the percentage of dye decolorization was calculated for each group. Another experiment was conducted by incubating flasks at different temperatures: 20, 25, 30, 35, and 40°C at pH 6. Three independent experiments were performed and the percentage of dye decolorization was determined for each flask.

Effect of different carbon and nitrogen sources on the dye decolorization

A. flavus AUMC8653 was inoculated in 50 ml of Czapek's Dox broth medium containing different carbon sources: glucose, fructose, sucrose, maltose, lactose, or starch with the addition of 50 mg/L of azo dyes, individually. Another experiment was conducted using different nitrogen sources (NaNO3, peptone, and yeast extract at 2 g/L and equivalent nitrogen for NH4NO3, KNO3, and urea). All flasks were then incubated at 30 ± 2 °C for 8 days. Each experiment was conducted three times, and the percentage of decolorization was measured.

Effect of some heavy metals on the decolorization activity

A. flavus AUMC8653 was inoculated in 50 ml of Czapek's Dox liquid medium with the addition of 100 ppm concentrations of Mg, Cu, Ca, Ni, and Co, individually. Two sets of experiments were conducted with and without 50 mg/L of azo dyes. Each experiment was repeated three times, and the percentage of decolorization was determined.

Effect of static and shaking incubation on dye decolorization

The Czapek Dox liquid medium was inoculated with A. flavus AUMC8653 along with the addition of 50 mg/L of azo dyes, individually. The flasks were then incubated at 30 ± 2 °C for 8 days under both static and shaking conditions at an agitation rate of 120 rpm. Afterward, samples were centrifuged at 10,000 rpm for 20 min. The supernatant was utilized to determine the percentage of decolonization.

Biochemical and enzyme assay

A. flavus AUMC8653 was inoculated in Czapek's Dox liquid medium with the addition of 50 mg/L of azo dyes for 120 h. The resultant pellet (fungal biomass) was utilized to analyze the protein and H2O2 content, and the resulting supernatant was employed to study extracellular enzymes as a crude enzyme extract.

According to Srinu et al. [42], the laccase activity (EC 1.10.3.2) was measured using guaiacol as the substrate. The absorbance of the sample was measured at 450 nm. In addition, manganese peroxidase (Mnp) (EC 1.11.1.13) was measured using guaiacol as the substrate. MnP activity was determined by measuring the absorbance of the obtained sample at λ465 nm using a UV–Vis spectrophotometer [43]. Lignin peroxidases (LiP, EC 1.11.1.14), the reaction was measured at 310 nm in a UV spectrophotometer [42]. The intracellular protein content was determined according to the method of Lowry et al. [44], and the H2O2 content of the fungal biomass was assessed using the procedure outlined by Velikova et al. [45].

FT-IR

FT-IR analysis investigated the degradation of various azo dyes and assessed the toxicity of degraded compounds through functional group changes. Ethyl acetate was used as a mid-polar solvent for the liquid–liquid extraction of the broken-down metabolites. To eliminate the moisture in the sample, potassium bromide was heated to 105 °C for 30 min. To make it in the ratio of 5:95 (w/w) for analysis, it was further powdered. A total of sixteen scans were conducted in the mid-IR 400–4000 cm−1 band to record the results [46].

Light microscopy

The mycelia were filtered from Czapek's Dox liquid medium containing azo dyes and photographed using a Microscope.

hytotoxicity study

Seeds of maize and common bean were obtained from the Agricultural Research Center in Giza, Egypt. The seeds of maize and common bean were sterilized with 0.1% sodium chlorite for 4 min, followed by five washes with sterile distilled water. Afterward, they were planted in pots filled with 8 kg of air-dried clay loam soil in a completely randomized design (CRD). The pots were split into four groups, each consisting of three replications, and kept at a temperature of 35 °C and a relative humidity between 60–75%. The first group contained only soil irrigated with water, serving as the control. The second group was irrigated with filtrate of A. flavus, the third group was irrigated with 50 mg/L azo dyes, and the fourth group was irrigated with azo dyes after biodegradation. The pots were watered every two days. The morphological criteria of the plants were observed 21 days after sowing.

Seed germination percentage, shoot length, root length, fresh and dry weight of seedlings, vigor index I, and vigor index II were measured after 21 days of treatment. Seeds germinated after 7 days were considered as germinated seeds, and germination% was calculated using the following formula [47, 48]:

$$\mathrm {Germination \, \%} = \frac {\mathrm {Number \,of \,seeds \,germinated}} {\mathrm {Total \,number \,of \,seeds}} \times 100$$

Seedling vigor index I and seedling vigor index II were determined using formulas:

$$\mathrm{Vigor \,index \,I }=\mathrm{Germination\, \%}\times \mathrm{ seedling \,length }$$
$$\mathrm{Vigor \,index \,II }=\mathrm{Germination \, \%}\times \mathrm{dry \,biomass }$$

Statistical analysis

Each treatment consists of three replicates. The greenhouse experiment was organized using completely randomized designs (CRD). Utilizing SPSS Software version 9, the data were subjected to an ANOVA, and the means of treatments were compared using the Duncan multiple range test (P < 0.05).

Results and discussion

As a result, the need to remediate wastewater contaminated by dyes is becoming essential [49]. It has been challenging for scientists to develop a unified and cost-effective technique for treating dyes in textile effluent [49].

Screening of the obtained fungal isolates for decolorization of azo dyes

The fungal isolates were tested for their ability to decolorize azo dyes at a concentration of 50 mg/L (Fig. 1). The results showed that A. flavus exhibited significantly higher decolorization ability followed F. oxysporium compared to the other isolates. A. flavus showed the greatest decolorization capacity of congo red, crystal violet, bromophenol blue, and malachite green by approximately 89.0%, 88.0%, 85.0%, and 82.0%, respectively. Hefnawy et al. [50] results are in line with this observation. They discovered that Aspergillus flavus and Penicillium canescens achieved the highest decolorization value of direct blue dye when the initial dye concentration was 0.01%. However, this value decreased at concentrations higher than 0.01%, likely due to dye toxicity.

Fig. 1
figure 1

Decolorization of azo dyes (50 mg/L) by different fungal strains. Data represent mean values of three replicates ± show standard error (SE)

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Effect of dye concentrations on dry weight of fungal

The data in Fig. 2 demonstrated that the fungal dry weight (g/100mL) of A. flavus AUMC8653 declined above a concentration of 100–200 mg/L and increased at lower concentrations of the dye in the growth medium up to a concentration of 50 mg/L. This indicates that the ability of A. flavus to remove dye was diminished as dye concentrations increased. The fungal dry weight increased most noticeably when they were cultivated in medium with 50 mg/L of congo red (1.02g/100mL), crystal violet (0.92g/100mL), bromophenol blue (0.88g/100mL), and malachite green (0.83g/100mL). Congo red consistently resulted in the highest fungal weight among all azo dyes tested. The data presented in Fig. 2 indicates that a dye concentration of 100 mg/L is considered sub-lethal, which negatively affects the dye removal capacity of the four azo dyes. High dye concentrations are toxic to fungal growth and hinder dye decolorization, and there is limited information available about fungal dye decolorization at this concentration. As a result, a dye concentration of 50 mg/L was selected for further investigation in this study. These findings align with Singh and Singh [51] study which found that the supplementation of bromophenol blue and congo red to potato dextrose agar medium individually suppressed the growth of A. flavus compared to their respective controls. Kunjadia et al. [52] found that the increment of crystal violet concentration caused a reduction in the mycelial growth of Pleurotus ostreatus MTCC142. Furthermore, increasing the dye concentration decreased the enzymes' ability to degrade azo dyes [53, 54]. Furthermore, 97.41% of the reactive red HE7B dye can be eliminated by Aspergillus salinarus strains cultivated on potato dextrose broth medium containing 50 mg/L of the dye [55].

Fig. 2
figure 2

The effect of different concentrations of azo dyes on dry weight of A. flavus. Data represent mean values of three replicates ± show standard error (SE)

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Many studies have shown the toxicity of azo dyes to microorganisms involved in biodegradation. The type, concentration, and dye particle blockage of azo reductase enzyme active sites all affect toxicity [56]. The toxicity of dyes to fungal cells, particularly at elevated concentrations, may result from factors such as high molecular weight, structural complexity, and the presence of inhibitory groups like sulfonic acid [57]. According to Fetyan et al. [58], Saccharomyces cerevisiae showed higher decolorizing levels in direct blue71 achieving 100% activity. However, at concentrations exceeding 200 ppm, the rate of decolorization decreased. This decline may be attributed to the potential toxicity of higher concentrations on microbial cells. Furthermore, compared to their respective controls, adding bromophenol blue and congo red to potato dextrose agar media significantly inhibited the growth of Aspergillus sp. [50].

Effect of different incubation periods on dye decolorization percentage

The data presented in Fig. 3 illustrated that extending the incubation period from 2 to 10 days led to a notable increase in the percentage of decolorization across all concentrations of azo dyes used. The highest decolorization percentages were detected for congo red (93.5%), crystal violet (90.4%), bromophenol blue (85.4%), and malachite green (80.6%) dyes during eight days of incubation and 50 mg/L of azo dye. The decolorization of dye through adsorption and breakdown is primarily influenced by the duration of the incubation period [59]. The degree of decolorization significantly decreased both before and after the 8-day incubation period. This decrease in decolorization may be due to the efflux mechanism in fungal cells, which reduces the dye's toxicity. After eight days, there could have been a decline in the decolorization process. The decrease in fungal growth and metabolic activities could have been attributed to enzymatic biodegradation activity, physical binding of dye to fungal biomass, and dye buildup [50].

Fig. 3
figure 3

Effect of incubation period, initial medium pH and temperature on decolorization percentage of 50 mg/L of azo dyes by A. flavus

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Effect of different pH on dye decolorization percentage

The rate of dye decolorization and biomass formation is significantly influenced by pH [60]. The pH level may affect the surface of mycelium, which is composed of biopolymers with various functional groups. Chitin, proteins, polysaccharides, and other compounds present on the mycelium surface contain NH3+ and COOH groups. These groups can interact with dye anions through electrostatic interactions [60]. Our results found that A. flavus is most effective at decolorizing azo dyes at pH 6 (Fig. 3). This suggests that the acidic pH range is ideal for A. flavus to decolorize these dyes. According to our findings, A. flavus AUMC8653 was able to decolorize dyes as follows: 94.0% of congo red, 91.0% of crystal violet, and 86.0% of malachite green at pH 6.0. The results align with those reported by Hefnawy et al. [50] who found that A. flavus can thrive and decolorize within a broad pH range of 3 to 8, with the optimum pH being 4. This indicates that the strain can be valuable for treating wastewater and for various industrial uses. Additionally, these findings suggest that acidic pH levels may affect the stability of the enzymes [59]. The decolorization efficiency of A. flavus A5p1 declines significantly as pH increases [61], due to the interaction of positively charged cell surfaces with negatively charged dye anions, as demonstrated in the instance of congo red and coir pith carbon [62]. Dyes tend to decolorize at higher concentrations in acidic environments, which facilitates their removal through enzymatic activity or adsorption onto the fungal cell wall, according to Namdhari et al. [57]. It is well known that [H +] ions regulate dye degradation, which is influenced by the pH of the medium.

Effect of different temperatures on dye decolorization percentage

The impact of incubation temperature on the decolorization of 50 mg/L of azo dyes was investigated throughout a temperature range of 20–40°C. According to the results, 30°C was the most effective temperature for decolorizing congo red (91.0%), crystal violet (90.3%), bromophenol blue (85.3%), and malachite green (84.5%) dyes. At higher temperatures (40°C), the decolorization % was decreased (Fig. 3). This decrease may be due to the loss of cell viability [50] or to the denaturation of ligninolytic enzymes. This means that the rate of color removal increases with temperature [50, 63]. The azo dye reactive violet 1 decolorization efficacy of Ganoderma cupreum AG-1 was higher at lower temperatures (25–35°C) [64]. According to the study of Ameen et al. [64] who found that the optimum temperature range for Aspergillus strains to decolorize dyes was 30–35°C. The optimum temperature for each organism promotes growth and metabolic activities. Furthermore, Singh and Dwivedi [65] found the maximum decolorization of congo red (92.60%) with A. flavus treatment at pH 6.0, 30°C, 120 rpm stirring speed, and 120-h duration. During the adsorption or decolorization process, the ideal temperature critically influences the kinetic energy and surface energy of dye molecules. It is also necessary for the release of enzymes during degradation that break down dye [66, 67].

Effect of different carbon sources on dye decolorization

Fungi need nutrient supplements like carbon and nitrogen to grow and to break down contaminants. Glucose, fructose, and sucrose have been extensively studied as carbon sources for fungal growth; while starch and xylan also appear to be important carbon sources [68]. The data in Fig. 4 presented that A. flavus AUMC8653 could effectively use certain carbon sources such as fructose, sucrose, glucose, and maltose to achieve high decolorization percentages. However, the growth of A. flavus AUMC8653 was decreased in the presence of starch and lactose, resulting in lower decolorization values. Sucrose was identified as the best carbon source for A. flavus AUMC8653, leading to the highest decolorization percentages for congo red (94.4%), crystal violet (93.2%), bromophenol blue (88.2%), and malachite green (88.3%) dyes. Similar results were observed in a study by Hefnawy et al. [50] when direct blue dye decolorization was evaluated after A. flavus and P. canescens were grown in various carbon sources, such as glucose, sucrose, fructose, or maltose. These factors are crucial for the efficient breakdown of dyes [69, 70]. The ability of microbes to break down and decolorize azo dye depends on the types and availability of carbon–nitrogen sources, which serve as electron donors during the decolorization process [24].

Fig. 4
figure 4

Effect of different carbon and nitrogen sources on decolonization of 50 mg/L of azo dyes by A. flavus. Data represent mean values of three replicates ± show standard error (SE)

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Effect of different nitrogen sources on dye decolorization

The highest decolorization percentage was achieved when NaNO3, NH4NO3, urea, and yeast extract were supplemented with the growth medium of A. flavus (Fig. 4). It was also noted that the lowest decolorization percentage of various azo dyes was obtained in the presence of KNO3 in the growth medium. On the other hand, NaNO3 was identified as the most effective carbon source utilized by A. flavus AUMC8653, leading to the highest decolorization percentages for congo red (95.7%), crystal violet (93.9%), bromophenol blue (90.4%), and malachite green (89%). Similar results were published by Hefnawy et al. [50], who noted that A. flavus and P. canescens displayed substantial growth and decolorization percentages of direct blue dye when using all nitrogen sources except for KNO3. The ideal concentration of nitrogen sources was 0.5% of ammonium sulfate and 0.2% of sodium nitrate for the degradation of azo dyes by Trichoderma virens, Phlebiopsis cf. ravenelii, Talaromyces stipitatus, and A. niger [71].

Effect of some heavy metals on the dye decolorization

Metal ions are considered to be potential laccase inhibitors because these ions bind to the enzymes and de/stabilize the protein and they change the enzyme activity [50, 65]. The results in Fig. 5 revealed that Mg+2, Ca+2, and Cu+2 substantially improved the decolorization of azo dyes compared to the control. In contrast, the presence of Ni+2 and Co+2 in the growth medium resulted in a decrease in the percentage of color reduction. Ca+2 proved to be the most effective metal utilized by A. flavus AUMC8653, resulting in the highest decolorization percentages for congo red (95.5%), crystal violet (92.2%), bromophenol blue (90.2%), and malachite green (89.4%) compared to the control. However, it is well known that heavy metals inhibit several enzymes from the primary and secondary metabolic pathways. However, the percentage of color reduction was lower in the presence of Ni and Co in the growth medium. The decolorization activity of A. flavus was enhanced by Ca, Ni, or Co, but inhibited by Cu, while, the decolorization activity of P. canescens was significantly decreased by these metals, especially Cu [50]. Singh and Dwivedi [65] found that A. terreus GS28 isolate showed the highest decolorization direct blue-1 of 94.67% with manganese, 85.56%-83.88% with copper and zinc, and the lowest decolorization of 70.26% with iron. Manganese induced the activity of manganese peroxidase in isolate A. terreus, which enhanced the degradation of direct blue-1 dye. Similarly, copper enhanced laccase activity, which also contributed to the degradation of the direct blue-1 dye [65].

Fig. 5
figure 5

Effect of different heavy metals (100 ppm) on decolonization of azo dyes by A. flavus. Data represent mean values of three replicates ± show standard error (SE)

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Effect of static and shaking incubation on dye decolorization

The dyes (congo red, crystal violet, bromophenol blue, and malachite green) were decolorized by A. flavus during static and shaking incubations. There are notable variations in the decolorization of different azo dyes by A. flavus. However, the static culture generally performed better than the shaking condition in terms of both dye removal and fungal growth (Table 1). The treatment with congo red resulted in higher fresh weight (11.4 g), dry weight of fungal (2.3 g), and decolorization percentage (96.8%) under static culture. The decolorization of dye in stationary cultures was caused by the absorption of the dyes on the fungal biomass, which was not detected in agitated cultures. Bettin et al. [72] used laccases from Pleurotus sajorcaju PS-2001 to study the decolorization of 22 different colors. They found that under the same pH (3.2) and temperature (30–30 °C), the decolorization percentages were much lower during agitation (100 min−1) compared to static conditions.

Table 1 Growth of A. flavus and dye decolorization in static and shaking culture (fresh weight, dry weight, and dye decolorization)
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Effect of azo dyes on enzyme activities

The release of several oxidoreductases, such as laccase, peroxidase, and azoreductase, enables microorganisms to remove dyes using enzymes, as previously documented [73, 74]. Oxidoreductases efficiently break down dyes and produce more environmentally friendly products. The cell walls of fungi contain various components, including heteropolysaccharides such as chitin, chitosan, and glucan, as well as lipids and phospholipids [75, 76]. These components serve as binding sites for functional groups like carboxyl, hydroxyl, and phosphoryl groups, which facilitate the biosorption process [77]. Additionally, filamentous fungi are a powerful source of enzymes capable of converting a wide range of colors. Fungal enzymes that are known to contribute to the breakdown of dyes include laccases, lignin peroxidases, and manganese peroxidases [78].

The results in Table 2 show a significant increase in laccase and manganese peroxidase in the supernatant of A. flavus AUMC8653 supplemented with different azo dyes compared to the supernatant of A. flavus without azo dyes. The most pronounced increases of MnP activity were detected in the supernatant of A. flavus AUMC8653 supplemented with congo red (3.3μm/L), crystal violet (2.9μm/L), bromophenol blue (2.5μm/L), and malachite green (2.3μm/L), respectively compared to the control after 7 days. According to Ning et al. [61], similar findings demonstrated a 3.2-fold increase in MnP activity with the presence of Direct Blue 86. It was suggested that the dye may act as an activator, enhancing the production of important enzymes by A. flavus A5p1. Moreover, manganese peroxidase (MnP) is regarded as an essential enzyme that plays a significant role during the elimination process [61]. The decolorization process involves various oxidoreductase enzymes, which transform the original dye molecule into a less harmful substance [77]. MnP belongs to the lignolytic peroxidase family and can oxidize phenolic compounds by converting Mn2+ to Mn3+ and mineralize azo dyes through redox reactions [65]. The Mn3+ compounds are active oxidants, which are typically stabilized by chelating organic acids such as oxalic acid [65].

Table 2 Enzymatic and biochemical parameters under 50 mg/L of azo dyes by A. flavus
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Our results showed that the higher concentrations of laccase enzyme were detected in the supernatant of A. flavus AUMC8653 supplemented with congo red (0.80μm/L), crystal violet (0.75μm/L), bromophenol blue (0.60μm/L), and malachite green (0.55 μm/L), respectively compared to the control after 7 days (Table 2). The process of dye degradation by fungi is straightforward, as it involves the oxidation of complex pollutants, such as azo dyes, into simpler compounds. This occurs due to the production of extracellular enzymes, including laccase [16]. Laccases are phenolic oxidases that contain copper ions, which contribute to the generation of non-toxic phenolic compounds during biodegradation. This process operates through a non-specific free radical mechanism. In the degradation of azo dyes, the primary steps include the breakdown of the N–N bond, along with deamination, oxidation, desulfonation, and dihydroxylation [16, 79].

Our results showed that the higher concentrations of lignin peroxidase (LiP) enzyme were detected in the supernatant of A. flavus AUMC8653 supplemented with congo red (4.2μm/L), crystal violet (3.3μm/L), bromophenol blue (3.0μm/L), and malachite green (2.9μm/L), respectively compared to the control after 7 days (Table 2). Lignin peroxidases decompose dyes by oxidizing the phenolic group attached to the azo bond-bearing carbon, which generates a radical group. This phenolic carbon is then attacked by water, leading to the formation of phenyldiazene. Subsequently, phenyldiazene can be oxidized through a one-electron process, resulting in the production of nitrogen gas (N2). LiP increased degradation capacity in the presence of H2O2 [76].

The supernatant of A. flavus AUMC8653 was supplemented with different azo dyes and showed a significant increase in hydrogen peroxide compared to the supernatant of A. flavus without azo dyes. In addition, the maximum H2O2 content (3.8nM/g FW) was observed at 50 mg/L of malachite green treated with A. flavus AUMC8653. The minimum H2O2 content (1.01nM/g FW) was observed at 50 mg/L of congo red treated with A. flavus AUMC8653. Comparable outcomes were stated by Singh and Dwivedi [65], who observed that the highest H2O2 content was seen at 100 mg/L of direct blue-1 dye-treated A. terreus fungal biomass, followed by 500 and 1000 mg/L of direct blue-1 dye. Manganese peroxidase is part of a similar family, and in this study, the slow release of H2O2 boosted the activity of manganese peroxidase. When there is a lack of nutrients, mycelia produce H2O2, which activates the lignolytic enzymes for complete substrate degradation [80]. Enzymatic activity has a role in the decolorization of congo red and is indirectly linked to the generation of H2O2 [24]. H2O2 has a special function in the radical-generating process that breaks down lignocellulose [65]. Although a significant amount of H2O2 is necessary for peroxidase activity as a co-factor, it can occasionally also impede enzymatic activities [74]. Some fungal strains, such as A. fumigatus and A. flavus, induce degradation and produce water as a byproduct. Similarly, peroxidases in fungi utilize hydrogen peroxide (H2O2) as an oxidizing agent [16].

The protein content of A. flavus decreased significantly in congo red (0.18 mg/g), crystal violet (0.16 mg/g), bromophenol blue (0.14 mg/g), and malachite green (0.13 mg/g) as compared to the control (Table 2). Similar results were obtained by Singh and Dwivedi [65] who reported that the protein content of A. terreus showed increasing trends with 100, 500, and 1000 mg/L of direct blue-1 dye, but was lower than the control. This could be due to the generation of more stress at increasing concentrations of direct blue-1 dye, while the control showed the maximum protein content without the stress of the dye.

FT-IR

FT-IR studies showed that this was mostly caused by laccase-mediated enzymatic breakdown, which produced less hazardous metabolites. The degradation process involved the conversion of toxic malachite green to benzaldehyde via Michler's ketone pathway [79]. Additionally, another study reported the biodegradation of malachite green using A. flavus, which successfully degraded malachite green completely within 8 days by synthesizing laccases and manganese peroxidase [81].

The functional groups and chemical bonds in A. flavus were analyzed before and after dye biodegradation using FT-IR spectroscopy. Based on Fig. 6a, the FT-IR spectrum of A. flavus before the azo dyes removal displays several major intense bands, around 3267, 2958, 2924, 2219, 1639, 1542, 1434, 1374, 1331, 1231, 1149, 1025, 926, 915, 436 and 426 cm−1. The peak at 3267 cm−1 indicated that there was a strong absorption band due to the symmetric stretching vibration of O–H and NH2 groups, while, the peak at 1542 cm−1 indicated the presence of –NH in A. flavus. The C–H asymmetric stretching peak is located at 2924 cm−1, the C = O stretching and N–H deformation peak is located at 1639 cm−1 (in the amide II area), the carboxyl group is located at 1542 cm−1, and the phosphate group is located at 1231 cm−1. The peaks at 1148 cm−1 correspond to groups containing phosphate and sulfide. The polysaccharide groups C–C, C = C, C–O–C, and C–O–P are represented by the peaks at 1025 cm−1.

Fig. 6
figure 6

Exemplary FT-IR band positions (cm-1) of A. flavus before and after biodegradation of azo dyes

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Referring to Table 3 and Fig. 6 b-e, following the biodegradation of azo dyes, certain peaks exhibited minimal variation; some peaks appeared, while other peaks disappeared. This behavior was explained as a result of the azo dyes aligning with an active functional group, which changed the vibration frequency. Furthermore, positively charged azo dyes can bind to the cell surface through the electrostatic force produced by negatively charged functional groups (such as hydroxyl, amino, carboxyl, phosphate, nitro, and halide groups).

Table 3 Exemplary FT-IR band positions (cm−1) of A. flavus before and after biodegradation of azo dyes
Full size table

Following dye biodegradation, the FT-IR spectra are shown in Fig. 6b. The FT-IR peaks shift as a result of interactions between congo red molecules and A. flavus active surface during adsorption. FT-IR spectrum shows multiple strong bands at around 3266, 2113, 1639, 1547, 1420, 1331, 1227, 1149, 1022, 928, and 424 cm−1. Along with the elimination of several bands around 2985, 2924, 1374, 915, and 436 cm−1, the FT-IR spectra also showed the emergence of additional bands around 2160, 1126, 1077, 971, and 474 cm−1. Similar results were published by Kaushik and Seth [66], who demonstrated that the degradation of congo red (CR) and malachite green (MG) was validated by comparing the FT-IR spectra of the control dye sample and the samples extracted following decolorization by A. flavus strain GKRS09. Dye degradation was indicated by the FT-IR study's decrease in peak spectra, which shifted between the treated and control dyes.

The FT-IR spectra following dye biodegradation are shown in Fig. 6c. The FT-IR peaks shift as a result of interactions between crystal violet molecules and A. flavus active surface upon adsorption. Following the elimination of the crystal violet, A. flavus FT-IR spectrum shows multiple prominent bands at approximately 3266, 2957, 2117, 1639, 1545, 1433, 1372, 1332, 1229, 1149, 1023, 927, and 418 cm−1. Additionally, the FT-IR spectra show the emergence of new bands at around 1076 and 470 cm−1 as well as the loss of existing bands at approximately 2924 and 436 cm−1.

FT-IR spectra following dye biodegradation were displayed in Fig. 6d. The interaction between bromophenol blue molecules and A. flavus active surface during adsorption causes the FT-IR peaks to shift. A. flavus FT-IR spectrum shows multiple prominent bands at approximately 3263, 1638, 1429, 1376, 1329, 1148, 448, and 418 cm−1 following the elimination of bromophenol blue. Additionally, the FT-IR spectra shows the emergence of new bands at approximately 2323 and 1009 cm−1 as well as the elimination of existing bands at around 2958, 2924, 2119, 1542, 1231, 1025, 926, and 915 cm−1.

The FT-IR spectra following dye biodegradation are shown in Fig. 6e. The shift in FT-IR peaks is caused by interactions between malachite green molecules and A. flavus active surface following adsorption. Following the elimination of malachite green, the FT-IR spectrum of A. flavus shows multiple strong bands around 3263, 2103, 1635, 1374, 1247, 1147, 1027, 928, and 413cm−1. The FT-IR spectrum also represents the appearance of new bands around 2167, 1077, 485, and 413 cm−1 and the disappearance of other bands around 2958, 2924, 1542, 1434, 1331, 915, and 436 cm−1. The metabolites produced when Lasiodiplodia sp. broke down malachite green dye were found to have fewer harmful components compared to the control group, according to FT-IR analysis of those metabolites. The analysis also revealed that malachite green dye is transformed into alkanes, carboxylic acids, and aromatic amines. This conversion could be a potential mechanism for Lasiodiplodia sp. degradation of malachite green dye [60]. The C = O and N–H stretch cause the formation of amines, amides, and carboxylic acids. The breakdown of MG into less hazardous metabolites is confirmed by the presence of dimer OH [82].

The emergence of new peaks and the disappearance of prominent IR peaks may indicate the mineralization of the parent compound (dye molecules) [80]. The C-N bond breaking and ammonium ion deformation could be indicated by the peaks disappearing [73]. Fungal cell walls in microorganisms contain a variety of charged functional groups, or active sites, including phosphate, hydroxyl, and carboxyl. Fungi release heteropolysaccharides, which are crucial to the adsorption process of indigo decolorization [83, 84]. Since A. flavus biomass showed increased peaks impacting the parent chemical of congo red, it proved to be more effective for the surface sorption of congo red compared to A. terreus biomass [24].

The ability of fungal mycelia to biosorb is closely linked to their surface area and the functional groups present on their surface [65]. The effectiveness of dye biosorption relies on the hetero-polysaccharide and lipid content of the cell wall. These components possess varied charged functional groups that establish a strong attractive interaction between azo dyes and the cell wall [65]. However, positively charged and negatively charged groups are important to attract both basic and acidic dyes through electrostatic attraction which is the basis of the biosorption mechanism of dye removal [65]. The biotransformation of azo dyes into several metabolites is indicated by the disappearance of the principal IR peak in the FT-IR spectra and the appearance of a new peak [85]. Azo dyes have positively charged nitrogen atoms that interact with negatively charged groups, such as carboxylates and phosphates on the cell wall. Chemical bonds, either covalent or hydrogen, can form a link between azo dyes and the cell wall [86].

Light microscopy

The findings are supported by the visual observation that the fungal cells have lost their original color. As a result, fungus mycelia are examined under a microscope to determine the intracellular fungal biomass, as depicted in Fig. 7. The findings show that A. flavus may accumulate the dyes within the fungal biomass without forming biotransformations, and this process would be the primary mechanism responsible for decolorization. Recent research discovered that A. flavus can accumulate dyes in its biomass without altering them, which is the primary mechanism for color removal. The dyes are only adsorbed on the surface of live cells, and fungal enzymes break down the dyes to create new chemicals. When cell pellets are brightly colored, it indicates dye adsorption, while the original color of the dye is preserved when biodegradation occurs [87].

Fig. 7
figure 7

Uptake the dyes inside the fungal biomass of A. flavus after 7 days of incubation with different azo dyes

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Phytotoxicity test

The untreated wastewater from dye factories is found in water bodies that are used for irrigation. This poses serious environmental concerns due to its impact on the health of plants and ecosystems. Water used for irrigation directly and indirectly affects soil fertility, when the wastewater biodegrades, it produces numerous degradation products. It is important to study the toxic effects of these breakdown products on plants [59]. The data in Tables 4 and 5 illustrated the effects of biodegradable azo dyes before and after treatment with A. flavus AUMC8653 on seed germination and growth of maize and common bean plants. It was found that the percentage of germination, shoot length, root length, seedling length, fresh weight of seedlings, dry weight of seedlings, vigor index I, and vigor index II increased significantly in plants treated with fungal metabolites. The increase in the germination percentage of both tested plants could be attributed to the less toxic nature of the dye residues and produced metabolites [88]. In addition, all morphological parameters showed a significant reduction when the plants were irrigated with azo dyes compared to control plants. On the other hand, irrigation of plants with azo dyes after biodegradation significantly increased all morphological parameters of maize and common bean plants compared to azo dyes before biodegradation. The inhibition of germination can be attributed to reduced water intake by seeds in the presence of high concentrations of wastewater. This, in turn, affects energy-forming compounds, total solids, and heavy metals [59].

Table 4 Phytotoxicity test of degraded products of azo dyes to plant seeds of Z. mays after 21 days
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Table 5 Phytotoxicity test of degraded products of azo dyes to plant seeds of Phaseolus vulgaris after 21 days
Full size table

According to Singh and Dwivedi [65], the germination index of S. leucopersicum and T. aestivum seeds was approximately 36% and 40%, respectively, when exposed to Since A. flavus biomass showed increased peaks impacting the parent chemical of congo red, it proved to be more effective for the surface sorption of congo red compared to A. terreus biomass [24] concentration, however, 66 and 70% germination index was noted when exposed to the degradation product of direct blue-1 dye. As a result, treating A. terreus decreased the toxicity of direct blue-1 dye degradation products. As a result, A. terreus was employed to decolorize the direct blue-1 dye, and this study demonstrated that their metabolites were less hazardous to plant growth [88].

Garden cress (Lepidium sativum) seedlings were irrigated with water treated with Phanerochaete chrysosporium to test for toxicity. Compared to untreated plants, the fungi-treated plants exhibited a significantly higher germination rate [89]. In another investigation, Phaseolus mungo (black gram) and Sorghum vulgare (jowar) seeds were irrigated with water treated by Marasmius sp. after they had been contaminated with Diazo Reactive dye. For S. vulgare, the maximum mean shoots and root lengths were observed [90]. Ali et al. [91] examined the phytotoxicity of A. niger and T. viride on Vigna radiata. They found that plants treated with water remediated with A. niger exhibited the highest rate of germination. The study conducted by Singh and Dwivedi [24] revealed that the germination rate, root length, and shoot length of S. leucopersicum and T. aestivum were higher in the control treatment and fungal-treated congo red dye solution compared to the pure dye treatment.

Conclusions

The utilization of fungi that efficiently decolorize and degrade synthetic dyes with diverse chemical structures is essential to the efficacy of the microbial cycle in eliminating or degrading dyes from wastewater. A. flavus exhibited the greatest percentage of azo dye decolorization after eight days of incubation at 30° C, pH 6, sucrose as carbon source and NaNO3 as nitrogen source under static conditions. Laccase, manganese peroxidase, and lignin peroxides carried out the oxidative pathway-mediated biodegradation of azo dyes, resulting in a less hazardous byproduct. In comparison to control plants, the phytotoxicity assay results indicated that the degraded product was less harmful to the growth and germination rates of maize and common bean plants. As a result, A. flavus may be a useful microorganism for efficiently degrading azo dyes (Fig. 8). According to our findings, the bioremediation method is the best for reducing dye toxicity because it is inexpensive and safe for the environment.

Fig. 8
figure 8

Decolonization of azo dyes by A. flavus

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Data availability

All the data available in the manuscript.

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Open access funding provided by The Science, Technology & Innovation Funding Authority (STDF) in cooperation with The Egyptian Knowledge Bank (EKB).

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  1. Department of Biological and Geological Sciences, Faculty of Education, Ain Shams University, Cairo, 11341, Egypt

    Amira M. Ghanaim, Omima M. El Mahdy & Heba I. Mohamed

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  1. Amira M. Ghanaim
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  2. Omima M. El Mahdy
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Contributions

AMG, OME, and HIM: Methodology, Acquisition and analysis of data; AMG, OME, and HIM: Investigation, interpretation of data: AMG, OME, and HIM:: Validation, interpretation of data. AMG, OME, and HIM: Resources, Conception, design of the work. AMG, OME, and HIM: Project administration, Supervision, Conception, Design of the work, Writing—Original Draft. All authors read and approved the final manuscript. All authors have approved the submitted version.

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Correspondence to Heba I. Mohamed.

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Ghanaim, A.M., Mahdy, O.M.E. & Mohamed, H.I. Biodegradation of azo dyes by Aspergillus flavus and its bioremediation potential using seed germination efficiency. BMC Microbiol 25, 7 (2025). https://doi.org/10.1186/s12866-024-03703-9

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BMC Microbiology

ISSN: 1471-2180

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