A new Brevibacillus laterosporus L77 isolate capable of rapid decolorization and detoxification of textile azo dyesAbstract
Synthetic dyes are widely used in the textile industry and constitute one of the main environmental pollutants worldwide. In the present study, 60 bacterial isolates were analyzed for decolorization capabilities against reactive azo dye. In this screening, one has emerged as the most promising strain to degrade the selected dye, which was identified as Brevibacillus laterosporus. The effect of different physical-chemical and temporal parameters (dye concentration, temperature, pH, shaking, and kinetics) on its decolorizing potential was assessed. B. laterosporus showed decolorization efficiency higher than 90% for high concentrations of dye mixture. The best decolorizing potential was found in the range of 30-40°C and 8-9.5 pH. Agitation had negative effects on decolorization performance, with decrease in bacterial dye degradation. Decolorization higher than 90% was found in the first 12 hours of culture. UV-Vis analyses suggested the cleavage of azo bonds, since the peaks observed in untreated dye mixture reached close to zero after 12 hours of treatment. The toxicity test showed that the treatment with the bacterium was efficient in removing the toxicity of the dye mixture. These findings suggest that B. laterosporus L77 is a promising strain to be used in bioremediation of textile effluents containing azo dyes.
Key words
Bioremediation; eco-friendly; environmental biotechnology; sustainability
INTRODUCTION
Despite its economic importance, the textile industry is one of the main environmental polluters, generating tons of effluents contaminated by several toxic compounds, including dyes. It is estimated that 280,000 tons of synthetic dyes are discharged into the environment worldwide, causing many damages, not only aesthetically, but especially ecologically, compromising aquatic life and water quality (Sinha et al. 2024). Several impacts of textile dyes on human health are also recognized, ranging from neurological problems to mutagenic and carcinogenic risks (Kishor et al. 2021, Rathi & Kumar 2022, Das et al. 2023, Ramamurthy et al. 2024, Gambino et al. 2025).
Azo dyes comprise the main class of dyes used in the textile sector as they have many desirable characteristics, such as stability, chemical versatility, resistance to light, and high fixation, providing strong coloration to clothing (Alzain et al. 2023). However, this class of dyes is a global concern due to their polluting potential and the environmental risks of their inappropriate disposal (Pinheiro et al. 2022, Das et al. 2023).
Physical, chemical, and biological methods can be used for degradation and detoxification of azo dyes, in order to properly treat effluents before their disposal. Biological methods have been universally accepted and desirable, since they generate lower costs compared to conventional physical-chemical treatments, and are eco-friendly. In many situations, the combined action of physical and biological methods is highly efficient and cost-effective, making it a desirable option for the treatment of industrial effluents (Samsami et al. 2020, Suzuki et al. 2020, Das et al. 2023, Liu et al. 2023, Nachiyar et al. 2023, Kumaravel & Shanmugan 2024).
Eukaryotic and prokaryotic microorganisms can be used in biodecolorization processes. Despite the fact that eukaryotes have been more widely used, the use of bacteria has the advantages of a rapid growth cycle as well as ability to degrade and mineralize several dyes, which has attracted special attention to these organisms (Moyo et al. 2022, Khan et al. 2023).
The search for descolorant bacteria can be directed, taking advantage of the natural selection through bioprospecting in contaminated environments (Pinheiro et al. 2022). Several bacterial isolates used in biodecolorization at laboratory scale have been collected globally from areas affected by textile effluent disposal, which are potentially selective environments for microorganisms with the ability to survive and metabolize dyes (El Bouraie & El Din 2016, Khan & Malik 2018, Meerbergen et al. 2018, Pinheiro et al. 2022, Moyo et al. 2022, Kushvaha et al. 2022). However, the potential of bacteria originating from non-impacted sites has been poorly explored.
It is also important to emphasize that many physical and chemical conditions may affect bacterial degradation ability, and for an effective biodegradation, and real application, these factors need to be understood and optimized (Moyo et al. 2022). Therefore, the aims of this study were to isolate and identify a bacteria capable of decolorizing azo textile dyes, as well as to evaluate the influence of physicochemical and temporal parameters on its decolorization potential and to assess the toxicity of the dye degradation product.
MATERIALS AND METHODS
Bacterial prospection, isolation and storage
Sixty bacterial isolates collected from four different sources were used in this study: two sources corresponding to environments affected by polluting agents, i.e. watercourses receiving textile wastewater (n=19) and domestic grease traps (n=17), and two from non-polluted environments - rhizosphere (n=20) and decomposing wood (n=4).
Effluent isolates were collected from seven different points, all of which from water that receives effluents from textile industries and/or laundries, located in Apucarana, in the North Central region of Paraná-Brazil. Approximately 100 mL were sampled at each point, downstream of the textile effluent release site and down to 10 cm of water depth. Rhizosphere isolates derived from 17 different sampling points not subjected to direct anthropogenic action, located in the North of Paraná-Brazil. The soil at the collection sites is classified as red latosol and the rhizospheric soil comes from the root system of various plant species, including grasses, fruit trees and arboreal species. Approximately 50 grams of rhizosphere were collected at each point, and the samples were taken from a depth of 5 to 10 cm from the soil surface.
Four pieces of decaying wood (50 grams each) were also collected in North of Paraná-Brazil, in dense vegetation areas, predominantly with arboreal species. The samples were taken to the laboratory for bacterial isolation. Grease trap isolates were collected from three different domestic grease traps, by submerging the bottle and sampling approximately 15 mL from each point.
All samples were collected using sterile containers and immediately transported to laboratory for processing. For rhizosphere and decaying wood samples, 10 grams were dissolved in 9 mL of sterile distilled water and serial dilutions were plated on Tryptic Soy Agar (TSA). After incubation, colonies were selected, purified, and stored. Aliquots of 100 µL of grease trap samples were inoculated by spreading on TSA, followed by serial dilution.
Wastewater samples were filtered through 0.22 µm pore cellulose ester-membranes (Millipore); the membranes were vigorously shaken in 10 mL of saline solution to detach the cells, and inoculated by spreading on TSA, followed by isolation, purification, and storage of the colonies.
All the isolates are stored in Agar Stock Culture. Cells were activated by seeding a loop of the stock solution in 3 mL of TSB (18 hour / 35°C).
Azo dyes
The following textile azo dyes were used: Reactive Blue BF-5G 130% (RB), Reactive Red BF-4B 140% (RR), and Reactive Yellow BF-3R 250% (RY), supplied by Texpal®, Valinhos - São Paulo - Brazil. To prepare the stock solutions (1000 or 500 mg.L-1), the dyes were dissolved in distilled water and sterilized by filtration through 0.22 µm pore nylon membranes (Filtrilo). For the analyses, dyes were added to a sterile culture medium at the concentrations analyzed. The dye mixture consisted of adding each dye (RB 5G, RR 4B, and RY 3R) to the sterile culture medium at the concentration under study.
Bacterial biodecolorization performance
Bacterial isolates were tested for their ability to decolorize azo dyes. Isolates were grown statically (18 hour/ 35°C) in Tryptic Soy Broth (TSB) added by RB azo dye 50 mg.L-1 for initial screening. The cultures were centrifuged (11,180 x g/5 min) and the supernatants underwent spectrophotometric evaluation. Sedimented cells were washed in saline solution and suspended in sterile water to determine the bacterial growth of each culture (O.D. 600 nm). The same conditions used in the tests were used to establish the decolorization controls, except for the absence of bacterial inoculum.
In order to test for possible enzymatic activity in the culture supernatant the isolate that showed the highest decolorization ability was grown in TSB broth either with or without dyes for 18 hour/35°C. Afterwards, the cultures were filtered through 0.22 µm nylon membranes (Filtrilo) to remove the cells, and the resulting supernatants were added to the azo dyes solution at a concentration of 50 mg.L-1. The samples were incubated for 18 hours and decolorization was assessed spectrophotometrically.
Morphological characterization and molecular identification
The isolate with the highest decolorization ability was characterized according to morphological features and molecular identification from 16S rDNA gene sequencing. Genomic DNA was extracted using the Pure Link® Genomic DNA kit (Invitrogen, USA) and part of the 16S rDNA gene was amplified by polymerase chain reaction (PCR) using the universal primers fD1 5’-AGAGTTTGATCCTGGCTCAG-3’ and rP1 5’-ACGGTTACCTTGTTACGACTT-3’ (Weisburg et al., 1991). PCR reactions were performed in a final volume of 15 μL, including: GoTaq® Green Master Mix (Promega Corp., USA), 0.65 µM of each primer, and 15 ng of DNA. Amplification was performed in a thermal cycler under the following conditions: initial denaturation of 95 °C for 5 minutes, followed by 35 cycles of 95 °C for 45 seconds, 50 °C for 30 seconds, and 72 °C for 1 minute, and a final elongation of 72 °C for 10 minutes. PCR products were purified using EXOSap IT® (Prodimol) and prepared for bidirectional Sanger sequencing using a BigDye Terminator v 3.1 kit (Applied Biosystems, USA), according to the manufacturer’s instructions. Sequence reading was performed using an ABI-PRISM 3500 XL automatic sequencer (Applied Biosystems, USA) and the new sequence was deposited in GenBank (NCBI - National Center for Biotechnology Information) under accession number PP263612.
The partial 16S rDNA sequence of the L77 isolate was subjected to BLAST (Basic Local Alignment Search Tool - https://blast.ncbi.nlm.nih.gov/Blast.cgi) to find the most similar sequences in the GenBank database, and to obtain data from the closest individuals and species that were included in the phylogenetic analysis. The set of sequences was edited and aligned using MEGA v.11.0, which was also used to reconstruct phylogenetic relationships by maximum parsimony method, based on MP trees inferred from 1000 repetitions, with a subtree pruning and regrafting (SPR) algorithm at research level 1, in which the initial trees were obtained by adding sequences at random (10 replicates).
Scanning Electron Microscopy
A 24-hour bacterial culture was fixed in a 3% glutaraldehyde solution in 0.2 M phosphate buffer pH 7.2 for 24 hours. The material was washed with phosphate buffer 3 times, frozen in an ultra-freezer (-80 °C) and subjected to freeze-drying. Subsequently, the samples were mounted on carbon tape in an aluminum stub, covered with a 25 nm layer of gold (Sputter Coater SDC 050 - BALTEC), observed in a scanning electron microscope operating in vacuum under an electron beam with 10 kV acceleration (FEI Quanta 200).
Optimization of L77 decolorization parameters and kinetics
The effect of different physico-chemical parameters on the decolorization of azo dyes by actively growing cells of Brevibacillus laterosporus L77 (GenBank accession number: PP263612) was analyzed. The following were evaluated: decolorization performance at higher dye concentrations (100, 200, 300, 400, 500, 600, and 800 mg.L-1), under different temperatures (15–45oC), cultures at different initial pH values (4,5; 5,0; 5,5; 7,5; 8,0; 9,6 and 10), rotation speed (0,0; 0,56 and 5,04 x g), and decolorization kinetics (from zero to 24 hours).
Spectroscopy and UV-Vis
To determine the percentage of decolorization under the culture conditions evaluated in this study, culture supernatants were read spectrophotometrically (Thermo Fisher Scientific - Genesis 150 UV-Vis) at different wavelengths according to the dye used (599 nm for RB, 542 nm for RR, 404 nm for RY, and 550 nm for the mixture of azo dyes). The following equation was applied to calculate the decolorization percentage %DES = [(initial ABS - final ABS)x100]/Initial ABS, where %DES represents decolorization efficiency, initial ABS represents the absorbance value of the control solution, and final ABS represents the absorbance value after cultivation (treatment) (Maniyam et al. 2020). UV-Vis spectral analyses were carried out in the range of λ 290 to λ 1000, with readings every 2 nm.
Phytotoxicity test
To evaluate the phytotoxicity of degradation intermediates from the dye mixture, a test was carried out on Cucumis sativus seeds (Top Seeds), based on the study by Pellizzer et al. (2024). B. laterosporus L77 was grown statically (35°C) for 3 days in TSB added by the dye mixture (50 mg.L1 of each dye), and after this period, the culture was subjected to agitation (5.04 x g; 35°C) for another 3 days. The culture supernatant was obtained by filtration (Filtrilo; NY 0.22 µm) and was diluted (25%) in sterile distilled water. Ten C. sativus seeds were placed in Petri dishes (8 cm diameter) on filter paper and moistened with 10 mL of each sample.
The plates were incubated under a completely randomized experimental design with three replicates and maintained in a climate chamber (Tecnal TE-4001-E7) with a 12hour/12hour dark/light photoperiod at 25°C and 50% humidity for five days of incubation. The culture medium added by the dye mixture were used as control, at the same concentration and dilution used in the test. Distilled water was used as seed growth control. Toxicity effects were recorded in terms of germination rate (G%) and seed vigor index (VI%), according to Abdul-Baki & Anderson (1973) and Jain et al. (2014), and were calculated using the following formulas:
Putative brAzo gene amplification and sequencing
The putative brAzo gene was amplified by PCR using genomic DNA as a template and the following primers: forward 50 -GATTACACATATGGCAAAAG TATTG-30 and reverse 50 -GTTTATTTT TCTCGAGCAAACGTTTTTGCG-30, according to Lang et al. (2013). The PCR and sequencing procedure were carried out as described for 16S rDNA gene.
Statistical analysis
A one-way ANOVA was performed to test significant differences (p < 0.05) between the source of the isolates and discoloration, as well as between incubation time and discoloration. A post hoc Tukey’s test was conducted to determine which groups differed significantly (p< 0.05). The assumptions of normality and homoscedasticity were tested, and when necessary, data were log-transformed to meet these assumptions. When the assumptions of ANOVA were not met, as the response variables did not show a normal distribution (p> 0.05 by the Shapiro-Wilk test), variations in decolorization as a function of dye concentration were analyzed using the Kruskal-Wallis test, followed by post hoc tests Wilcoxon-Mann-Whitneytest. All analyses were carried out in the R environment. Significance tests were calculated using the aov function, TukeyHSD, kruskal.test, and pairwise.wilcox.test. Graphs were constructed using the ggplot2 package.
RESULTS
The screening of bacterial decolorization potential showed that all 60 isolates evaluated had partial potential to decolorize RB azo dye in 18-hour cultures, ranging from 17.8 to 96.3%. Individual decolorization results for each isolate are shown in Table I. Analysis by isolation source showed that, as a whole, wastewater isolates promoted higher decolorization than rhizosphere isolates (p<0.05). There were no significant differences between the other sources (Figure 1).
In our study, out of 60 bacterial isolates, L77 emerged as the most effective in terms of biodegradation potential of the tested azo dyes. This isolate was obtained from rhizosphere of Musa paradisiaca, located at latitude -23.174637 and longitude -50.670606 and showed complex colony structuring in 10-day cultures. Micromorphologically, it appears as a spore-forming Gram-positive diplobacillus (Figure 2). Analysis of the partial 16S rRNA sequence identified L77 isolate as Brevibacillus laterosporus (GenBank accession number: PP263612), showing similarities of over 99.8% with other B. laterosporus, according to BLAST analysis at NCBI. In the phylogenetic analysis, the topology exhibited by the Maximum Parsimony tree revealed a well-supported monophyletic group formed by L77 together with other B. laterosporus, as shown in Figure 2. Regarding the possible brAzo gene, a 570 bp segment was amplified and sequenced, showing over 99% similarity with FMN-dependent NADH Azoreductase gene from other B. laterosporus, based on BLAST analysis (GenBank accession number: PV173308).
Morphological and molecular characterization of the L77 isolate. a) Colony morphology in 10-day cultures; b) Scanning electron microscopy of decolorizing cells. Bars correspond to 10 µm; c) Phylogenetic relationships of Brevibacillus laterosporus L77 according to Maximum Parsimony analysis of partial 16S rRNA gene sequence.
In 18-hour cultures in the presence of 50 mg.L-1 azo dyes, B. laterosporus L77 showed a mean decolorization efficiency of 96.3%, 90%, and 75.8% for RB, RR, and RY dyes, respectively, and 95.9% for the mixture of the three azo dyes. No coloration was detected in the cell sediment, which remained white regardless of the dye applied (Figure 3). Cell-free supernatants of L77 cultures failed to promote decolorization of azo dyes, under the evaluated conditions.
Decolorization scale of azo dyes by 18-hour cultures of B. laterosporus L77. n RB; b) RR; c) RY; and d) Mixture; Bars represent decolorization (%) and lines represent O.D. 600 nm. e) visual decolorization (18-hour culture) of the dye mixture (50 mg.L-1 of each dye: RB, RR, and RY); the microtube on the left represents the control and on the right it represents the test situation. The arrow shows cells sediment after centrifugation. Different lowercase letters represent statistical differences (p<0,05) in decolorization. Results from 3 individual experiments.
Considering the high decolorization performance of B. laterosporus L77 with azo dyes at 50 mg.L-1, its decolorization profile at higher concentrations (100 to 800 mg.L-1) was also evaluated. Under these conditions, the best performance was observed for the RR dye, where there was no significant difference in decolorization up to a concentration of
500 mg.L-1, compared to that promoted at
50 mg.L-1 (Figure 3). The bacterial decolorization performance decreased from 600 mg.L-1 onwards (p<0.05). Poor cell growth (O.D 600 nm = 0.166) was detected at the highest concentration evaluated (800 mg.L-1), as well as a marked decrease in decolorization potential (6.3%). For the RB dye, the percentage of decolorization remained unchanged up to a concentration of 400 mg.L-1, compared to that found at 50 mg.L-1. At 500 mg.L-1, B. laterosporus L77 decolorization potential was lower than at the previous concentrations (p<0.05). The isolate was unable to decolorize higher concentrations of the dye, although cell growth was observed (Figure 3). Decolorization of the RY dye remained above 70% up to a concentration of
400 mg.L-1, with no differences in decolorization performance between 50 mg.L-1 and
400 mg.L-1. At higher concentrations, B. laterosporus L77 decolorization performance decreased (p<0.05) (Figure 3). In the mixture of azo dyes, the mean percentage of decolorization promoted by B. laterosporus L77 remained above 95% up to 200 mg.L-1. At 300 mg.L-1 a decrease in decolorization potential (p<0.05) was detected, and there was no decolorization at concentrations above 400 mg.L-1, despite bacterial growth (Figure 3).
Decolorization efficiency and growth of B. laterosporus L77 was significantly affected by temperature variation (Figure 4). However, decolorization performance remained high in the 30-40°C temperature range. For 18-hour cultures in the presence of RB, RR, and dye mixture, B. laterosporus L77 maintained decolorization over 90% in this temperature range, with a drastic reduction at either lower or higher temperatures (p<0.05). For cultures in the presence of RY, maximum decolorization was achieved at 35°C (p<0.05).
Effect of physical chemical parameters on decolorization by B. laterosporus. a-d) Effect of temperature on the decolorization of the dyes a) RB, b) RR, c) RY, and d) dye mixture; e-h) Effect of pH on decolorization of the dyes e) RB, f) RR, g) RY, and h) dye mixture; i) Effect of agitation on decolorization of the dye mixture. Bars indicate decolorization (%) and lines show O.D. 600 nm. Different lowercase letters represent statistical differences (p<0,05) in decolorization. Results from 3 individual experiments.
B. laterosporus L77 maintained decolorization efficiency (50 mg/L; 35°C; 18-hour cultures) at a broad range of pH values (Figure 4), with optimum values between 7.5-10 for RB, 8-9.5 for RR and mixture, and 5-8 for RY. More acidic conditions (pH 4.5) radically affected decolorization capacity (p<0.05).
Shaking cultures (5,04 x g; 18 h; 35°C) of B. laterosporus L77 caused a significant decrease in decolorization compared to those detected under static conditions (p<0.05), as well as at a lower agitation intensity (0,56 x g), as shown in Figure 4. Interestingly, bacterial growth was more pronounced with increased agitation speed, contrary to the findings for decolorization (Figure 4).
The kinetics of dye decolorization were also evaluated over a 24-hour period. No significant difference in decolorization was found between 0 and 4 hours, neither was there any visual decolorization of the dyes (Figure 5). The start of the logarithmic phase was achieved between 4 and 6 hours of cultivation, and from 6 hours onwards decolorization was found to be higher (p<0.05) than that promoted by the 4-hour cultures; it was higher than 70% for RB and between 40 and 50% for the other dyes.
Decolorization of azo dyes over time by cultures of B. laterosporus L77. Letters a, b, c, and d represent decolorization of RB, RR, RY, and the mixture, respectively. In the graphs, bars indicate decolorization (%) and lines show O.D. 600 nm. Letters e and f show visual decolorization over time and UV-Vis spectral analysis, respectively, of the dye mixture (50 mg.L-1 of each dye: RB, RR, and RY). In f, the arrowhead indicates the control; the wide arrow indicates 6 hours; the thin arrow indicates times 12 and 24 hours. Different lowercase letters represent statistical differences (p<0,05) in decolorization. Results from 3 individual experiments.
In the presence of RB or RR, there was no significant difference in decolorization between 12- and 24-hour cultures, with decolorization of over 90% being detected in the first 12 hours of culture. For RY dye, decolorization rates between 70 and 80% were detected in the 12- and 24-hour period (p<0.05). When the mixture of azo dyes was analyzed, significant differences in decolorization were observed at all cultivation times evaluated after the first 6 hours. In this situation, the average decolorization in 24-hour cultures was higher than 96%.
To detect the possible occurrence of biodegradation, control and treated dye samples were analyzed using UV-Vis spectroscopy, and changes in peaks were recorded. The analysis showed that the peak detected in the first 4 hours of cultivation decreases over time, reaching close to zero after 12 hours for all dyes evaluated, as well as for the mixture (Figure 5).
Finally, experiments were performed to assess the toxicity of degradation products of dye mixtures by B. laterosporus L77. Germination of C. sativus seeds showed that there were no statistical differences between control (water) and the supernatant of the bacterium dye treatment. Similarly, the vigor index of C. sativus seeds was examined for growth of seed germination and showed a non-significant decrease in bacterium treatment compared to control. Seed germination and seedling development are shown in Figure 6.
Toxicity effect of azo dye mixture degradation product by B. laterosporus L77 on C. sativus. a) Growth control (distilled water); b) Azo dye mixture (50 mg.L1 of each dye: RB, RR, and RY in culture medium); c) Azo dye mixture treated by L77; d) C. sativus seedlings grown in water (growth control); e) C. sativus seedlings grown in the presence of dye mixture treated by L77.
DISCUSSION
Azo dyes comprise the main class of dyes present in textile effluents, and their relevance and importance are likely to increase in the future. However, the structural properties of these dyes make them difficult to degrade under natural conditions and even through some conventional treatments. Consequently, the development of new cost-effective and sustainable technologies for the treatment of these industrial effluents has been encouraged worldwide. Microbial degradation has emerged as one of the main methods for detoxifying azo dyes, and some characteristics of bacteria, such as their rapid proliferation and enzymatic diversity, render them potentially useful for wastewater treatment (Benkhaya et al. 2020, Moyo et al. 2022, Rathi & Kumar 2022). This study assessed the decolorizing potential of bacterial isolates prospected in environments with different characteristics, ranging from pollutant-impacted environments, such as stream water containing textile effluents, to environments with low anthropogenic action. Microbial prospecting in environments impacted by textile effluents can provide higher success in biodecolorization, given the natural selection promoted by environmental pressure exercised by the presence of high concentrations of dyes in these effluents (El Bouraie & El Din 2016, Khan & Malik 2018, Meerbergen 2018, Pinheiro 2022, Moyo 2022, Kushvaha 2022, Harish 2024). Taken together, the isolates collected from effluents analyzed in this study showed higher decolorization rates compared to those from the rhizosphere. However, only one of the 60 isolates evaluated, collected from the rhizosphere, stood out as an excellent biodecolorizer. This highlights the importance of prospecting bacteria in uncontaminated environments, which can also be promising sources for bioremediating microorganisms.
This isolate was molecularly identified as B. laterosporus and this species is often known for its potential as a biological pest control agent (Bedini et al. 2020, Barbieri et al. 2021, Li et al. 2021, Hamze & Ruiu 2022, Smirnova et al. 2023, Su et al. 2024), as well as other bioactivities, such as plant growth promotion (Wang et al. 2022, Swiatczak et al. 2023), antibiosis (Zayed et al. 2022, Abdulsahib & Rauf 2023) and anticancer activity (Chen et al. 2022, Zayed et al. 2022). Few studies have reported biodecolorizing activity for isolates of this species (Gomare & Govindwar 2009, Kurade et al. 2013, 2016). The present data emphasize the potential of B. laterosporus in bioremediation, particularly in the biodecolorization of azo dyes.
To date, most studies concerning bacterial biodecolorization have focused on the degradation potential of one or more dyes, separately (Gomare & Govindwar 2009, Fareed et al. 2022, Ikram et al. 2022, Khandare et al. 2023, Sma-Air & Richie 2025). In this study, a new isolate of B. laterosporus is described that is capable of efficiently decolorizing a mixture of azo dyes, aside from single dyes, which is an important differential for real biotechnological potential, since industrial textile effluent is composed of a combination of dyes (Moyo et al. 2022).
Bacteria can decolorize dyes by adsorption into biomass or enzymatic biodegradation. In adsorption there is no breakdown of the dye, which is retained in the cells, while in biodegradation the chemical structure of the dye is broken down, or possibly completely destroyed, which is the most desirable route. Many bacteria are recognized to use biological enzymes- such as azoreductase, laccase, and peroxidase- to catalyze the breaking of azo bonds (Khan et al. 2021, Cong et al. 2022, Rathi & Kumar 2022). B. laterosporus L77 cells remained colorless after decolorization, suggesting that the process occurred through enzymatic degradation of the dye. The enzymatic decolorization by B. laterosporus has already been reported, and taking into account the pattern of decolorization observed, the participation of several enzymes at appropriate concentrations was suggested (Gomare & Govindwar 2009, Kurade et al. 2013).
The present study failed to show decolorization by cell-free supernatants of B. laterosporus L77. Although dyes can be eliminated by using purified enzymes, there are few studies on the performance of cell-free supernatants, and this application in wastewater treatment has been limited so far, since even the use of pure enzyme alone occasionally has no evident impact on the degradation of organic pollutants (Singh & Arya 2019).
We analyzed the influence of physical and chemical factors on the decolorization potential of B. laterosposus L77. Dye concentration is known to have an impact on decolorization, as decolorization rates decrease gradually with increased dye concentration. This occurs probably due to blockage of enzyme sites, toxic effects on bacteria, causing their death, and/or insufficient biomass concentration, leading to improper cell-to-dye ratio (Kumaravel & Shanmugan 2024). In this study, B. laterosporus L77 showed ability to decolorize high concentrations of individual dyes. Regarding the dye mixture, high decolorization efficiency was observed at concentrations up to 200 mg.L-1 of each dye. Higher concentrations probably inhibited bacterial enzymatic activity, since growth was observed at concentrations higher than 500 mg.L-1, although there was no decolorization at this concentrations.
Temperature variations generally have an impact on bacterial growth, survival, and metabolic activity, affecting virtually every process of microbial life. Our results show that B. laterosporus L77 is sensitive to wide temperature variations, in terms of both growth and decolorization activity, which may occur since temperatures below or above the optimum value can lead to loss of cell viability and inactivation of metabolic pathways, affecting biodecolorization capabilities, among other factors (Moyo et al. 2022, Kumaravel & Shanmugam 2024). However, the broad range of temperatures in which optimum decolorization capacity was maintained renders B. laterosporus L77 more suitable for dye bioremediation.
We found that the best decolorization performance of B. laterosporus L77 occurred under basic to alkaline conditions, particularly in the pH 7.5-9.5 range. Sodium hydroxide and other aqueous alkaline solution are used in the textile dyeing process, which can keep the effluent at a basic pH (Khattab et al. 2020, Pinheiro et al. 2022). Thus, the maintenance of optimum decolorization capacity in alkaline conditions is a favorable attribute of B. laterosporus L77.
Our results also showed that the decolorization performance of B. laterosporus L77 is better under static conditions, although the isolate decolorized about 70% of the dye mixture even with vigorous agitation (5,04 x g). This study identified a putative brAzo gene in B. laterosporus L77, as described by Lang et al. (2013). The enzyme encoded by this gene require NADH as an electron donor to catalyses the reductive cleavage of azo bonds. Under shaking conditions there is an increase in the amount of dissolved oxygen, as well as greater microbial growth, and the intense bacterial respiratory activity can impair azo bond reduction by competition for NADH, needed as electron acceptor for the azo bonds, which is also used by aerobic respiration (Kalme et al. 2007, Pinheiro et al. 2022, Moyo et al. 2022, Kumaravel & Shanmugan 2024). Different authors have reported improved performance in bacterial azo dye decolorization under static conditions compared to agitation (Khan & Malic 2016, Ajaz et al. 2019, Thapa et al. 2021). However, other papers have reported superior decolorization under agitation (Saranraj et al. 2018), showing that different bacteria respond differently to similar culture conditions, probably due to their diversity in degradation pathways. It should be emphasized that the ability to promote decolorization without needing to shake is a favourable characteristic of the isolate, as this makes it less expensive to industrially employ.
Rapid bacterial growth and production of a diversity of enzymes are factors that highlight the biotechnological interest and are desirable in biodecolorization since they can provide rapid degradation of the dye. In this study, B. laterosporus L77 showed biodecolorization rates of over 90% after 12 hours of incubation, suggesting that the supposed enzymatic expression and activity occur during the log phase of growth. Decolorization rates exhibited by B. laterosporus L77 in 12 hour-cultures are comparable to those exhibited by other bacterial species at times higher than 90 hours, highlighting its decoloring potential (Parshetti et al. 2010, Franciscon et al. 2012, Wang 2013, Khan & Malik 2016, Maniyam et al. 2020, Guembri et al. 2021, Srivastava et al. 2022, Ikram et al. 2022, Haque et al. 2024).
Our Uv-Vis results suggested that during the decolorization reaction, the azo bond was cleaved, most likely mediated by azoreductase, similar to the findings by Oturkar et al. (2011), Qu et al. (2012), and Khan & Malik (2016). In addition, our toxicity test indicated that the treatment with the bacteria was efficient in removing the toxicity of the dye mixture, which reinforces the suitability of B. laterosporus L77 as a potential, sustainable, and cost-effective decolorization agent that should be studied at a larger scale.
CONCLUSIONS
B. laterosporus L77 is a promising isolate for the eco-friendly treatment of textile effluents, remarkable for its effectiveness, speed, ability to degrade a mixture of azo dyes at high concentrations, and for its detoxification of a dye mixture.
Acknowledgements
We would like to thank the Department of Science, Technology and Higher Education (SETI); the Araucária Foundation to Support the Scientific and Technological Development of the State of Paraná (FA); the Multiuser Laboratory/Universidade Estadual do Norte do Paraná, Campus Cornélio Procópio and the Laboratory of Materials and Molecules Analysis/Universidade Estadual de Londrina.
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