Heterogeneous biocatalytic system for effective decolorization of textile dye effluent

Abstract

The current physicochemical methods for decolorizing toxic synthetic dyes are not sustainable to halt the environmental damage as they are expensive and often produce concentrated sludge, which may lead to secondary disposal problems. Biocatalysis (microbes and/or their enzymes) is a cost-effective, versatile, energy-saving and clean alternative. The most common enzymes involved in dye degradation are laccases, azoreductases and peroxidases. Toxic dyes could be converted into less harmful byproducts through the combined action of many enzymes or the utilization of whole cells. The action of whole cells to treat dye effluents is either by biosorption or degradation (aerobic or anaerobic). Using immobilized cells or enzymes will offer advantages such as superior stability, persistence against harsh environmental conditions, reusability and longer half-lives. This review envisages the recent strategies of immobilization and bioreactor considerations with the immobilized system as the effective treatment of textile dye effluents. Packed bed reactors are the most popular heterogeneous biocatalytic reactors for dye decolorization due to their efficiency and cost-effectiveness.

Introduction

Textile effluents are the prime source of water pollution. The chemicals in effluent include unutilized synthetic dyes, acids and alkalis, heavy metals, trace elements, etc. The textile industrial effluents predominantly consist of synthetic dyes. Synthetic dyes are produced from carcinogenic and highly toxic elements like mercury, lead, sulphur and aromatic compounds. The mordants such as chromium and iron used for fixation increase the toxicity of natural and synthetic dyes. World Health Organization reported that textile industrial effluent contributes around 20% of industrial water pollution. Approximately 2,00,000 tonnes of various dyes are discharged along with effluent per annum (Ogugbue and Sawidis 2011). Dyes are categorized based on their source, structure, ionic charge and applications (Fig. 1).

Fig. 1.

Classification of dyes

Apart from the intense colour imparted by the dyes, the effluent has extreme pH, a significant amount of suspended solids, chemical oxygen demand (COD) and biological oxygen demand (BOD) (Yaseen and Scholz 2017); high chemical and photolytic stability; and persistence in the natural environment (Guadie et al. 2017). Discharging hazardous effluents into different water bodies may resist light penetration and oxygen solubility, affecting aquatic flora and fauna. The impact of dye effluents on the living system is based on their origin, chemical structure, mode of application, and concentration, which results in various health hazards, including carcinogenesis (Mahida and Patel 2016). Hence it is inevitable to effectively treat the effluent prior to release into the environment.

So far, many physical (filtration, flocculation, sedimentation, adsorption, and reverse osmosis), chemical (Fenton’s process, oxidation, ozonation and advanced oxidation processes such as photocatalysis and sonocatalysis) and biological methods (microorganisms and their enzymes) have been employed for the efficient treatment of dye effluents (Emam and Ahmed 2019). However, physical methods simply transfer recalcitrant dyes from one medium to another resulting in secondary wastes. Whereas, in chemical methods such as Fenton’s process, chemical oxidation and ozonation result in toxic, unstable metabolites imparting adverse effects on animal and human health. In the advanced oxidation process (AOPs), highly reactive hydroxyl radicals (OH*) are produced to non-selectively oxidize many recalcitrant compounds, including dyes (Ajmal et al. 2014). These radicals are produced with the help of one or more primary oxidants (e.g. ozone, hydrogen peroxide, oxygen) and/or energy sources (eg- ultraviolet light, visible light, sonic waves) or catalysts (e.g.titaniumdioxide, ZnO, Fe₂O₃) (Emam et al. 2020, 2019). However, the operating cost, the flexibility of operation and effectiveness over a large area are the key challenges (Emam et al. 2020). At the same time, the biological treatment of textile dye effluents is eco-friendly, cost-effective, easy to handle, versatile and clean technology for the decolorization of textile dye effluents (Fig. 2).

Fig. 2.

Biological route of dye decolorization

Biological treatment is catalyzed by the systematic action of various enzymes produced by different organisms. Enzymes such as laccase, azoreductases and peroxidases were reported to decolorize textile dye effluents significantly. The combined action of various enzymes can be ensured by either immobilizing multiple enzymes or whole cells.

Degradation of textile dyes employing bacterial, fungal and algal species has become a significant area of research as it can degrade toxic wastes into less-toxic or non-toxic. Microbial degradation can convert toxic dyes into carbon dioxide, biomass and other inorganic compounds (Kolekar et al. 2012). Compared to bacterial cells, algal and fungal biomass could be used as effective biosorbents. Immobilized enzymes/cells are superior in decolorization due to better stability under various conditions. The present review focuses on efficiently using heterogeneous biocatalytic systems to handle toxic textile dye effluents effectively.

Enzymes for dye degradation

The enzymatic approach for the biodegradation of dyes is considered an efficient molecular tool for treating phenolic pollutants in dye effluents. The biological route of dye degradation is by the action of several oxidoreductase enzymes such as laccase, peroxidases and azoreductases (Fig. 3). Microbes are the largest and most economical source of these enzymes as they can be easily handled and purified (Sarkar et al. 2017). Azoreductases are highly specific in cleaving the azo bonds. While phenoloxidases, such as peroxidases and laccases, act non-specifically on aromatic rings (Abadulla et al. 2000).

Fig. 3.

Enzymes involved in dye decolorization: (a) laccase, (b) azoreductase and (c) peroxidases

Laccase for dye degradation

Laccases (EC 1.10.3.2; benzenediol: oxygen oxidoreductases) belong to the multicopper oxidases (MCOs) superfamily, a group of enzymes consisting of cupredoxin like domain which enables it to reduce O₂ to water without releasing harmful reactive oxygen species. Laccase is a key enzyme used in industries to treat dye effluents. It facilitates the oxidation of organic dyes in the effluent, thereby removing the toxic compounds. The dye degradation ability of laccases can be improved by adding specific mediators such as 1-hydroxybenzotriazole (HBT) (Senthivelan et al. 2016).

Laccases are present in bacteria, fungi, plants and animals. Plant laccases participate in lignin biosynthesis, whereas in insects, laccase aids the sclerotization of epidermal cuticular proteins. Microbial laccases participate in morphogenesis and copper homeostasis (Giardina et al. 2010). Microbial laccases used in dye decolorization are listed in Table 1.

Table 1.

Laccases from the microbial origin for dye decolorization

Source	Dye decolorized (% removal)	Conditions	Activity	Km	Vmax	References
Enterobacter sp. AI1	Malachite green (78%) Congo red (85.7%) Methyl orange (77%) Remazol Brilliant Blue R (81%) Reactive blue 4 (73%)	60 ºC, pH- 4.0	187.88 U/mg (Specific)	0.23 mM	51.28 μM/min	Edoamodu and Nwodo (2021)
Bacillus pumilus ZB1	Congo red (16.43%) Crystal violet (54.05%) Reactive blue 4 (41.61%)	80 ºC, pH- 5.0	10.18 U/L	35.454 µM		Liu et al. (2021)
Arthrographis kalrae	Remazol Brilliant Blue R (97.18%)	35 ºC, pH- 7.0, Time- 2 h	1525 IU /mL	306.3 µM	3.492 µM/ min	Yadav et al. (2021)
Oudemansiella canarii	Congo red (80%)	30 ºC, pH- 5.5, Time- 24 h	-	46.180 ± 6.245 µM	1.840 ± 0.101 µmol/min	Iark et al. (2019)
Escherichia coli CD-2	Indigo dye (90%) Amaranth (50%) Congo red (50%)	60 ºC; pH- 8.0; Time- 1 h	512.81 U/mg (Specific)	0.737 μM	100.5 U/mg	Javadzadeh and Asoodeh (2020)
Trametes versicolor TV-6	Aniline purple color (91%)	55 ºC; pH- 4.0; Time- 72 h	54.59 U/mg (Specific)	–	–	Tišma et al. (2020)
Aspergillus sp. Omeje	Basic violet 2 (36 ± 11.31%) Azo yellow 6 (41 ± 8.48%) Acid red 337 (39 ± 4.24%) Basic blue 22 (58 ± 2.83%) Azo purple (73 ± 22.63%) Azo yellow (65 ± 8.49%) Azo Brilliant Black (62 ± 5.65%) Vat blue 5 (82 ± 12.73%) Acid blue 74 (58 ± 5.66%)	37 ºC; pH- 5.0; Time- 2 h	336 ± 11.31 U/L	0.14 mM	0.45 μmol/min	Omeje et al. (2020)
Trametes versicolor	Malachite Green (95%)	25 ºC; pH- 4.5; Time 2 h	24.8 U/g	–	–	Xu et al. (2020)
Fusarium oxysporum HUIB02	Bromothymol blue (99.68%), Methyl orange (99.56%), Remazol brilliant blue R (98.66%), Indigo carmine (97.75%), Malachite green (75.47%) Evans blue (57.64%)	45 ºC; pH- 5.0; Time- 15 days	186 U/mL	–	–	Huy et al. (2020)
Spirulina platensis CFTRI	Remazol Brilliant Blue R 89% with syringaldehyde; 63% with 1-hydroxybenzotriazole	30 ºC; pH- 6.0; Time- 48 h	66.74 U/mL	–	–	Afreen et al. (2017)

While most of the laccases from plant, fungi, insects and some bacteria contain three cupredoxin like domains (3dMCO), a few laccases from bacteria were also grouped into 2dMCO containing two such domains (Janusz et al. 2020). Laccases can simultaneously oxidize a broad spectrum of substrates, including phenolic compounds, complex cyanides, metal ions and non-aromatic compounds, with oxygen molecules as the final electron acceptor.

The catalytic reaction cycle of laccase is illustrated in Fig. 4. Typically, laccases contain one Cu1 (T1- type-1 copper), one Cu2 (T2- type 2 copper) and two Cu3 (T3- type 3 copper), with Cu2 and Cu3 forming a trinuclear cluster. Cu1 is the initial electron acceptor, which is present in the cavity near the enzyme's active site. The reduction of Cu1 is the rate-determining step for the laccase-catalyzed reactions. The relatively low values of Cu1 reduction potential (420–790 mV vs. normal hydrogen electrode, NHE) limit the substrates with molecules containing phenolic moieties. Laccase converts phenol to phenoxyl radicals, which are then converted into endproducts via coupling polymerization or radical rearrangement. The electron accepted by Cu1 is then shuttled to the trinuclear cluster, where oxygen is reduced to water (Chaurasia et al. 2013). In the MCOs superfamily, the enzymes having laccase-like catalytic activity are grouped as polyphenol oxidases, which includes tyrosinases, catechol oxidases and laccases, according to their substrate specificity and reaction mechanism (Janusz et al. 2020; Endo et al. 2003; Machczynski et al. 2004).

Fig. 4.

Laccase activity on phenolic dyes to generate radical intermediates. T1-type-1-copper (T1-Cu²⁺) accepts the electron from the substrate, which is then shuttled to the trinuclear cluster, consisting of two T3-type 3-copper (T3-Cu²⁺) and one T2- type 2-copper (T2-Cu²⁺). Laccase is completely reduced by Cu²⁺ converting to Cu¹⁺. Oxygen is the final electron acceptor, which is reduced to water

Among the laccases identified and characterized so far, fungal laccases account for the maximum number with various physiological roles, such as development and morphogenesis, lignin degradation, host–pathogen interaction and stress tolerance. Fungal laccases are mostly extracellular monomeric globular proteins of about 50–130 kDa (Senthivelan et al. 2016) with 10–25% glycosylation and isoelectric point of around 4.0 (Giardina et al. 2010). Many white rot fungi such as Dichomitus squalens (Šušla et al. 2007), Ischnoderma resinosum (Kokol et al. 2007), Pleurotus pulmonarius (Leo et al. 2019), Trametes hirsute (Couto et al. 2005) and Pycnoporus sanguineus (Salazar-López et al. 2017) were previously reported to be involved in laccase mediated dye-degradation. Laccase from Aspergillus proliferans decolorized synthetic dye solution containing aniline blue and Congo red around 40.9–70% and textile effluent 88.6% within 14 h (Kanmani et al. 2011).

As fungal laccases lack thermal and pH stability, the use and application of bacterial laccases are preferred. It has a broader range of operational parameters, such as temperature and pH. Moreover, industrial applications of bacterial laccases are cheap due to their broad specificity, short production time and easiness of expressing the gene in suitable host organisms (Chauhan et al. 2017). Many bacterial producers of laccases such as Bacillus (Kumar et al. 2022; Bu et al. 2020), Geobacillus (Jeon and Park 2020), Streptomyces, Nitrosomonas, Marinomonas, Pseudomonas, Enterobacter and Proteobacterium have been reported (Chauhan et al. 2017).

Azoreductases for dye degradation

Azoreductases (EC 1.7.1.6) are a class of flavoenzyme that reduces the azo bonds (–N = N–) present in the aromatic structure of azo dyes. Azoreductases from different sources have different structures, functions and cofactor requirements. Based on cofactor requirement, azoreductases are categorized into two groups—FMN (Flavin mononucleotide)—dependent and FMN-independent. FMN-dependent azoreductases are further grouped as NADH-dependent, NADPH-dependent, or both. Based on oxygen transfer, azoreductases are classified into oxygen-sensitive and oxygen-tolerant azoreductases (Misal and Gawai 2018). Azoreductases produced by anaerobes are highly sensitive to oxygen, whereas azoreductases produced by aerobes have a high tolerance to oxygen. Further, based on location, azoreductases are either membrane-bound or intracellular enzymes. Intracellular azoreductases are often thermostable, hydrophilic and highly efficient in degrading azo dyes (Sarkar et al. 2020). Microbial azoreductases used in dye decolorization are given in Table 2.

Table 2.

Azoreductases from the microbial origin for dye decolorization

Source	Dye decolorized (% removal)	Conditions	Activity	Km	Vmax	References
Streptomyces sp. S27	Methyl red (99%)	Whole cell biocatalyst, 30 ºC; pH- 6.0; Time- 2 h	13.92 U/g (NADH as co-enzyme) 15.59 U/g (NAD+ as co-enzyme)	–	–	Dong et al. (2019)
Halomonas sp. GT	Acid Brilliant Scarlet GR	35 ºC; pH- 6.0; Time- 2 h	0.27 μmol min−1 mg−1	12 μM	–	Tian et al. (2019)
Pseudomonas aeruginosa ASU3	Methyl red	40 ºC; pH- 7.0; 200 µM NADH;	0.3 ± 0.005 U/mg protein	125 µM	0.5 µM/min/mg	Elfarash et al. (2017)
Pseudomonas aeruginosa ASU6			0.92 ± 0.11 U/mg protein	3240 µM	10.6 µM/min/mg
Escherichia coli	Brilliant black	pH-7.2	3.82 U (NADH + FMN)	–	–	Zahran et al. (2019)
Enterococcus faecalis			12.89 U (NADH)
Enterococcus avium			7.93 U (NADH + FMN)
Bacillus cereus			2.19 U (NADH + FMN),
Chromobacterium violaceum	Methyl red	35 ºC; pH- 7.2	–	38.23 μM	2.30 μM/s	Verma et al. (2019)
	Amaranth			59.94 μM	2.61 μM/s
	Methyl orange			56.44 μM	1.63 μM/s
Shewanella sp. IFN4	Acid red 88	45 ºC; pH- 8; NADH as coenzyme	1.26 U/mg	–	–	Imran et al. (2016)
	Reactive black 5		3.29 U/mg
	Direct red 81		0.97 U/mg
	Disperse orange 3		1.56 U/mg
	Acid yellow 19		0.09 U/mg

Azoreductases break the azo linkages under aerobic and anaerobic conditions during the dye metabolism to generate aromatic amines, which will be eventually degraded by microbial enzymes such as mono-, di-oxygenases and hydrolases (Sarkar et al. 2020). Moreover, due to nitro-reduction ability, azo dyes can reduce toxic nitro-aromatic compounds. Azoreductases have a broad spectrum of stability for both pH and temperature. Azoreductases are usually stable in a pH range of 5–9, with optimal activity under physiological conditions (Misal and Gawai 2018). In contrast, azoreductase from Rhodococcus opacus has a higher dye degradation rate between pH of 3.8 and 5 (Qi et al. 2017). Azoreductase is stable within the 25–85 °C range with optimal activity within 35–40 °C (Misal and Gawai 2018; Maier et al. 2004). However, azoreductase produced by Bacillus badius was found to be stable till 85 °C and most efficient at 60 °C (Misal et al. 2011).

Azoreductases are present either as a monomer or as a homo-dimer. FMN-independent azoreductases are generally monomeric in nature (Misal and Gawai 2018), whereas FMN-dependent azoreductase from Enterococcus faecalis (Liu et al. 2007), E. coli (Langer et al. 2013) and Bacillus sp. B29 (Ogata et al. 2010) were found to be homo-dimers. However, NADPH-dependent azoreductase from S. aureus was a tetramer (Chen et al. 2005).

The catalytic mechanism of azoreductases is illustrated in Fig. 5. Crystallographic studies of FMN-dependent azoreductases confirmed that the degradation of azo dyes is by a ping-pong mechanism. At first, the oxidized FMN, which is present in the reactive site, is reduced by reducing equivalents such as NADH and NADPH (Ogata et al. 2010). Upon reduction, FMN transfers one or two hydrides from N atoms to the azo bonds present in the dye in one or two cycles with the formation of hydrazines which are further converted to two amines (Misal and Gawai 2018; Liu et al. 2007).

Fig. 5.

Azoreductase on azo dyes with the formation of colourless amines. In this example, NADH serves as an essential reducing agent for the enzyme located in the bacterial cellwall. The reduced enzyme reduces the redox mediator which in turn reduces azo dyes to two amines

Peroxidases for dye degradation

Peroxidases belong to the family of oxidoreductases. Most peroxidases are heme proteins comprising iron-porphyrin IX as the prosthetic group in the catalytic site. Heme peroxidases are divided into three types: peroxidase-cyclooxygenase superfamily (animal peroxidase), peroxidase-catalase superfamily (non-animal peroxidase) and dye-decolorizing peroxidases (DyPs) (Xu et al. 2021; Pandey et al. 2017; Chen and Li 2016). Based on the homology peroxidase-catalase superfamily is subdivided into class I, II and III. Peroxidases such as lignin peroxidases (LiPs; EC 1.11.1.14), manganese peroxidase (MnP; EC 1.11.1.13), and versatile peroxidases (VP; EC 1.11.1.16) belongs to class II whereas, catalase peroxidase (CP; EC 1.11.1.6) belong to class I. Plant peroxidases such as horseradish peroxidase (HRP) and barley grain peroxidase are class III peroxidases (Lin et al. 2018).

Among these peroxidases, LiPs, MnPs, HRP and DyPs are associated with the decolorization and/or degradation of many synthetic textile dyes. DyPs act on a broad range of synthetic dyes like anthraquinone derivatives, such as Reactive Blue 5, which is a poor substrate for other peroxidases. DyPs, with unique primary and secondary protein structure, can act on some pertinacious substrates such as dyes, β-carotene, aromatic compounds and sulphides (Xu et al. 2021). The catalytic mechanism of peroxidases is illustrated in Fig. 6. Peroxidases utilize H₂O₂ as their terminal electron acceptor while oxidizing the dye molecule. Lignin peroxidase directly oxidizes the target dyes, whereas Manganese peroxidases first oxidize Mn²⁺ to Mn³⁺ preferably, which further oxidizes the target dye molecules. The degradation of azo dyes by peroxidases is by either symmetric or asymmetric cleavage (Goud et al. 2020).

Fig. 6.

Mechanism of dye degradation by peroxidases. Peroxidases degrade azo dyes either by symmetric or asymmetric cleavage

Microbial peroxidases used in dye decolorization are given in Table 3. Rajhans et al. (2020) reported that among the enzymes involved in decolorizing methyl orange, Congo red, trypan blue and Eriochrome black T by Geotrichum candidum, DyP-type peroxidase showed the maximum activity (900 mU ml⁻¹) followed by laccase (405 mU ml⁻¹) and lignin peroxidase (324 mU ml⁻¹). The addition of lignin to the growth medium induces the production of peroxidases (Ali et al. 2022a).

Table 3.

Peroxidases from the microbial origin for dye decolorization

Source	Dye decolorized (% removal)	Conditions	Activity	Km	Vmax	References
Bacillus amyloliquefaciens	Crystal violet (63%)	30–40 ºC pH- 4.0	–	–	–	Zhang et al. (2021)
Cerrena unicolor BBP6	Remazol Brilliant Blue R (79.7%) Methyl orange (69.2%) Crystal violet (51.2%) Brilliant Blue R (61.6%) Bromophenol blue (28.4%)	30 ºC pH- 4.6	154.5 U/L (MnP)	–	–	Zhang et al. (2020)
Aspergillus flavus	Direct red 31 (94%) Acid black 234 (85%)	35 ºC pH 4.0	732 U/L mg (MnP)	0.0886 mM	0.126 U/mg	Kalsoom et al. (2022)
Meyerozyma caribbica	Acid orange 7 (93.8%)	28 ºC	27 U/mL	–	–	Ali et al. (2022b)
Trametes sp. 48424	Indigo carmine (94.6%) Remazol Brilliant Blue R (85%) Remazol Brilliant Violet 5R (88.4%) Methyl green (93.1%)	70 ºC (MnP)	1983.8 U/L	–	–	Zhang et al. (2016)
Aspergillus niger CTM10002	98% Acid blue 158	40 ºC pH 5.0	372 U/mg	0.169 mM	372 U/mg	Hamdi et al. (2022)
Bacillus albus	99.27% Methylene blue	30 ºC pH 7.0	–	–	–	Kishor et al. (2021)
Pleurotus eryngii	73% Carmine indigo dye	35 ºC pH 7.0	10.60 mmol/mL	–	–	Lopes et al. (2020)

Whole cells for dye decolorization

Whole cells of dye-degrading microbes could be effectively used for the decolorization of textile dyes (Table 4). Textile effluents are treated using whole cells through biosorption and biodegradation (Azeez and Al-Zuhairi 2022). The introduction of potentially pathogenic species during the dye effluent treatment could have a negative impact which is very difficult to manage (Pinheiro et al. 2022). This could be possibly reduced by using consortia of effective microorganisms (EMOs) or by genetically engineered microorganisms (GMOs).

Table 4.

Whole cells for dye decolorization

Microbe name	Dye name	Degradation/biosorption	Single/consortia	Operational conditions	Percentage removal	References
Enterobacter sp. CV–S1 (B, S, L)	Crystal violet	Degradation	Single	72 h at 35 °C and pH 6.50	100%	Roy et al. (2018)
Sphingomonas paucimobilis (B, S, L)	Malachite green	Degradation	Single	24 h at 25 °C and pH 9	99 to 75% for 2.5 to 50 mg/l dye after 24 h	Ayed et al. (2009)
Klebsiella pneumoniae K2, Enterobacter sp. K16b and Vibrio tritonius K20	Congo red	Degradation	Consortia	24 h, 37 °C and pH 7.3	86.7%	Chaieb et al. (2022)
	Malachite green				93.43%
Aspergillus foetidus (F, S)	Reactive black 5	Biosorption	Single	30 and 50 °C; pH:2.3; agitation 200 rpm; amount of pellets: 0.2 ± 0.05 g (w/w)	67% for live biomass and 97% for alkali pretreated	Patel and Suresh (2008)
Alcaligenes sp. BDN1, Bacillus sp. BDN2, Escherichia sp. BDN3, Pseudomonas sp. BDN4, Provedencia sp. BDN5, Acinetobacter sp. BDN6, Bacillus sp. BDN7 and Bacillus sp. BDN8 (B, C)	Reactive blue 160	Biodegradation	Consortia	4 h, 37 °C and pH 7	100% for 50 to 100 mg/L of dye	Balapure et al. (2014)
Consortia dominated by strains of Acinetobacter, Comamonas, Erwinia, Trichococcus, Dysgonomonas and Citrobacter	Congo red	Biodegradation	Consortia	96–120 h	85 to 97%	Neihsial et al. (2022)
Aspergillus niger, Aspergillus terreus and Rhizopus oligosporus (F, S, L)	Procion red MX-5B and Acid blue 161	Biodegradation		2 h	60% using Aspergillus terreus	Almeida and Corso (2019)
Aspergillus fumigatus (F, D)	Methylene blue	Biosorption				Kabbout and Taha (2014)
Chlorella pyrenoidosa (A)	Direct red 31			180 min	80.12%	Behl et al. (2019)

Biosorption is a metabolically passive process involving adsorption and absorption. The rate and efficiency of biosorption are directly correlated to the microbial cell wall composition, as the charged groups of the dyes could attract lipids and heteropolysaccharides present in the cell walls of live or dead cells. Dead cells are preferred over live cells as they can be used, stored and regenerated with minimum maintenance (Pinheiro et al. 2022). The cells of bacteria, fungi, algae and yeast species could be utilized to remove dyes from the textile effluent by acting as a chelating and complexing sorbent (Almeida and Corso 2019; Abbas et al. 2014). The ultrasound-assisted (35 kHz) adsorption of reactive yellow dye -145 using marine Chlorella sp. was highly effective, with 99% dye removal in 1 min (Amin et al. 2020). However, biosorption is not a sustainable method for large-scale dye decolorization as the secondary disposal of the biomass could be a problem (Pinheiro et al. 2022).

Bacterial degradation is economical and eco-friendly, with a broad range of substrate specificity. During degradation, enzymes such as reductase, dehydrogenase, kinase, hydrolase, transferase, catalase and dioxygenases were reported to be induced, which may result in complete degradation (Chaieb et al. 2022).

The bacterial cell system can be used in single or consortia. The use of mixed bacterial cultures is advantageous as it has broad substrate specificity and synergistic action of microbial consortia with diverse catabolic capabilities (Balapure et al. 2014). For instance, azo dyes are transformed into aromatic amines by azoreductases produced by anaerobes, which are further completely degraded by aerobes (Li et al. 2019). Using bacterial community from the polluted environment may impart the capability to degrade the dyes as it constantly evolves due to the selective local pressure.

Fungi that can withstand a broad range of temperatures, pH and inhibitory agents are also effective in decolorizing dyes (Soliman et al. 2013; Fu and Viraraghavan 1999). Fungi, specifically white rot fungi, produce several non-specific, extracellular oxidative enzymes (laccase, lignin peroxidase, manganese peroxidase, etc.) and hydrolytic enzymes (xylanases, cellulases, etc.) which could degrade a broad range of tenacious compounds including synthetic dyes (Tišma et al. 2010). The implementation of filamentous fungi in the decolorization process is a potent substitute because of less cost and the potential complete mineralization (Silva et al. 2018). Algal biomass can degrade numerous recalcitrant compounds, including dyes, heavy metals and pesticides (Kalesh and Nair 2005; Jothinayagi and Anbazhagan 2009; Alp et al. 2012; Palmer 1969).

Immobilization strategy

Immobilization is the process of confining the enzymes/cells using suitable support material by either physical or chemical methods, thereby retaining their activity and enhancing their reusability. Moreover, the use of immobilized systems is advantageous, with superior degradation capability and reduced sensitivity to environmental parameters like temperature, pH, amount of harmful contaminants, etc., compared with the suspended culture (Kurade et al. 2019). Immobilization strategies include adsorption, entrapment, covalent immobilization and cross-linking (Fig. 7). Heterocatalytic systems employing immobilized enzymes and whole cells for dye decolorization are summarized in Tables 5 and 6, respectively.

Fig. 7.

Immobilization types used for heterogeneous catalytic degradation of textile dyes

Table 5.

Immobilized enzyme system for dye decolorization

Enzyme used	Type of immobilization	Target dyes	Support for immobilization	Reactor and reaction conditions	References
Laccase from Trametes versicolor	Adsorption	Mixture of Brilliant blue, Reactive brilliant blue, Methyl orange and Crystal violet	Graphene oxide- polyethersulfone membrane	Nanofiltration system with biocatalytic membrane	Xu et al. (2018)
Recombinant dye decolorizing peroxidase from Aspergillus oryzae	Adsorption	Remazol Brilliant Blue R	silica-based mesocellular foam	1.5 mL vials; 30 °C; 100 strokes per minute	Shakeri and Shoda (2010)
Laccase Tplac from white rot fungus Trametes pubescens	Entrapment	Anthraquinone and azo dyes	Entrapped onto chitosan beads with glutaraldehyde	Shake flask; dark chamber at 50 °C with shaking speed at 150 rpm	Zheng et al. (2016)
Ginger peroxidase	Entrapment	Textile effluent	Agarose/agarose- guar gum hydrogel	Continuous packed bed reactor; 30 °C, Flow rate 10 mL/h, Residence time 12 min	Ali and Husain (2018)
Laccase from strain Brevibacterium halotolerans	Entrapment	Congo red	alginate -gelatin mixed gel	Shake flask, 100 rpm, 24 h, 50 °C, pH- 5.5	Reda et al. (2018)
Laccase from Bacillus sp. MSK-01	Entrapment	Textile dye effluent	Laccase-ABTS system on Copper alginate beads	Continuous flow packed bed bioreactor, Flow rate- 0.8 mL/ min, optimum temperature and pH were 85 °C and 10 respectively	Sondhi et al. (2018)
Manganese peroxidase from Ganoderma lucidum	Entrapment	Reactive Red 195A, Reactive Blue 21 and Reactive Yellow 145A	Agar–Agar matrix	Shake flask, 30 °C, 150 rpm for 12 h	Bilal et al. (2016b)
Manganese Peroxidase from G. lucidum IBL-05	Encapsulation	Complex sandalfix Red C4BLN dye	Gelatin hydrogel	Shake flask, 30 °C, 150 rpm for 5 h	Bilal et al. (2016a)
Crude Enzymes (Trametes versicolor and Pestalotiopsis sp. NG007)	Encapsulation	Textile dyes Lefavix Blue 16 (LB16), Reactive Remazol Violet 9 (RRV9) and Reactive Remazol Navy 4 (RRN4)	Double layer of Alginate beads	Packed bed bioreactor, Flow rate 1.5 mL/min	Yanto et al. (2014)
Laccase from genetically modified Aspergillus	Covalent attachment	Textile dyes	Green coconut fiber	Shake flask, 35 °C, pH- 7.0, 240 rpm	Cristóvão et al. (2012)
Recombinant Azoreductase from Brevibacillus laterosporus TISTR1911	Histidine-tagged enzyme is immobilized on nickel-chelating enzyme	Methyl orange	Double hexahistidine-tagged enzyme immobilized onto a nickel chelating column	Packed bed metal affinity reactor; Flow rate- 600 mL/h with an intermitting feed of NADH at 0.5 h interval	Lang et al. (2013)
Laccase from Ganoderma sp. KU-Alk4	Entrapment	Indigo carmine	Copper alginate beads	Airlift bioreactor; aeration 4 L/min	Teerapatsakul et al. (2017)

Table 6.

Whole-cell immobilized system for dye decolorization

Cells used	Type of immobilization	Target dyes	Support for immobilization	Reactor and reaction conditions	References
Enterobacter agglomerans	Entrapment	Methyl red	Calcium alginate beads, vermiculite cells, cooper beech, polyacrylamide Gel	Fluidized bed bioreactor, Shake flask, 25 °C; 100 rpm	Moutaouakkil et al. (2004)
Bacillus firmus	Entrapment	C. I direct red 80	Ca-alginate tubular polymeric gel	Tubular polymeric gel bioreactor; 30 °C; aeration- 100 mL/min	Ogugbue et al. (2012)
Stenotrophomonas maltophilia AAP56	Entrapment	Methylene blue, Toluidine blue, Methyl green, Indigo blue, Neutral red, Congo red, Methyl orange and Reactive pink	alginate gel, agar gel, polyacrylamide gel	Downflow fixed column reactor; Agitated condition at 30 °C, Residence time- 4.03 h, Dilution rate 0.3 h−1	Galai et al. (2010)
Pseudomonas luteola	Entrapment	Reactive red 22	Alginate- silicate sol–gel beads	500 mL flask in static incubation condition at 50 °C	Chen and Lin (2007)
Phanerochaete chrysosporium	Adsorption	Methylene blue	Mineral kissiris	Shake flask, 37 °C, 11 days	Karimi et al. (2006)
Sphingomonas sp. BN6	Entrapment	Mordant yellow3 (MY3)	Sodium Alginate beads (anaerobic) and calcium alginate beads (aerobic)	anaerobic expanded-bed reactor, and an aerobic airlift-loop reactor	Kudlich et al. (1996)
Lysinibacillus sphaericus D3	Entrapment	Xylidine orange	Immobilized beads in natural gel sodium alginate	Shake flask, 120 rpm, pH 7.2	Devi et al. (2017)
Trametes versicolor	Entrapment	Direct Blue 15	Immobilized in sodium alginate beads	Packed bed bioreactor; jacketed glass column with temperature maintained at 26 °C;	Pazarlioglu et al. (2010)
Trametes hirsuta	Entrapment	Phenol red, Indigo carmine	Calcium alginate polymer beads	Airlift bioreactor; Batch mode; 30 °C; aeration 1 L/min	Domínguez et al. (2005)
Aeromonas jandaei SCS5	Entrapment	Methyl red	Polyvinyl alcohol-sodium alginate -kaolin mixture with antraquinone-2,6-disulfonate or Fe3O4 nanoparticles as mediators	Flask level, 35 °C; pH 7.0	Sharma et al. (2016)
Proteus vulgaris NCIM 2027	Entrapment	Reactive blue 172	Calcium alginate, k-carrageenan, polyacrylamide, phosphorylated Polyvinyl alcohol gel beads	Fixed bed bioreactor; Volumetric flow rate- 15 mL/h	Saratale et al. (2011)
Escherichia coil CD-2	Entrapment	Azo dyes	Polyvinyl acetate/ calcium alginate/ gelatin	Flask level, 37 °C; Aerobic condition	Cui et al. (2015)
Bacillus cohnii RAPT1	Biofilm formation	Reactive red 120	Polyurethane foam	Packed bed reactor; 35 °C; pH 8.0	Padmanaban et al. (2016)
Bjerkandera sp. strain BOL 13	Fungal mycelia overgrown in carrier discs	Reactive red 2 and Reactive blue 4	Birchwood discs	continuous rotating biological contactor reactor; 20 rpm; aeration 900 mL/min; 5.5 mL/h; pH- 6.0	Axelsson et al. (2006)
Bacillus subtilis	Biofilm formation	Congo red	Polyurethane foam	Multistage fluidized bed bioreactor; air flow rate 0.2 LPM	Setty (2019)
Bjerkandera adusta OBR105	Self-immobilized fungal pellet	Acid dyes (red 114, blue 62, and black 172) and reactive dye (blue 4)	Self-immobilization	Air lift bioreactor, aeration 1 L/min	Sodaneath et al. (2017)

Adsorption

Adsorption is the most widely adopted immobilization approach, as the native conformation of the enzyme is conserved. Adsorption of the enzyme on the carrier material uses weak forces of interaction such as the van der Waals force of attraction, ionic interactions and hydrogen bonding. A wide range of carriers can be employed for enzyme adsorption and the selection of the carrier for successful adsorption depends upon the cost, availability and stability. Moreover, carriers should possess suitable morphology, pore size and specific surface area. In case such groups are absent, chemical modifiers are required to amend the binding efficiency (Jesionowski et al. 2014). This method is simple but vulnerable to leaching due to the minimal interaction between the carrier and the enzyme (Imam et al. 2021).

Laccase showed more affinity towards organic carriers than inorganic carriers. Many organic carriers (chitin, chitosan, alginate, cellulose and synthetic polymers) and inorganic carriers (silicas, hydroxyapatite and titania) have been reported (Jesionowski et al. 2014). According to the literature, peroxidases were immobilized by adsorbing on silica-based carriers (Shakeri and Shoda 2010), magnetic nanoparticles (Mohamed et al. 2017), carbon nanotubes (Oliveira et al. 2018) and mesoporous activated carbon (Torres et al. 2017). Peroxidases were immobilized on nanohybrids fabricated with Fe₃O₄/Ag/Graphene oxide nanoparticles to remove 2, 4 dichlorophenols (Sarnoab and Iulianoc 2020).

Entrapment

Enzymes can be irreversibly entrapped inside a porous matrix that permits substrate and products through the matrix support. Here, the matrix is a solid or gel that develops due to an elemental reaction, which becomes part of the support in the presence of an enzyme to be immobilized. The matrix prevents the direct contact of the enzyme with the bulk reaction liquid. Organic materials like biopolymers or synthetic polymers are frequently used as entrapment matrixes. The entrapment of enzymes inside these gels is achieved typically in four ways, i.e., sol–gel method, polymerization to develop insoluble polymers, cross-linking of biopolymers and supramolecular assembly (Imam et al. 2021).

Imam et al. (2021) confirmed that the gel, which comprises of cross-linked matrix and a confined liquid renders high resistance to the inactivation of the enzymes as the native structure is preserved (Imam et al. 2021). Previous studies proved that the entrapped peroxidases could be reused for several cycles, in which the decolorization efficiency was protected even after several repeated uses. Despite all these merits, entrapment is not widely used on an industrial scale due to its drawbacks, such as mass transfer limitation, leaching of enzymes through larger pores, etc. (Jun et al.  2019).

A purified Pleurotus ostreatus IBL02 laccase entrapped in the sol–gel matrix comprised of trimethoxysilane and proplytetramethoxysilane was used in the decolorization of reactive textile dye Drimarene brilliant red K-4BL. The entrapped enzyme shower higher K_m and V_max in comparison with that of the free enzyme (Asgher et al. 2012).

Covalent immobilization

Enzymes can attach covalently to a support substance, typically a polymer that can be either organic or inorganic. The organic supports include chiefly polysaccharides (such as modified celluloses, dextran, chitosan, agarose, etc.), vinylic (such as polyvinyl alcohol), acrylic polymers (polyacrylamide) and polyamides. Inorganic supports include silica-based and oxide-based materials (such as ceramics, titania, zirconia, alumina, etc.) (Zucca and Sanjust 2014). These support materials are chemically modified to provide specific binding sites for the enzymes. This is usually done by functionalization of the support followed by chemical modifications to get desired electrophilic groups. Among all the immobilization methods, covalent immobilization offers minimal leakage through the high bond strength between carrier and enzyme. Moreover, covalently immobilized enzymes are commercially viable with better stability, recyclability and minimal mass transfer resistance (Imam et al. 2021; Zucca and Sanjust 2014). However, covalent binding may affect enzyme activity by altering its polarity.

Cross-linking

Enzymes are crosslinked with each other or with another protein and/or with a support matrix using a crosslinking agent in a two-step process. During the initial step, the aggregates or crystals of enzymes to be immobilized are produced by adding aggregating agents. The aggregating agents may be organic or inorganic precipitants such as glycol, dimethoxyethane, acetone, acetonitrile and ammonium sulphate. The aggregate/crystal thus formed is chemically coupled to each other to form crosslinked enzyme aggregates (CLEAs) using crosslinking agents. The most commonly used crosslinking agents are glutaraldehyde, benzoquinone, or dextran-polyaldehyde for lysine-specific crosslinking, while polyethyleneimine and carboxylate activating carbodiimide are used for glutamic or aspartic acid-specific crosslinking (Imam et al. 2021). This type of immobilization provides high specific enzymatic activity, better storage and operational stability and less cost as the carriers are not mandatory (Bilal et al. 2017a).

Bilal et al. (2017a) immobilized manganese peroxidase (MnP) from white-rot fungus, Ganoderma lucidum IBL-05, by glutaraldehyde cross-linking of aggregates prepared by using aggregating agents such as acetone, ammonium sulfate, ethyl alcohol, 2-propanol and tert-butyl alcohol. This immobilized MnP was used to degrade endocrine disruptors and dye-based chemicals in a packed-bed reactor. Kulkarni et al. (2020) prepared microcapsules of CLEAs with extracellular enzymes from the lichen Dermatocarpon vellareceum using ammonium sulphate as the aggregating agent and glutaraldehyde as cross-linking agent for the decolorization of several dyes such as Brilliant Blue R (BBR), Direct Red 2B, Brilliant Blue GRL, Green HE4B, Red M5B, Blue 2RNL, Reactive Orange 3R, Yellow HE4G and Orange 2RX.

Other strategies

Micellar system of laccase enzyme using ionic liquids such as tetraalkylammonium- and imidazolium-based cationic surfactants and cholinium-based anionic surfactants was used for the degradation of highly hydrophobic dye, indigo carmine with high efficiency. An aqueous solution of ionic liquids above their critical micellar concentration formed micelles that could encapsulate both enzyme and the liquid (Bento et al. 2020). Nanoparticles with a size similar to the enzymes can be used for immobilization. The immobilized peroxidase enzyme on Fe₃O₄ magnetic nanoparticles prepared by coprecipitation with glutaraldehyde showed remarkable stability upon storage for three months and recycling up to 100 cycles (Darwesh et al. 2019).

Bioreactor consideration with immobilized system

In general, to treat textile dye effluent, various heterogeneous bioreactors such as packed bed reactors, rotating biological contactors (RBCs), fluidized bed reactors and air lift bioreactors have been used (Pocedič et al. 2009). Among them, packed-bed reactors are effectively used as biofilters in treating textile dyes because of their ease of operation, high yield, high liquid residence times and ease of scale-up.

Packed-bed biofilters are contacting columns packed with immobilized systems through which the effluent to be treated passes through. For the economical and effective textile effluent treatment, microbial biofilters are reported to be superior to the enzymatic system (Chaudhary et al. 2003). Microbial biofilters are commonly used in continuous mode with enhanced contact time and lower capital costs. It consists of biofilms attached to the surface of the porous solid matrix and is operated either in submerged or trickling mode (Pocedič et al. 2009). Packed-bed biofilters operating in trickling mode are more suitable when the ligninolytic fungal mycelium is used for decolorizing dye effluent (Svobodová and Novotný 2018). Padmanaban et al. (2016) demonstrated a packed bed reactor with immobilized Bacillus cohnii RAPT1 cells on polyurethane foam as the packing material for the degradation of Reactive Red 120. Kurade et al. (2019) used an up-flow fixed bed reactor to decolorize Remazol red, an azo dye, using an immobilized microbial consortium of Brevibacillus laterosporus and Galactomyces geotrichum on various carriers such as calcium alginate, polyvinyl alcohol, stainless steel sponge and polyurethane foam.

Similarly, immobilized enzymes could also be used as biofilters (Lopez-Barbosa et al. 2020). Bilal et al. (2017b) employed a packed bed reactor to degrade Remazol brilliant blue R, reactive black 5, Congo red and crystal violet using chitosan-immobilized HRP, with consistent decolorization for continuous six batches. Lang et al. (2013) used a packed bed metal affinity reactor with double hexahistidine tagged azoreductase immobilized on a nickel chelating column for complete degradation of methyl orange.

Rotating biological contactors (RBC) consist of a closely spaced circular disc mounted on a horizontal shaft which will slowly rotate over the effluent in a basin where microbial biofilms are formed over the discs as a static biofilm (Abraham et al. 2003). The mild rotation of the discs enables the removal of excess biomass and alternate exposure to media and air. The biofilm formation on the support of RBC depends on initial adhesion and the stability is provided by forming a glycoconjugate exopolysaccharide (EPS) matrix (Möhle et al. 2007). Earlier studies using RBC with immobilized fungi Coriolus versicolor (Kapdan and Kargi 2002), Phanerochaete chrysosporium (Pakshirajan and Kheria 2012), and Trametes versicolor (Del Álamo et al. 2020) confirmed a significant dye decolorization. Abraham et al. (2003) used an aerobic bacterial consortium to degrade sulfonated azo dyes in RBC without any additional carbon source.

Fluidized bed reactor (FBR) is used for the biological treatment of industrial effluent as it can be used continuously with less area requirements. Moreover, it provides excellent particle mixing, a sizeable liquid–solid interface area and superior isothermal temperature distribution. FBR consists of a static bed which is fluidized by passing a fluid through the bed (Tisa et al. 2014). Previously FBR has been used for dye decolorization with solid catalysts employing chemical approaches such as Fenton’s process (Farshchi et al. 2018; Wang et al. 2015; Su et al. 2011) and photocatalytic degradation (Couto et al. 2002). Apart from this, FBR is used in biological dye decolorization with immobilized enzymes (Zolfaghari et al. 2018) or cells (Moutaouakkil et al. 2004) on supports as the bed. However, during fluidization, several operational difficulties, such as bumping, spouting and slugging of the bed, may occur (Sodaneath et al. 2017).

Three-phase airlift bioreactor consisting of immobilized cells (Sodaneath et al. 2017) or enzymes (Teerapatsakul et al. 2017) is reported for dye degradation. The potential washout of the bed in FBR can be controlled by using an internal gas draft tube to convert it into an airlift bioreactor (Zhu 2007). Here the gas draft tube acts as a riser through which air is sparged, resulting in the upflow of the liquid inside the riser. Eventually, the air escapes from the top and the gas-free liquid medium with suspended particles will fall through the down comer resulting in pseudo-homogeneity inside the reactor (Valdivia-Rivera et al. 2019). The cyclical flow inside an airlift bioreactor offers several advantages, such as preventing bubble coalescence, even distribution of shear stresses and enhanced heat and mass transfer (Wang and Zhong 2007).

Conclusion

Microbial cells and their enzymes are commonly used for the dye decolorization process. The action of the enzymes depends upon the class of the dye. Dyes are classified based on source, ionic strength, structure and applications. Microbial enzymes involved in dye decolorization are mainly laccases, azoreductases and peroxidases. The utilization of whole cells can ensure the synergistic action of several enzymes. The decolorization of dyes by whole cells is either by biosorption, bioaccumulation and/or biodegradation. Sometimes the use of solo enzymes of single microbes could produce toxic by-products. Immobilized multienzyme systems and microbial consortiums may be considered in this regard. Moreover, immobilization will improve reusability and reduce the impact on the environment by the biological system. The immobilized system of whole cells or enzymes could be operated in a packed bed reactor, rotating biological contactors, fluidized bed reactors, or airlift bioreactor. From the presented survey, using an immobilized system in a heterocatalytic reactor could be the best method for the economical and effective treatment of textile dye effluent.

Data availability

The authors confirm that the data supporting the findings of this study are available within the article.

Declarations

Conflict of interest

All the authors declare that they have no potential conflict of interest.

Ethical approval

This article does not contain any studies with human participants or animals performed by any of the authors.

Informed consent

None.