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https://doi.org/10.1007/s42398-020-00150-w
REVIEW
Mycoremediation of synthetic dyes by yeast cells: a sustainable
biodegradation approach
M. Danouche1,2
· H. EL Arroussi1 · N. El Ghachtouli2
Received: 19 December 2019 / Revised: 25 October 2020 / Accepted: 20 November 2020
© Society for Environmental Sustainability 2021
Abstract
Dye effluents released from various industries, notably the textile sector, are hazardous, and can cause significant damage
to the environment. Thus, treatment and detoxification of these toxic dyes are of major concern for compliance with environmental legislations. So far, a number of physicochemical dye-removal methods have been proposed. However, despite
their effectiveness in dye decolorization, by-products of chemical degradation methods may be more toxic than their parent
dye molecules. The cost of these processes is also very high, thereby limiting their large-scale application. The use of yeast
cells for the removal of toxic dyes is a comparatively effective, eco-friendly and cost-effective method. In this review, we
describe the adverse effects of synthetic dyes on living organisms and enzymatic biodegradation mechanisms involved in
mycoremediation processes of synthetic dyes. In addition, the influence of various physico-chemical factors on the decolorization performance of yeast cells, the analytical techniques used to identify the intermediates of dye biodegradation, the
assessment of their toxicity and the molecular aspects of their biodegradation are also highlighted. This study may provide
a basis for the development of dye bio-removal methods using yeast cells.
Graphical abstract
Keywords Yeast · Decolorization · Enzymatic biodegradation · By-product identification · Toxicity assessment
Introduction
* M. Danouche
mohammed.danouche@usmba.ac.ma
Extended author information available on the last page of the article
Since the beginning of the industrial revolution in the nineteenth century, the equation for balanced economic growth
while protecting the environment has not yet been resolved.
In fact, soil, air, and water pollution are among the main
global challenges facing the world today. More critical than
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ever, there is a growing concern about the impact of water
contamination on aquatic life and public health. Compared
to other industrial sectors, the textiles industry consumes
large quantity of water; it was estimated that about 200–500
L of water is required to produce 1 kg of finished textile
products (Waghmode et al. 2012a). Therefore, this sector generates significant volume of wastewater composed
of suspended solids, surfactants, heavy metals and several
synthetic dyes (Balapure et al. 2015; Chen et al. 2009). The
exact amount of dyes released into the environment is uncertain, it has only been estimated that the overall use of dyes
in the textile sector is more than 10 000 tons per year, and
about 10–15% of this amount is lost as waste during the
dyeing process, due to the low chemical affinity of the dyes
used on textile fibers (Kunamneni et al. 2008; Saratale et al.
2011). Consequently, the release of such xenobiotics leads
to significant pollution, which not only affects the aesthetic
quality of water but also leads to serious impacts on exposed
organisms including humans (Lellis et al. 2019; Puvaneswari
et al. 2006). To cope with this issue, legislations have been
enacted worldwide for the management and the treatment
of these contaminants prior to their release into the environment. A variety of physicochemical methods have therefore
been used, particularly membrane processes, photochemical oxidation, and electrochemical processes (Arslan et al.
2016). Although these methods are effective, they require
a large amount of chemicals and energy-intensive facilities, which makes them expensive, thereby limiting their
large-scale applications (Barakat 2011; Fu and Wang 2011).
The most recommended approach is the use of biological
techniques, due to their advantages, including treatment efficiency, appropriate cost, and a minimal ecological impact
(Srinivasan and Viraraghavan 2010). The emergence of
this promising solution is based on the exploitation of the
ecological principles of bacteria, fungi, and microalgae in
the treatment of toxic dyes (Chen et al. 2009; Jafari et al.
2014; Miranda et al. 2013). While bioprocesses have several advantages, some drawbacks may also be noted, such
as the long time required for the bio-removal of dyes using
filamentous fungi or microalgae (Singh and Arora 2011).
Moreover, the decolorization of dyes using bacteria requires
two phases; the anaerobic reduction of Azo bonds, followed
by aerobic mineralization of the resultant aromatic amines
(Abiri et al. 2017; Roșu et al. 2019). However, the use of
yeast cells overcomes this drawback; they do not only grow
rapidly like bacteria, but exhibit great plasticity and ability to adapt to adverse growth conditions like pH fluctuation or temperature changes. Also, yeast does not require
special biphase growth conditions like bacteria. In addition,
the biodegradation mechanisms of dyes using yeast cells
involve different oxidases, that can directly break the azo
dyes through non-specific free radical mechanisms, avoiding
therefore the production of toxic intermediates like aromatic
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amines, that are typically produced after the specific cleavage of the azo bond of synthetic dyes by bacteria (Pandey
et al. 2007; Dave et al. 2015). Furthermore, yeast have a
special flocculating characteristic that allows them to aggregate into multicellular masses (flakes), facilitating, therefore,
their recovery after treatment of colored effluents (Soares
and Soares 2012). Hence, the use of yeast cells in the bioremoval of synthetic dyes has attracted more interest (Jafari
et al. 2014; Sen et al. 2016). Several studies focusing on the
decolorization mechanisms of the ascomycete and basidiomycete strains of yeast have demonstrated the involvement
of three bioremoval strategies, with active and/or passive
metabolism pathways, namely: extracellular biosorption (Dil
et al. 2017; Mahmoud 2016), intracellular bioaccumulation
(Das et al. 2010; Gönen and Aksu 2009) and/or intra and
extracellular biodegradation (Martorell et al. 2018; Tan et al.
2019). The present review focuses on the harmful effects
of synthetic dyes on aquatic environments and the state-ofthe-art of the biodegradation mechanisms of synthetic dyes
by yeast cells. Sequential summary studies on the biodegradation of synthetic dyes using different yeast species are
also discussed. Much emphasis will be projected towards
oxidoredactase enzymes involved in the biodegradation of
dyes, as well as the influence of operating factors on this bioprocess. Besides, the analytical methods used in the chemical characterization of dye biodegradation by-products and
the evaluation of their toxicity will be discussed. Throughout
the last section, the molecular aspects of the degradation of
dyes by yeast cells will be addressed in order to identify the
future scope of this research.
Adverse effects of synthetic dyes on aquatic
ecosystems
Since the dawn of human civilization, colorants have
played a crucial role in different aspects of everyday life.
Before the end of the nineteenth century, all dyes used
were obtained from natural sources, until the first artificial
dye was discovered by W.H Perkins (1838–1907) through
an accidental production of Mauveine. This successful discovery paved the way for the synthesis of chemical dyes
as we know them today. Since then, many chemical compounds have been identified as coloring substances (Rai
et al. 2005). Thus, the use of these chemicals has continued to increase, notably in textile and leather industries.
In addition to the type of color emitted by these chemicals,
other physicochemical requirements have become more
stringent in textile sectors including resistance to various
environmental conditions, such as persistence of colored
fabrics against washing, exposure to light, chemicals
and biological attacks (Khan et al. 2013). However, the
same requirements that confer higher resistance qualities,
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could complicate the dye’s removal from wastewater by
traditional methods. Meanwhile, the release of untreated
or inadequately treated colored wastewater into aquatic
environments can lead to various signs of toxicity to
exposed living organisms (Moopantakath and Kumavath
2018). The high concentration of dyes in aquatic ecosystems may prevent the penetration of light into the depths,
thus disturbing the photosynthetic activity of autotrophic
organisms, and impacting the re-oxygenation potential of
the receiving waters (Carmen et al. 2009). Moreover, the
toxicity of synthetic dyes may occur in aquatic ecosystems
across the food chains. Numerous studies have focused on
the adverse effect of dyes on species from various trophic
levels, including producer and consumer species (Puvaneswari et al. 2006; Tkaczyk et al. 2020). For instance, the
microalgae’s morphological, biochemical, and metabolic
properties can be influenced by increasing concentrations
of dyes. Gita et al. (2019) reported that a concentrationdependent decrease in the specific growth rate and pigment contents of Chlorella vulgaris was observed after
exposure to increasing concentrations of the textile dyes
Drimarene Blue, Optilan Yellow and Lanasyn Brown.
Similarly, Indigo dye induced a significant growth reduction of Scenedesmus quadricauda, and altered its morphological characteristics (Chia and Musa 2014). Likewise,
zooplankton organisms can be directly or indirectly influenced by dyes. Hernández-Zamora et al. (2016) showed
that exposure to low concentrations of Congo Red or
feeding on microalgae that already accumulated this dye,
induced adverse effects on the survival and reproduction of
Ceriodaphnia dubia. Additionally, many toxic dyes have
shown a direct effect on primary consumers like fish. For
instance, Kaur and Kaur (2014) investigated Poikilocytosis
occurrence in Labeo rohita fish as an indicator of stress
caused by the azo dye Basic Violet-1, with a highly toxic
effect (96-h LC50) at 0.45 mg L−1 concentrations. Srivastava et al. (2017) also noted that the exposure of L. rohita
to sublethal concentrations of Eriochrome Black T caused
considerable histopathological abnormalities and altered
antioxidant enzyme activities in the epidermis. Hence, the
accumulation of these xenobiotics in aquatic organisms
and their transmission to humans through the consumption of sea food from polluted sources may provoke many
human health problems (Chung 2016; Mendes et al. 2011).
To prevent these environmental issues, it is necessary
to introduce standards and framework laws to limit the use
of hazardous dyes or to substitute them with new synthetic
dyes that can meet industrial requirements, and that can also
be easily biodegradable. Therefore, research in synthetic
chemistry of dyes must not only compete with the criteria of resistance, but also their fate in nature. On the other
hand, it is important to establish promising approaches for
the treatment of these pollutants before their release into the
environments.
Biodegradation of synthetic dyes by yeast
Biodegradation is defined as an energy-dependent process,
involving the decomposition of organic compounds, into
smaller and simpler by-products, through different enzymatic reactions (Kaushik and Malik 2009). When the biodegradation is completed, the bioprocess is called mineralization. This form of treatment produces simpler products,
such as H2O, CO2, NH3, CH4, H2S and PO3, which are less
harmful. The same process is defined as biotransformation if the organic compounds are not completely mineralized (Martorell et al. 2017a). Many microorganisms
belonging to various taxonomic classes (bacteria, fungi,
and microalgae) have shown the ability to decompose a
broad variety of anthropogenic compounds, including
synthetic dyes (Chen et al. 2009; El-Sheekh et al. 2009;
Jafari et al. 2014; Miranda et al. 2013). In recent years,
mycoremediation or the biotechnological application of
fungi has become a model example for bioremoval of pollutants. It has been documented that a diversity of fungal
species could be used for removing a variety of toxic dyes,
mainly with white-rot fungi like Trametes versicolor and
Phanerochaete chrysosporium. Other fungi species such
as Rhizopus oryzae and Aspergillus niger, have also been
reported (Sen et al. 2016). Regarding the use of yeast in
the bioremediation of synthetic dyes, it was first mooted
back in 1992, when the yeast strain of Candida curvata
was used for the treatment of colored wastewater (Kakuta
et al. 1992). Until today, reports on the biodegradation of
synthetic dyes using yeast cells are limited compared to
other investigations on the biodegradation of dyes using
other microorganisms. The most studied yeast strains for
the biodegradation of synthetic dyes are Candida sp., Saccharomyces cerevisiae, and Pichia sp. which belong to
the Ascomycetes phylum, and only a few reports involve
Basidiomycetous yeast strains, such as Trichosporon sp.
and Pseudozyma rugulosa (Pajot et al. 2014).
Enzymes involved in the biodegradation
of dyes by yeast
The biodegradation of synthetic dyes using yeast cells is
performed with various intra-extracellular oxidase and
reductase enzymes (Fig. 1). Oxidases are the most studied
enzymes for the biodegradation of dyes by yeast. Moreover,
the activity of certain reductases has also been reported in
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Fig. 1 Intracellular and extracellular enzymatic biodegradation of synthetic dyes in yeast cells
some yeast strains (Table 1). The suggested mechanism for
the enzymatic biodegradation of azo dyes is shown in Fig. 2.
Oxidases are a class of enzymes that use O2 as an electron acceptor to catalyze oxidation–reduction reaction, and
thereby generating H2O or H2O2 as products. The oxidases
typically contain metal or Flavin coenzyme on their active
site (Phale et al. 2019). It was reported that the biodegradation of synthetic dyes by yeast cells can be accomplished through lignin-modifying enzymes, particularly
phenoloxidase like Laccase (Lac) and tyrosinase (Tyr) as
well as peroxidase such as manganese peroxidase (MnP)
and lignin peroxidase (LiP) (Martorell et al. 2012; Solís
et al. 2012).
Laccase (EC 1.10.3.2) Lac is a benzenediol oxygen reductase that can be expressed in different species
including lichens, bacteria and fungi. In fact, this enzyme
belongs to the classes of urushiol and P-diphenol oxidase, distinguished by a multicopper atoms in their catalytic center (Arregui et al. 2019). A high redox potential
(780 mV) allows these enzymes to oxidize many organic
compounds, such as polyphenols, methoxy-substituted
phenols, aromatic diamines, and several organic pollutants inclining dyes (Upadhyay et al. 2016). The fungal
Lac was found present in Ascomycetes, Deuteromycetes,
Basidiomycetes species, but is particularly abundant in
white-rot fungi that degrade lignin (Brijwani et al. 2010).
In addition, fungal Lac is an extracellular enzyme, this
peculiarity facilitates their isolation and purification compared to Lac from other organisms. Therefore, they have
gained particular commercial interest in organic syntheses,
pulp/textile bleaching, bioremediation, chemical grafting,
and surface modification of polymers (Jeon et al. 2012;
Viswanath et al. 2014). As shown in the Table 1, a number
of studies have documented the role of Lac in yeast cells in
the biodegradation of synthetic dye. The use of Lac for the
biodegradation of dyes does not require cofactors, since
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it directly breaks azo dyes via non-specific free radical
mechanisms, which does not create toxic by-products, such
as aromatic amines, typically produced after the specific
cleavage of the azo bond of azo dye (Dave et al. 2015).
Tyrosinase (E.C. 1.14.18.1) Try is a copper-containing
enzyme, known as polyphenol oxidase or monophenol
mono-oxygenase. It has been found in plants, filamentous
fungi, bacteria, and some yeast strains. This enzyme has a
range of industrial interests, for example in the biosynthesis
of melanin, the manufacture of L-dihydroxy phenylalanine,
as well as several environmental biotechnologies like the
detoxification of phenol-containing wastewater (Ates et al.
2007; Kim and Uyama 2005; Zaidi et al. 2014). With regard
to the involvement of Try in the decolorization of synthetic
dyes in yeast cells, only a few species revealed their role,
notably Galactomyces geotrichum MTCC (Waghmode et al.
2012a, b), S. cerevisiae MTCC 463 (Jadhav et al. 2007) and
Candida krusei strains (Charumathi and Das 2011). The
catalytic reaction of dyes with Try is performed in two
steps: The primary reaction catalyzing the o-hydroxylation
of monophenols to the corresponding catechols (monophenolase activity), followed by the second oxidizing monophenols to the corresponding o-quinones (Diphenolase activity)
(Duckworth and Coleman 1970).
Lignin peroxidase (EC 1.11.1.14) LiP is a glycosylated
enzyme that belongs to the oxidoreductase family that acts
on peroxide as an acceptor. This ligninolytic enzyme has a
high redox potential (700–1400 mV), with optimum activity
at acidic range (pH 3–4.5). Lip is a non-specific substrate
enzyme, which means that this characteristic confers it the
ability to catalyze the degradation of different phenolic and
non-phenolic aromatic compounds, including β-O-4 linkagetype arylglycerol-aryl ethers with a redox potential of up
to 1.4 V (in comparison with the normal hydrogen electrode) (Choinowski et al. 1999; Chowdhary et al. 2018). For
example, the biodegradation mechanism of sulphonated azo
Dyes
Conditions
D (%)/Time
Enzymatic activity
Sterigmatomyces halophilus
SSA-1575
Reactive Black 5
98/24 h
G. geotrichum GG
Acid Scarlet GR
P. occidentalis G1
Acid Red B
G. geotrichum MTCC 1360
Methyl Red
S. cerevisiae MTCC 463
Methy Red
50 mg L−1
30 °C/0 rpm
pH 5.0
100 mg L−1
30 °C /180 rpm
pH 7.0–8.0
50 mg L−1
30 °C/160 rpm
pH 5.0
100 mg L−1
30 °C/150 rpm
pH 3.0
100 mg L−1
30 °C/0 rpm
pH 6.5–9.0
G. geotrichum MTCC 1360
Remazol Red
Rubine GFL
50 mg L−1
30 °C /0 rpm
pH 7.0
96/36 h
87/96 h
S. cerevisiae MTCC 463
Malachite Green
95/23 h
S. cerevisiae
Malachite Green
C. krusei
Basic Violet 3
D. rugosa
Indigo
C. samutprakarnensis
Acid Red B
Candida sp VITJASS
Reactive Green
100 mg L−1
33 °C/150 rpm
pH 7.2
100 mg L−1
30 °C/150 rpm
pH 7.2
10 mg L−1
28 °C/120 rpm
pH 6.5
10 mg L−1
30 °C/0 rpm
pH 2.0
50 mg L−1
30 °C/160 rpm
pH 6.0
100 mg L−1
30 °C/120 rpm
pH 5.0–6.0
Al-Tohamy et al. (2020)
Oxidase: LiP, MnP and Lac
Reductase NADH-DCIP
reductase
Guo et al. (2019)
Oxidase: LiP and Lac
Reductase: NADH-DCIP
reductase
Song et al. (2018a, b)
Oxidase: LiP, MnP and Lac
Reductase: NADH-DCIP
reductase
Jadhav et al. (2008b)
Oxidase: LiP and Lac
Reductase: NADH–DCIP
reductase, MG-reductase
Jadhav et al. (2007)
Oxidase: LiP, Tyr, Lac and
Aminopyrine N-demethylase
Reductase: NADH-DCIP reductase, AzoR
Waghmode et al. (2012a, b)
Oxidase: Tyr and Lac
Reductase: NADH-DCIP reductase, AzoR, and Riboflavin
reductase
Jadhav and Govindwar (2006)
Oxidase: LiP and Lac
Reductase: NADH-DCIP reductase, MG-reductase
Biradar et al. (2016)
Oxidase: LiP and Lac
Reductase: NADH-DCIP reductase, MG-reductase
Charumathi and Das (2011)
Oxidase: LiP, Tyr and Lac
Reductase: NADH-DCIP reductase, MG-reductase and AzoR
Bankole et al. (2017)
Oxidase: LiP
Reductase: NADH-DCIP
reductase
Song et al. (2018a, b)
Oxidase: LiP
Reductase: NADH-DCIP
reductase
Sinha et al. (2018)
Oxidase: Lac
Reductase: NADH-DCIP reductase, AzoR
92/10 h
98/16 h
100/1 h
100/16 min
99/2 h
100/24 h
99.9/5 5 d
97/18 h
84/96 h
References
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Yeasts species
Environmental Sustainability
Table 1 Studies on the biodegradation of dyes by yeasts species
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Table 1 (continued)
Conditions
D (%)/Time
Enzymatic activity
References
Candida sp MM 4035
T. porosum MM 4037
C. satwnus MM 4034
Barnettozyma californica MM
4018
D. polymorphus
C. tropicalis
Yellow 4R-HE
Black B-V
Blue RR-BB
Red 7B-HE
200 mg L−1
25 °C/250 rpm
pH 4.0
64–96/24 h
Oxidase: LiP, MnP, Tyr, Lac
and N-demethylase
Martorell et al. (2012)
Reactive Black 5
100/18–24 h
Oxidase: MnP
Yang et al. (2008)
T. multisporum
T. laibachii
63–98/36–72 h
Oxidase: LiP, MnP and Lac
Pajot et al. (2007)
C. boidinii MM 4035
Yellow 4R-HE
Red 7B-HE
Blue RR-BB
Green RR-4B
Reactive Black 5
200 mg L−1
28 °C/200 rpm
pH 5.0–6.0
200 mg L−1
26 °C/200 rpm
pH 4.5
100/24 h
Oxidase: MnP, Lac and peroxidase
Martorell et al. (2017a)
T. akiyoshidainum HP2023
Reactive Black 5
100/12 h
Oxidase: phenol oxidase and
peroxidase enzymes
Martorell et al. (2018)
T. beigelii NCIM-3326
Reactive Blue 171
95/24 h
Reductase: NADH-DCIP reduc-Saratale et al. (2009a)
tase and AzoR
P. kudriavzevii CR-Y103
Reactive Orange 16
95/72 h
Reductase: NADH-DCIP reduc-Rosu et al. (2018)
tase and AzoR
C. oleophila
Reactive Black 5
100/24 h
Reductase: AzoR-like
Lucas et al. (2006)
I. occidentalis
Methyl Orange
Orange II
80/15 h
Reductase: AzoR
Ramalho et al. (2004)
G. geotrichum KL20A
Methylene Blue
70/48 h
Biodegradation
Contreras et al. (2019)
C. tropicalis TL-F1
Acid Brilliant Red GR
95/24 h
Biodegradation
Tan et al. (2013)
P. rugulosa Y48
C. kruseia Gl
Reactive Brilliant Red K-2BP
99/24 h
Biodegradation
Yu and Wen (2005)
200 mg L−1
25 °C/250 rpm
pH 4.0
200 mg L−1
25 ºC/250 rpm
pH 7.0
50 mg L−1
37 °C/0 rpm
pH 6.6
400 mg L−1
30 °C/120 rpm
pH 6.0
200 mg L−1
26 °C/120 rpm
pH 7.6
0.2 mM dye
26 °C/120 rpm
Acidic pH
50 mg L−1
35 °C
pH 7.0
100 mg L−1
35 °C/160 rpm
pH 5.0–6.0
200 mg L−1
28 °C/200 rpm
pH 5.0–6.0
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Dyes
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Yeasts species
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Tan et al. (2014b)
Biodegradation
98/10 h
50 mg L
30 °C/200 rpm
pH 6.0
D (%) percentage of decolorization
Magnusiomyces ingens LH-F1 Acid Red B
Enzymatic activity
D (%)/Time
Conditions
Dyes
Yeasts species
Table 1 (continued)
−1
References
Environmental Sustainability
dyes by LiP may involve two consecutive one-electron oxidations of the H2O2-oxidized forms of LiP in the phenolic
ring, where the corresponding carbonium ion carrying the
azo link contributes to the formation of quinone and phenyldiazine through a nucleophilic attack by H2O. The phenyldiazine product is then oxidized by O2 to a phenyl radical
and the azo bond is removed as N2, and the phenyl radical
extracts hydrogen from its environment to produce a stable
aromatic compound (Chivukula et al. 1995). LiP activity
was detected during the biodegradation of various dyes
using Ascomycota yeast strains of Pichia occidentalis (Song
et al. 2018a, b), G. geotrichum (Guo et al. 2019), S. cerevisiae (Jadhav et al. 2007), C. krusei (Charumathi and Das
2011), Diutina rugosa (Bankole et al. 2017), Cyberlindnera
samutprakarnensis (Song et al. 2018a, b), as well as strains
belonging to the Basidiomycota phylum like Trichosporon
laibachii and Trichosporon multisporum (Pajot et al. 2007).
Manganese peroxidase (EC 1.11.1.13) MnP is a glycoprotein enzyme belonging to the oxidoreductase’s family, having a molecular weight ranging from 38 to 62.5 kDa (~ 350
amino acid residue). It is a substrate specific enzyme that
oxidizes Mn2+ to Mn3+, which diffuses from the enzyme
surface and in turn oxidizes the phenolic substrate, such
as lignin model compounds or other organic contaminants
(Zhou et al. 2013). It was reported by Hofrichter (2002), that
MnP production is limited to certain soil litter decomposing and wood-decaying fungus. But some research on the
biodegradation of synthetic dyes using yeast reported the
involvement of MnP in yeast strains of P. occidentalis (L.
Song et al. 2018a, b), Debaryomyces polymorphus, Candida tropicalis (Yang et al. 2008), T. multisporum, and T.
laibachii (Pajot et al. 2007).
Reductases They are enzyme classes that catalyze the
reduction of the azo bonds (–N=N–) of azo dyes to produce a colorless aromatic amine. The involvement of such
enzymes in the biodegradation of dyes was mainly recorded
in bacteria (Hu 2001). However, other studies demonstrated
the biodegradation activity of this enzyme in microalgae
(Sinha et al. 2016), filamentous fungi (Bankole et al. 2018)
and yeast (Lucas et al. 2006). The main reductase enzymes
described in yeast strains are:
Azoreductase (EC 1.7.1.6) AzoR is present in diverse
microorganisms and higher eukaryotes. Although their
structure and function are diverse, they have a common ability to reduce azo bonds of organic compounds including azo
dyes, nitroaromatic and azoic drugs (Misal and Gawai 2018).
The AzoR classification is primarily based on their secondary and tertiary structures. Thus, they can be divided into
two classes: Flavin dependent and independent AzoR. The
flavin-dependent AzoR can also be classified according to
the required co-enzymes NADH, NADPH, or both (Saratale
et al. 2011; Solís et al. 2012). Breaking the azo bond by
AzoR act as a critical step in the dye biotransformation
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Fig. 2 Suggested azo bond
cleavage via a reduction and
oxidation, adapted from Kandelbauer and Guebitz (2005)
Fig. 3 Reactions catalyzed by
NADH-DCIP reductase (a) and
MG- reductase (b)
mechanisms (Elfarash et al. 2017). The involvement of AzoR
during the dye biodegradation was detected in yeast strains
of S. cerevisiae MTCC 463 (Jadhav et al. 2007), C. krusei (Charumathi and Das 2011), Issatchenkia occidentalis
(Ramalho et al. 2004), Trichosporon beigelii NCIM-3326
(Saratale et al. 2009a).
NADH-DCIP reductase (EC 1.6.99.3) The NADH-preferring 2,6-dichloroindophenol reductase is an oxidoreductase
that reduces 2,6-dichloroindo-phenol (DCIP) using NADH
as an electron donor. DCIP is blue in its oxidized state and
becomes colorless after reduction with reductase. This feature confers to this enzyme the ability to be useful in many
clinical settings such as the determination of NADH and
other dehydrogenases with a colorimetric reaction, when
coupled to formazan dye-forming chromogens that act as
hydrogen acceptors (Nishiya and Yamamoto 2007). Some
studies have shown a significant increase in the activities of
NADH-DCIP reductase, during the decolorization of azo
dyes by yeast cells (Bankole et al. 2017; Rosu et al. 2018;
Saratale et al. 2009a; Song et al. 2018a, b).
Malachite green reductase It is an enzyme that also uses
NADH as an electron donor to convert green malachite to
green Leucomalachite. Only few researchers reported (MG)reductase as a marker enzyme for the reduction of synthetic
dyes. The first study showing the role of (MG)-reductase in
the biodegradation of green malachite dye was discovered
in a yeast strain of S. cerevisiae MTCC 463 (Jadhav and
Govindwar 2006). Two years later, Jadhav et al. (2008b)
demonstrated that the biodegradation of methyl red by G.
geotrichum MTCC 1360 strain involved also the (MG)reductase. Thereafter, Charumathi and Das (2011), reported
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that (MG)-reductase activity was increased in C. krusei cells
following the biodegradation of Basic Violet 3. Biradar et al.
(2016) also, reported that the biodegradation of Malachite
green by the S. cerevisiae yeast strain involved an increase
in the enzymatic activity of (MG)-reductase. The reactions
catalyzed by MG-reductase (B) and DCIP-reductase (A) are
reflected in Fig. 3.
The enzymatic biodegradation of dyes using yeast cells
can be improved through omics approaches, especially protein engineering, which would pave the way for further studies. In addition, the exploitation of enzymes from yeast cells
after the dye mycoremediation process may be another form
of valorization of the resulting biomass.
Factors affecting the biodegradation of dyes
by yeast
The effective biodegradation of dye contaminants in wastewater remains a challenging process, due to many factors
that influence this bioprocess. Therefore, it is important to
evaluate and optimize the effect of physicochemical operating conditions, in order to make the process faster, more
efficient and suitable for large-scale applications.
Effects of temperature In the microbial environment, temperature is a critical factor that plays several roles in cell viability, physiological status and metabolic performance. Optimal temperature for yeast growth is typically between 25 °C
and 37 °C (Fu and Viraraghavan 2001). In fact, the optimum
temperature for dye decolorization using yeast cells, is often
correlated with the optimum growth temperature. Changes
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Environmental Sustainability
in temperature can affect the biological response of yeast
cells, especially enzyme activities (Ali 2010). Numerous
studies (Table 1) have reported that the decolorization rate
increases with increasing temperatures (from low to optimum temperature). However, a decolorization decline was
noted at high temperatures (Tan et al. 2013, 2014b, 2016),
which can be attributed to the loss of cell viability, or the
denaturation of enzymes (Saratale et al. 2009b).
Effects of pH It’s known that fungi, including yeast species show greater biodegradation behaviors in acidic or
neutral mediums (Ali 2010). As showed in Table 1, almost
all yeast decolorization studies demonstrated that the bioremoval of dyes is higher at optimum pH between 4 and 7
(Bankole et al. 2017; Jadhav et al. 2008b; Ramalho et al.
2004), and tended to decrease significantly when the pH of
the medium was within the alkaline range. The pH effects
are also related to cell viability, as shown by Peña et al.
(2015), where the cell cycle of S. cerevisiae was interrupted at pH 9.0. This growth inhibition might be attributed to the decrease in cell transport of amino acids or the
incorporation of proteins. On the other hand, the change in
pH of the medium could also inhibit the transport of dye
molecules through the cell membrane, which is considered
as a limiting step for intracellular decolorization (Khan
et al. 2013).
Effects of dye structure and concentration The chemical
structure and concentration of dyes are among the influencing factors of decolonization efficiency. Research studies
have suggested that increasing dye concentration gradually reduces their decolorization (Aksu 2003; Jadhav et al.
2007; Martorell et al. 2018), which could be attributed to
an increase in dye toxicity. Moreover, at high concentrations, dyes can bind to the active site of enzymes mainly of
AzoR, thereby preventing their activity (Jadhav et al. 2008a;
Saratale et al. 2009a). Meanwhile, the chemical structure of
dyes could also influence their degradation. In a comparative
study on monoazo, diazo or triazo dyes, Franciscon et al.
(2012) reported that simple-structured azo dyes with low
molecular weight were easily biodegradable. Additionally,
the azo bond is more susceptible to be broken when the
substituent is in the para position of the phenyl ring relative to the ortho and meta position (Hsueh et al. 2009). For
example, the substitution of electron withdrawal groups
(-SO3H) by (-SO2NH2) in the phenyl ring para position relative to the azo bond, increases the rate of reduction (Walker
and Ryan 1971). Also, amino and hydroxyl-based dyes are
more resistant to degradation than dyes with nitro, methyl,
sulpho, and methoxy groups. This can be explained by the
reduction mechanism that is conducted in two steps: a fast
one-electron transfer reaction occurs to the radical anion,
followed by a second, slower electron transfer process to
create a stable dianion. The functional group of azo bond
with a higher electronic density may not be suitable for this
second electron transfer forming the dianion (Pearce et al.
2003; Rau et al. 2002). Metal-complex dyes might also have
an adverse effect on the decolorization efficiency (Chen et al.
2003; Libra et al. 2004).
Effects of oxygen and shaking There are differing views
on the effect of agitation on decolorization efficiency.
According to results of several studies, the highest decolorization efficiency was observed in agitated cultures compared
to static cultures. Shaking allows aeration to the medium,
and equal distribution of nutrients, it also facilitates the
exchange of gas formed by yeasts during the degradation of
dyes (Martorell et al. 2017b; Pajot et al. 2007; Yang et al.
2008; Yu and Wen 2005). As mentioned above, several oxidative enzymes require oxygen (Shahid et al. 2015). Meanwhile, some researchers obtained the best decolorization
results under static conditions (Table 1). The negative effect
of medium agitation on the decolorization efficiency was
attributed to the fact that oxygen acts as an electron acceptor
with high redox potential, thereby preventing dye reduction
(Li et al. 2004; Kalyani et al. 2008; Al-Tohamy et al. 2020).
Effect of carbon and nitrogen sources Despite the presence of dyes and other organic compounds in textile wastewater, no yeast strain can achieve an effective decolorization
via the biodegradation mechanisms with dyes as the only
carbon source, unless bioremoval is based on biosorption
mechanisms with non-viable yeast biomass (Mahmoud
2016). It was mentioned that the addition of glucose to
yeast culture medium stimulates the biodegradation of dyes
(Chang et al. 2000; Waghmode et al. 2011). Other studies
also showed a strong correlation between glucose depletion
and the rate of dye removal (Lucas et al. 2006; Yang et al.
2008). Glucose can play many roles in metabolic pathways,
including its role as regenerator of the redox mediators like
NADH and FADH, that act as substrates for the production
of H2O2, through an enzymatic reaction by glucose1oxidase
and glucose-2-oxidase (Jafari et al. 2014). In turn, H2O2
acts as a co-substrate for extracellular peroxidase activity
(Swamy and Ramsay 1999). Meanwhile, decolorization in
a medium containing high carbon source concentrations
appears to be less effective. In this case, the yeast cells prefer
to use readily available carbon sources instead of using dyes
as a carbon source (Saratale et al. 2009b). On the other hand,
the supplementation of nitrogen sources to the medium (peptone, yeast extract, urea), can also regenerate NADH (Bras
et al. 2001). However, similarly to carbon, high nitrogen
concentrations may influence the decolorization efficiency.
Tatarko and Bumpus (1998) reported that supplementation
of high nitrogen concentrations to the medium inhibited
Congo red decolorization. Conformingly, a study by Kaushik and Malik (2009), demonstrated that higher nitrogen
concentrations (25–60 mM), suppressed ligninolytic enzyme
13
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Environmental Sustainability
activity. It is therefore important to determine the appropriate concentration of carbon and nitrogen sources that
improve the decolorization process (Pearce et al. 2003; Van
der Zee et al. 2001).
Since several factors can influence the process of biodegradation of dyes by yeast cells, optimizing these factors
is essential to limit losses, whether related to chemicals,
growing conditions, or process time. Applying the design of
the experiment can be a promising solution that can reduce
treatment costs and time, making, therefore, the use of yeast
more competitive than the other treatment processes.
Chemical characterization and toxicity
assessment of dye biodegradation
by‑products
Irrespective of the technique used (physical, chemical,
or biological) in the treatment of textile wastewater, the
main concern is not only the dye removal, but mainly the
guarantee of its effective detoxification after treatment.
Hence, the ecotoxicological evaluation of post-treatment
is crucial. The oxidation processes of synthetic dye using
Fenton reactions, non-metallic oxidation catalysts based
on hydroxyl radicals, and other chemical techniques, can
ensure a successful decolorization (Javaid and Qazi 2019).
However, these techniques can produce various oxidation
by-products that may be more toxic than the original dye
molecules (dos Santos et al. 2007; Sun et al. 2009). In order
to highlight the biodegradation efficiency of dyes from a
toxicological point of view, a detailed characterization of
the metabolites produced during and after complete decolorization must be assessed. Particularly, the colorless aromatic amines resulting from the reduction of the cleavage of
chromophore azo-bonds, because they are classified among
the most hazardous compounds (Forss and Welander 2009;
Puvaneswari et al. 2006). Based on the available literature,
detailed characterization of dye by-product degradation can
be carried out with different chromatography techniques
such as thin-layer chromatography (TLC), high performance liquid chromatography (HPLC), high performance
thin layer chromatography (HPTLC), liquid chromatography–mass spectrometry (LC–MS), gas chromatographymass spectrometry (GC–MS), and spectroscopy techniques
such as ultraviolet–visible spectroscopy (UV–VIS), Fourier
transform infrared spectroscopy (FTIR) and nuclear magnetic resonance (NMR) spectroscopy. In addition, some
studies (Saratale et al. 2009a, b; Tan et al. 2014b) reported
that biodegradation could also be calculated by measuring the level of mineralization using total organic carbon
(TOC), chemical oxygen demand (COD) and biochemical
oxygen demand (BOD). Table 2 summarizes the analytical
technique used for the identification of the by-products of
13
dye biodegradation by yeast species. As showed in Fig. 4
and Table 2, the degradation of the same dye by different
yeast strains produces different by-products. This suggests
the need for ecotoxicological analyzes to ensure the safety
of these metabolites. Indeed, a variety of ecotoxicological
approaches can be used. Among the existing ecotoxicological bioassays, phytotoxicity tests are the most commonly
used, because of their high sensitivity, low cost and easy
use. Saratale et al. (2009a) studied the phytotoxicity of Navy
Blue HER dye and its biodegradation by-products on two
plant models, Sorghum vulgare and Phaseolus mungo. The
results showed a higher germination percentage and significant growth of the plumule and radical in seedlings treated
with degradation by-products compared to those treated with
the original dye, confirming thus the detoxification of HER
by T. beigelii. Similarly, Waghmode et al. (2012a) assessed
the phytotoxicity of Remazol Red and its by-products on
S. vulgare and P. mungo seeds, and demonstrated that the
by-products produced were less toxic compared to the
original dye. Likewise, Tan et al. (2014a) reported that the
investigation of metabolites derived from Acid Orange G
decolorization on P. mungo and Oryza sativa seeds, confirmed the biotransformation of this dye into less toxic byproducts after complete decolorization using C. tropicalis
TL-F strain. The other common bioassay used to estimate
the acute toxicity of the dye and their biodegradation intermediates may be performed using the Microtox test. Tan
et al. (2016) assessed the shift in acute toxicity before and
after biodegradation of Acid Scarlet 3R using Scheffersomyces spartinae TLHS-SF1 strain, with the Microtox test, at
the initial dye concentration of 20 mg L−1and 50 mg L−1.
Moderate (IRs 60% within 30 min) and high (IRs 75% within
30 min) toxicity were recorded against Vibrio fischeri. The
dye treatment toxicity sharply decreased to non-toxic levels.
Similarly, Song et al. (2018a, b) studied the toxic effects of
decolorization intermediates of ARB dye using a yeast strain
of P. occidentalis G1 through Microtox test. It was shown
that the IRs of 20 mg L−1 and 50 mg L−1 were respectively
about 68% and 78%. After dye treatment, the toxicity of
both dye solutions decreased to non-toxic levels as indicated
by the IRs, which decreased to 8% and 11%, respectively.
Likewise, Tan et al. (2019) assessed the toxicity of ARB
(50 mg L−1) and its decolorization by-products produced
by C. tropicalis SYF-1 by using the Microtox test. The IR
was about 75.4% after 5 min exposure, suggesting that the
initial dye concentration possessed high acute toxicity. After
a 6 h treatment, the dye displayed the 5 min IR of about
67.2%, slightly lower than initial IR. However, the 5 min IR
of the dye solution sharply decreased to 9.4% after complete
decolorization (12 h), suggesting a non-toxic level. Also, the
combinations of different bioassays were used to assess the
toxicological effects of synthetic dyes or their metabolites.
Waghmode et al. (2012b) studied the toxicity of Rubine GFL
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Environmental Sustainability
Table 2 By-products detected after the biodegradation of some azo dyes by yeast
Yeast strain
Dye name and chemical
structure
Analyses
C. tropicalis SYF-1
Pichia sp. TCL
Acid Red B
M. ingens LH-F1.
UV–vis
HPLCMS
P. occidentalis G1
S. cerevisiae ATCC 9763
Acid Scarlet 3R
S. spartinae TLHS-SF1
Monoazo disperse dye
C. tropicalis
HNMR
TLC
HPLC
LC-MS
Reactive blue 13
UV-Vis
MS/MS
GC-MS
C. rugopelliculosa
HXL-2
Methyl red
S. cerevisiae ATCC 9763
UV-Vis
FTIR
TLC
ESI-MS
UV-Vis
FTIR
HPLC
Remazol Red
UV-Vis
FTIR
TLC
HPTLC
G. geotrichum MTCC 1360
Rubine GFL
Reactive Black 5
T. akiyoshidainum
HPTLC
HPLC
FTIR
GC-MS
UV–vis
FTIR
HPLCGPC
GC–MS
Reactive green dye
Candida sp. VITJASS
UV–Vis
FTIR
HPLC
GC–MS
Biotransformation products
4-hydrazinylnaphthalene-1- sulfonic acid
4-hydroxynaphthalene-1- sulfonic acid
4-aminonaphthalene-1-sulfonic acid
3,4-dihydroxynaphthalene-1-sulfonic acid
4-hydroxynaphthalene-1,2-dione
4-aminonaphthalene-1-sulfonic acid
naphthalene1,2,3,4-tetraol
3-7-dihydroxy-octahydronaphthalene-2,6-dione
4-aminonaphthalene-1-sulfonic acid
3-amino-4-hydroxynaphthalene-1-sulfonic acid
naphthalen-1-ol
3,7-dihydroxy-hexahydronaphthalene-2,6 (1H,7H)dione
4-amino-naphthalene-1-sulfonic acid
3,4-dihy droxy-naphthalene-1-sulfonic acid
naphthalene-1,2,4-triol
4-Amino-1-naphthalene sulfonic acid sodium salt
3-amino-4-naphthol-1- sulfonic acid sodium salt
Disodium 4-hydroxy-2-[(E)-(4-sulfonato-1-naphthyl)
diazenyl]naphthalene-1-sulfonate
Disodium 4-hydroxy-2-[(E)-(4-sulfonato-1-phenyl)
diazenyl]naphthalene-1-sulfonate
Disodium 4-hydroxy-2-[(E)-(4-sulfonato-1-naphthyl)
diazenyl]benzene-1-sulfonate
4-aminonaphthalene-1-sulfonic acid
7,8-dihydroxynaphthalene-1,3-disulfonic acid,
3,4-dihydroxynaphthalene-1-sulfonic acid
naphthalene-1,2,6,8-tetraol
naphthalene-1,2,4-triol
References
(Tan et al.
2019)
(Qu et al.
2012)
(Tan et al.
2014b)
(L. Song et al.
2018)
(Kiayi et al.
2019)
(Tan et al.
2016)
2-amino-4-methyl- 5-ethoxycarbomylthiazole
(Arora et al.
2005)
1-chloro-3- aniline 2,4,6-triazine
2-chloro-1,3,5-triazine
Phenol
aniline
N, N-dimethyl-p-phenylene diamine (DMPD)
2-Aminobenzoic acid.
N, N’-dimethyl- p-phenylenediamine
2-authentic 2-aminobenzoic acid
3-amino [4, 5 (6-chloro-1, 3, 5 triazine-2yl) amino]
naphthalene 2, 4, 7 benzene trisulfonic acid.
2[(3-aminophenyl) sulfonyl] ethane sulfonic acid.
2- amino naphthalene
N-phenyl-1, 3, 5 triazine
2[(3-aminophenyl) sulfonyl] ethane sulfonic acid
ethylsulfonyl-benzene
2-ethylphenyl sulfone
2-(aminomethyl)-4-nitroaniline
1-(3-nitrophenyl)methana- mine
N-(3-aminopropyl)benzene-1,4-diamine
4-((chlorodifluoromethyl)sulfonyl) aniline
4- ((diazenilfenil) sulfonyl) methyl hydrogen sulfate
5,7-dimethoxy-1-naphthol
5-amino-4-hydroxy-3,6-dioxo-2,3, 5,6tetrahydronaphthalene2,7-sodium disulfonate.
sodium 3-[(dichloro-1,3,5-triazin-2-yl) amino]
benzene-1-sulfonate
sodium 1-amino-4-[(4-aminophenyl) amino]-9,10dioxo-9,10- dihydroanthracene-2-sulfonate
2-chloro-1,1-diphenyethane
Diphenylmethane
(Liu et al.
2011)
(Vatandoostar
ani et al. 2017)
(Jadhav et al.
2008b)
(Waghmode et
al. 2012a)
(Waghmode et
al. 2012b)
(Martorell et
al. 2017b)
(Sinha et al.
2018)
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C.tropicalis SYF-1
(Tan et al. 2019)
4-hydrazinylnaphthalene-1- sulfonicacid
4-hydroxynaphthalene-1- sulfonicacid
4-aminonaphthalene-1-sulfonic acid
3,4-dihydroxynaphthalene-1-sulfonic acid
4-hydroxynaphthalene-1,2-dione
HPLC-MS
C.samutprakarnensisS4
(Song et al., 2018)
Acid Red B
S. cerevisiae ATCC 9763
(Kiayi et al., 2019)
LC-MS
HPLC-MS
Pichia sp. TCL
(Qu et al., 2012)
4-Amino-1-naphthalene sulfonicacid
3-amino-4-naphthol-1- sulfonicacid
4-hydroxy-2-[(E)-(4-sulfonato-1-naphthyl)
diazenyl]naphthalene-1-sulfonate
4-hydroxy-2-[(E)-(4-sulfonato-1-phenyl)
diazenyl]naphthalene-1-sulfonate
4-hydroxy-2-[(E)-(4-sulfonato-1-naphthyl)
diazenyl]benzene-1-sulfonate
Fig. 4 Proposed partial biodegradation pathways of ARB by various yeasts using HPLC–MS and LC–MS analysis techniques
dye and its by-products produced from the biodegradation by
G. geotrichum MTCC 1360 yeast strain. Analytical methods
in this study included genotoxicity and cytotoxicity tests,
oxidative stress, activity of antioxidant enzymes, lipid peroxidation and protein oxidation on root cells of Allium cepa
as well as the phytotoxicity of the dye and dye by-products
using P. mungo and S. vulgare. The outcomes showed that
Rubine GFL dye exerted oxidative stress and subsequent
toxic effects on the root cells, whereas its metabolites were
less toxic. Likewise, Roșu et al. (2019) suggested that after
treatment of BB41 dye by Pichia kudriavzevii CR-Y103,
the degraded metabolites was found to be less toxic than the
parent dye compound, by using phytotoxicity, cytotoxicity,
and genotoxicity assays on Trifolium pratense and Triticum
aestivum seedlings. This indicates the detoxification of this
azo dye.
The chemical characterization of the by-products of dye
biodegradation not only makes it possible to recognize the
pathways of their biodegradation, but may also indicate the
toxicity of the aromatic compounds formed during biodegradation. This will assess the performance of treatment from
a toxicological point of view.
13
Molecular aspects of dye degradation
by yeasts
In all areas of life sciences, the use of omics methods,
namely metagenomics, transcriptomics, proteomics, and
metabolomics, has recently become widespread. The integration of these approaches allows a holistic view of the biological systems. Understanding the molecular mechanisms
of biodegradation of aromatic compounds such as synthetic
dyes is essential for the development of successful bioremediation strategies. Numerous studies have been carried
out on the molecular aspects of dye degradation by fungi
(Mäkinen et al. 2019; Sun et al. 2015) and bacteria (Joshi
et al. 2020; Ma et al. 2019). However, very little research
has focused on the molecular mechanisms involved in the
enzymatic biodegradation of synthetic dyes by yeast cells.
María et al. (2018) unraveled the genetic basis of the enzymatic activities of Mnp, Lac, and Phenoloxidase after complete decolorization of Reactive Black 5 by Trichosporon
akiyoshidainum HP-2023, their genome comprises 30 MB
with a G + C content of 60.75% and 9019 gene models.
Thus, thirty-three putative carbohydrate-active enzymes
with auxiliary activity have been identified in the annotated
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Environmental Sustainability
genome, nineteen hydrogen peroxide-producing enzymes,
four benzoquinone oxidoreductases, four extracellular fungal heme-peroxidases, and two Lacs. For the decolorization
of ARB at high osmotic environment, Wang et al. (2020)
reported that the halotolerance enhancement through the
detection of different genes expressed in P. occidentalis used
for the decolorization of ARB was related to the upregulated genes encoding the enzymes or functional proteins
related to intracellular synthesis of glycerol. Also based on
the transcriptome sequencing, Tan et al. (2020) suggested
that the halotolerance capacity of C. tropicalis SYF-1 yeast,
used for the decolorization of ARB was enhanced by the
regulation of the cell wall component. Some recent studies
have focused on enhancing the biodegradation of dyes by
improving enzyme activity through protein engineering. For
example, Zhang et al. (2019) have reported that the recombinant enzyme rMnP3-BBP6 that resulted from cloning of new
MnP gene (mnp3) of the white-rot fungus Cerrena unicolor
BBP6 in Pichia pastoris, has shown a high decolorization
activity of various dyes: Brilliant blue R, Methyl orange,
Crystal Violet, Bromophenol Blue, and Remazol Brilliant
Blue. Karla et al. (2020) have used saturation mutagenesis
to alter two amino acids in the catalytic tryptophan environment positions V160 and A260, of Versatile peroxidase (VP)
from Pleurotus eryngii. In order to effectively use VP and its
variants for the degradation of azo dyes (Evans Blue, Amido
Black 10B, and Guinea Green B), VP was immobilized on S.
cerevisiae EBY100 cell surface and cell wall fragments were
used after lysis. Thus, the embedded VP retained ∼ 70% of
its initial activity after 10 cycles of decolorization.
State-of-the-art technology in the biochemical characterization of dye biodegradation pathways coupled with further
studies on genomic, proteomic, and metabolomic aspects,
could improve fundamental knowledge in the mycoremediation field. This could offer a wide range of possibilities for
improving the performance of yeasts or their enzymes for
applications in the bioremoval of toxic dyes or other contaminants from wastewater.
Future perspectives
Despite the scientific and technical advancements that have
been developed for the depollution of textile wastewater,
there is no universal approach to achieve effective detoxification with low-cost investment. Each technique has its
own advantages and limitations. Therefore, the use of hybrid
and/or integrated processes may be a promising solution.
In fact, mycoremediation can be an efficient approach to be
combined with other chemical or physical processes (Akhtar
et al. 2020). In order to promote such bioprocesses and make
them more attractive, the first crucial concern is related to
the choice of tolerant yeast strains. Furthermore, optimizing
operating conditions such as agitation, temperature, and the
use of low-cost substrates as carbon and nitrogen sources
can significantly reduce the processing cost. From the point
of view of environmental sustainability, the chemical characterization of the by-products of the enzymatic biodegradation of dyes by yeast cells and the evaluation of their
toxicity after complete decolorization will highlight the
environmental safety of this bioprocess compared to other
chemical dye-degradation processes. Thus, the combination
of characterization studies of biodegradation by-products
with further studies based on omics approaches, notably
transcriptomics, proteomics and metabolomics techniques,
will greatly contribute to the understanding of dye biodegradation pathways by yeast cells (Wang et al. 2020; Zheng
et al. 2020). Also, the use of yeast oxidoreductive enzymes
in the bioremediation of dyes, and the improvement of their
performance in terms of stability, selectivity, and catalytic
activity, can be carried out with various genetic engineering
techniques (Karla et al. 2020). Indeed, over the past decade,
research has become increasingly interested in the application of genetic engineering techniques to develop enzyme
formulation for bioremediation applications. Meanwhile,
the fate of the biomass after the biodegradation of dyes
is among the main questions that needs to be solved. The
resulting biomass can be valued in several areas, mainly in
the energy sector, for the production of biodiesel (Ali et al.
2021). The mycoremediation of synthetic dyes using yeast
is simple, reliable, and cost-efficient at laboratory scales,
however approval of this approach should be carried out
on pilot schemes prior to large-scale industrial operations.
Conclusion
Based on the studies reviewed in this review, it can be concluded that mycoremediation using yeast cells can be an
efficient and eco-friendly strategy for the treatment of toxic
dyes. The biodegradation ability of yeasts has been related to
the activity of oxidases (Lac, Tyr, LiP, and MnP), and reductases (AzoR, NADH-DCIP reductase and MG-reductase).
The operating physicochemical parameters that influence the
biodegradation process have to be optimized for successful
decolorization. For most yeast strains, the highest bioremoval capacity is achieved at neutral or acidic pH ranges, in
shaken cultures at 30 °C. The addition of appropriate supply of nitrogen or carbon sources significantly improves the
dye-removal efficiency. Meanwhile, textile dyes can affect
the biodegradation capacity of yeast, both by their chemical properties and/or concentration. During the process of
dye biotransformation, the chemical characterization of
the degradation intermediates using chromatography and
spectroscopy techniques enables the identification of the
dye degradation pathways. Besides, toxicological bioassays
13
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Environmental Sustainability
revealed that the resulting intermediate metabolites of dye
biodegradation using yeast are typically less toxic than the
initial dye. We can therefore deduce that the biodegradation of synthetic dyes by yeasts does not only allow dye
decolorization, but ensures the effective detoxification of
these contaminants and their by-products. This makes the
use of yeast cells or their enzymes in the biodegradation
of synthetic dyes a promising future technology to ensure
environmental sustainability.
Acknowledgements The authors gratefully acknowledge Moroccan
Foundation for Advanced Science, Innovation and Research (MAScIR) for the financial and technical support. Authors wish also to thank
Chanda Mutale Joan for reviewing the quality of the English language
in this article.
Compliance with ethical standards
Conflict of interest On behalf of all authors, the corresponding author
states that there is no conflict of interest.
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Affiliations
M. Danouche1,2
1
· H. EL Arroussi1 · N. El Ghachtouli2
Green Biotechnology Center, Moroccan Foundation
for Advanced Science, Innovation and Research (MAScIR),
Rue Mohamed Al Jazoulie Madinat Al Irfane, 10 100 Rabat,
Morocco
13
2
Microbial Biotechnology and Bioactive Molecules
Laboratory, Faculty of Sciences and Technologies, Sidi
Mohamed Ben Abdellah University, Route Immouzer,
P. O. Box 2202, Fez, Morocco