azo dyes removal by bacteria isolated from industrial wastewater

AZO DYES REMOVAL BY BACTERIA ISOLATED FROM

INDUSTRIAL WASTEWATER

Mehvish Ajaz

M.Phil (Punjab)

Department of Microbiology and Molecular Genetics

University of the Punjab, Lahore,

Pakistan

AZO DYES REMOVAL BY BACTERIA ISOLATED FROM INDUSTRIAL WASTEWATER

A Thesis Submitted to the University of the Punjab, in Partial Fulfillment

of the Requirements of the Degree of

Doctor of Philosophy

By

Mehvish Ajaz

JANUARY, 2019

Department of Microbiology and Molecular Genetics

University of the Punjab, Lahore,

Pakistan

Dedicated To

MY PARENTS

for the love and untiring efforts to make my life successful

Acknowledgement

Praise to the grace of ALLAH ALMIGHTY, The Gracious, and The Merciful. He bestowed

man with intellectual power and understanding and gave him spiritual insight enabling him to

discover his “Self” know his Creator through His wonders and conquer nature. I bow in

obedience before my Lord Who bestowed upon me His love and Blessings and made it possible

for me to accomplish this task and get a drop of experience from already existing ocean of

knowledge with label of being called as a “Student of Research”.

Next to all His Messenger HAZRAT MUHAMMAD (PEACE BE UPON HIM) who is an

eternal torch of guidance and knowledge for humanity.

It is indeed a pleasure and honor for me to express my deepest gratitude towards my

research supervisor Dr. Abdul Rehman, Assistant professor, Department of Microbiology and

Molecular Genetics, University of the Punjab, Lahore. What to say about his knowledge, advices,

guidance and patience throughout my work. His intellectual, hard work, patience and caring

nature are just few of many qualities which had motivated me to complete this research task. I

regard myself lucky to be under the supervision of such a humble and nice person from which I

learned a lot as a researcher and more as a human.

I also want to say word of thanks and deepest appreciation to Prof. Dr. Shahida Hasnian,

(Former Chairperson MMG) and Dr. Anjum Nasim Sabri (Chairperson, Department of

Microbiology and Molecular Genetics, University of the Punjab, Lahore) for providing me an

opportunity to work in the department and for her invaluable guidance and motivation which

made this research project possible.

I also want to say thanks to all the teachers of the department for their cooperation and

help in the completion of this research work. I would like to express special word of thanks to

my seniors Ms. Amina Elahi and Dr Sidra Ilyas for their supreme support, help and making my

time so special, and all the research fellows Ms Hafsa Shahbaz, Ms Wajiha Kanwal, Ms Asma

Akhtar, whose kind behavior and guidance offered great help and encouragement throughout

my research work

I am also extremely thankful to the technical staff MMG, our lab attendant, Mr Nadeem,

Mr. Shehbaz, Mr Mazhar and Mr Saif for their incredible support and cooperation.

Last but not least, I must acknowledge the most important persons in my life, which are,

my family members. I would like to thanks to my entire family for their support and inspiration

they have given me throughout my life. My sincere thanks to all those not mentioned here but

who contributed in one way or other throughout my life.

MEHVISH AJAZ

\

CONTENTS

Title Page No.

List of Tables i

List of Figures ii

Summary vii

Introduction 1

Dyes 3

Classification of dyes(s) 6

Azo dyes 8

Mechanisms of azo dye decolorization by microorganisms 10

1. Biosorption 10

2.Enzymatic degradation 11

Parameters influencing dye degradation 13

1.Structure of azo dyes 14

2. Carbon and nitrogen source 15

3. Salinity, dye concentration, pH, temperature and oxygen 16

Biological systems involved in the decolorization of azo dyes 17

Azo dyes degradation by algae 18

Azo dyes degradation by yeast 19

Azo dyes degradation by filamentous fungi 20

Azo dyes degradation by bacteria 21

Azo dyes degradation by using genetically modified microorganisms and enzymes 22

Azo dyes degradation by using consortia 23

Condition of dye pollution in Pakistan 24

Aims and Objectives 25

Material and Methods 26

Sample collection 26

Sample enrichment 26

Dyes 26

Isolation of dye degrading bacteria 26

Evaluation of dye degrading potential 27

Morphological and biochemical characterization of bacterial isolates

27

Determination of optimum growth conditions of bacterial isolates 28

Molecular characterization of bacterial isolates 29

DNA isolation 29

Agarose gel electrophoresis 30

PCR amplification 31

Gene cleaning 31

Sequencing and submission of sequences to GenBank 31

Construction of phylogenetic tree 32

Optimization of biodegradation conditions 32

Decolorization of multiple dyes 33

Growth decolorization relationship 33

Consortium studies 34

Development of consortium 34

Aerobic consortium 34

Anaerobic consortium 34

Mixed consortium 34

Analysis of dye degraded products 35

Sample Preparation 35

Thin layer chromatography (TLC) 35

High performance liquid chromatography (HPLC) 35

Fourier transform infrared spectrometer (FTIR) 36

Gas Chromatography Mass Spectrometry (GC-MS) 36

Dye degradation at large scale 36

Microbial toxicity 37

Phytotoxicity 37

Statistical analysis 37

Results 38

Physiochemical characteristics of wastewater samples 38

Isolation and evaluation of azo dye degrading potential of bacteria 39

Morphological and biochemical characterization of bacterial isolates 39

Determination of optimum growth conditions of bacterial isolates 43

Optimum temperature 43

Optimum pH 43

Growth curves 43

Molecular characterization of bacterial isolates 49

Optimization of bio decolorization conditions 54

Decolorization of multiple dyes 55

Growth Decolorization relationship 55

Consortium studies 66

Analysis of dye degraded products 67

TLC 67

HPLC 68

FTIR 68

GC-MS 75

Dye degradation at large scale 79

Microbial toxicity 83

Phytotoxicity 85

Discussion 89

References 103

Publications Annex I

HPLC chromatograms Annex II

i

LIST OF TABLES

Table No. Description Page No.

Table 1 Physiochemical parameters (temperature, pH) of wastewater collected

from industrial area of Sheikhupura and Lahore.

38

Table 2 Decolorization (%) of bacterial isolates, isolated from 9 samples. 40

Table 3 Morphological and biochemical characteristics of bacterial isolates. 42

Table 4 The nearest homolog and their percentage homology with the azo dye

degrading bacterial strains after 16S rRNA sequencing.

51

ii

LIST OF FIGURES

Figure No. Description Page No.

Figure 1 Schematic diagram showing various treatment methods for textile

effluents.

3

Figure 2 Structure of Synozol Red HF-6BN (CI Reactive Red 195). 4

Figure 3 Chemical structure and some characteristics features of major textile

dyes.

5

Figure 4 Classification scheme of dyes. 7

Figure 5 Proposed mechanism for degradation of azo dyes by azo reductase. 13

Figure 6 Change in color of Synazol Red HF-6BN and Sumifex Tourqi blue as

compared to the control in flask containing culture of bacterial strain

(a) 1b (b) 1m (c) 2p (d) 4v (e) 3c (f) 3d (g) CBL-2 isolated from the dye

contaminated wastewater.

41

Figure 7 Growth of bacterial strains at various temperatures after incubation of

16 hours (a) anaerobic and (b) aerobic conditions.

44

Figure 8 Growth of bacterial isolates at various pH after incubation of 16 hours

in (a) anaerobic and (b) aerobic conditions.

45

Figure 9 Growth curves of bacterial isolates (a) 1b, (b) 1m, (c) 2p, (d) 4v, (e) 3c,

(f) 3d and (g) CBL-2 in LB medium at their respective optimum

temperatures. Optical densities were taken at 600 nm after regular time

interval.

49

Figure 10 Lane M, DNA marker (GeneRuler 100bp DNA ladder #SM0323

Fermentas); Lanes 1b, 1m, 2p, 4v, 3c, 3d, and CBL-2 represent bacterial

isolates. The gel is 1% agarose and stained with ethidium bromide.

50

Figure 11 Lane M, DNA marker (GeneRuler 100bp DNA ladder #SM0323

Fermentas); Lanes 1b, 1m, 2p, 4v, 3c, 3d, and CBL-2 represent

bacterial isolates. The gel is 1% agarose and stained with ethidium

bromide.

50

iii

Figure 12 Phylogenetic tree of all the selected bacterial isolates based on neighbor

joining method (a) S. lentus 1b, (b) S. lentus 1m, (c) B. subtilis, (d) S.

lentus 4v, (e) A. aquatilis and (f) A. faecalis.

53

Figure 13 Phylogenetic tree of all the selected bacterial isolates based on neighbor

joining method in combined form.

53

Figure 14 Optimization of decolorization condition for S. lentus 1b (a)

temperature, (b) pH, (c) incubation condition, (d) inoculum percentage,

(e) carbon source, (f) nitrogen source and (g) dye concentration.

56

Figure 15 Optimization of decolorization condition for S. lentus 1m (a)



57

Figure 16 Optimization of decolorization condition for B. subtilis (a) temperature,

(b) pH, (c) incubation condition, (d) inoculum percentage, (e) carbon

source, (f) nitrogen source and (g) dye concentration.

58

Figure 17 Optimization of decolorization condition for S. lentus 4v (a)



59

Figure 18 Optimization of decolorization condition for A. aquatilis 1b (a)



60

Figure 19 Optimization of decolorization condition for A. faecalis 1b (a)



61

Figure 20 Potential of (a) S. lentus 1b, (b) S. lentus 1m, (c) B. subtilis, (d) S. lentus

4v, (e) A. aquatilis and (f) A. faecalis to decolorize multiple dyes

incubated at 37°C after suitable time interval.

63

Figure 21 Effect of decolorization on growth of (a) S. lentus 1b, (b) S. lentus 1m,

(c) B. subtilis, (d) S. lentus 4v, (e) A. aquatilis and (f) A. faecalis under

optimum growth conditions.

65

iv

Figure 22 Effect of (a) Aerobic, (b) Anaerobic and (c) Mixed consortium on

degradation of dye.

66

Figure 23 TLC chromatogram of extracted metabolites of (a) S. lentus 1b, (b) S.

lentus strain 1m, (c) B. subtilis, (d) S. lentus strain 4v, (e) A. aquatilis

and (f) A. faecalis decolorized dye sample visualized under UV range

of 254nm.

67

Figure 24 HPLC analysis of reactive Synazol red HF-6BN with a mobile phase of

profile of methanol at 1.0 mL/min. The column was C-18 (250×4.6

mm) (a) represents dye chromatogram while (b) represents S. lentus 1b,

(c) S. lentus strain 1m, (d) B. subtilis, (e) S. lentus strain 4v, (f) A.

aquatilis and (g) A. faecalis degraded dye products extracted after 4

days of incubation at 37°C.

72

Figure 25 FTIR spectrum of control dye (Synazol red HF-6BN) with (a) S. lentus

1b, (b) S. lentus 1m, (c) B. subtilis, (2p) (d) S. lentus 4v, (e) A. aquatilis

(3c) and (f) A. faecalis (3d).

75

Figure 26 GC-MS chromatograms of (a) control dye (b) S. lentus 1b, (c) S. lentus

1m, (d) B. subtilis, (2p) (e) S. lentus 4v, (f) A. aquatilis (3c) and (g) A.

faecalis (3d).

79

Figure 27 Change in color of dye with respect to control in flasks containing

culture of bacterial isolates (a) S. lentus 1b, (b) S. lentus strain 1m, (c)

B. subtilis (2p), (d) S. lentus strain 4v, (e) A. aquatilis (3c) and (f) A.

faecalis (3d) from the original industrial wastewater.

80

Figure 28 Decolorization of dye (Synazol red HF-6BN) by bacterial isolates (a) S.

lentus 1b (b) S. lentus strain 1m, (c) B. subtilis (2p), (d) S. lentus strain

4v, (e) A. aquatilis (3c) and (f) A. faecalis (3d) from industrial

wastewater after incubation of 2, 4, 6 and 8days of incubation at room

temperature.

82

v

Figure 29 The broth decolorized by S. lentus 1b used for incubation of (a) B.

megatarium, (b) B. subtilis and (c) B. cereus at 37°C for 48 hours. No

zone of inhibition was appeared in any case.

83

Figure 30 The broth decolorized by S. lentus 1m used for incubation of (a) B.



83

Figure 31 The broth decolorized by B. subtilis used for incubation of (a) B.



84

Figure 32 The broth decolorized by S. lentus 4v used for incubation of (a) B.



84

Figure 33 The broth decolorized by A. aquatilis used for incubation of (a) B.



84

Figure 34 The broth decolorized by A. faecalis used for incubation of (a) B.



85

Figure 35 The broth decolorized by Alishwanella sp. used for incubation of (a) B.



85

Figure 36 Use of treated wastewater by (a) S. lentus 1b, (b) S. lentus strain 1m, (c)

B. subtilis (2p), (d) S. lentus strain 4v, (e) A. aquatilis (3c) and (f) A.

faecalis (3d)., tap water and untreated wastewater for growth of Vigna

radiata (mung beans) for a 10 days incubation period under 1:1 light

and dark conditions.

88

Figure 37 Synazol Red HF 6BN dye enters into the cell (A. aquatilis 3c) by

unknown mechanism. Upon entrance into the cell the dye is

enzymatically processed into various end products. Most probably, azo

group (-N=N-) is reduced, followed by cleavage reaction and different

100

vi

end products are produced. Secondly, desulfonation and oxidative

deamination results in synthesis of pyrrolo[1,2-a]pyrazine-1,4-dione

derivative which can be used as substrates in amino acid metabolism.

The amino acid catabolism can synthesize pyruvate (3C compound)

which can be converted into acetyl-CoA. The acetyl-CoA undergoes

Krebs cycle to produce NADH2 and FADH2 (substrates of electron

transport chain). Moreover, dye desulfonation, oxidative deamination

and carboxylation lead to produce phthalate derivatives, which can be

transformed into different fatty acids and aldehydes. The phthalate,

fatty acids and aldehydes can directly/indirectly enter into fatty acid

oxidation reactions (β-oxidation) to produce acetyl-CoA, NADH2 and

FADH2.

Figure 38 Synazol Red HF 6BN dye enters into the cell (Staphyloccocus lentus 1M) by

unknown mechanism. Upon entrance into the cell the dye is enzymatically

processed into various end products. Most probably, azo group (-N=N-) is

reduced, followed by cleavage reaction and different end products are

produced. Secondly, desulfonation and oxidative deamination eventually

results in synthesis of 4-guanidinobutanoic acid derivative which can be used

as substrates in amino acid metabolism. The amino acid catabolism can

synthesize pyruvate (3C compound) which can be converted into acetyl-CoA.

The acetyl-CoA undergoes Krebs cycle to produce NADH2 and FADH2

(substrates of electron transport chain). Moreover, dye desulfonation,

oxidative deamination and carboxylation lead to produce phthalate

derivatives, which can be transformed into different fatty acids and aldehydes.

The phthalate, fatty acids and aldehydes can directly/indirectly enter into fatty

acid oxidation reactions (β-oxidation) to produce acetyl-CoA, NADH2 and

FADH2.

101

Figure 39 Flow diagram demonstrating the microbial treatment of textile effluent

for dye degradation and use of treated wastewater for the growth of

plants.

102

vii

SUMMARY

The present study deals with the isolation and characterization of dye degrading bacteria

from industrial effluent. Based on dye decolorization percentage seven bacterial isolates i.e., 4

anaerobic (1b, 1m, 2p and 4v) and 3 aerobic (3c, 3d and CBL-2) were selected for further study.

The isolates were recognized as Staphylococcus lentus 1b (KX950707), Staphylococcus lentus 1m

(KX950708), Bacillus subtilis (KX950709), Staphylococcus lentus 4v (KX950710), Alcaligenes

aquatilis (KY009932), Alcaligenes faecalis (KY009933) and Alishwanella sp. (JN546609) on the

basis of morphology, biochemical characterization and 16S rRNA ribotyping.

All the bacterial isolated had varying ability to decolorize dye. The dye degrading

percentage of S. lentus 1b, S. lentus 1m, B. subtilis, S. lentus 4v, A. aquatilis, A. faecalis and

Aliswanella sp. were 71, 70, 73,79,82, 78 and 76 respectively. All the isolates showed optimal

growth at 37 oC and pH7.

The dye decolorizing conditions of the isolates were optimized in order to get optimum

rate of decolorization. All the strains showed maximum decolorization rate at 37 oC and pH 7. The

shaking anaerobic conditions were best to decolorize dye for aerobic isolates while anaerobic one

decolorize dye optimally at static conditions. S. lentus 1b, S. lentus 1m and S. lentus 4v gave

maximum decolorization rate when wheat bran was used as carbon source while B. subtilis

decolorize dye optimally in the presence of sugarcane baggase. A. aquatilis and A. faecalis relied

more on saw dust as carbon source. As nitrogen source yeast extract was utilized by all the strains

to decolrize dye at maximum rate. S. lentus 1b, B. subtilis, A. aquatilis and A. faecalis decolrized

dye efficiently when inoculum percentage was 6 while S. lentus 1m and S. lentus 4v at 4 percent.

All the strains gave maximum decolorization rate when dye concentration was 10 mg/L except S.

lentus 4v whose optimum dye concentration was 20 mg/L.

viii

The potential of the strains to degrade multiple dyes was also evaluated. S. lentus 1b, S.

lentus 1m and S. lentus 4v, A. aquatilis and A. faecalis decolorized mixture of multiple dyes

(Congo Red, Methyl Red, Phenol Red, Brilliant Black and Navy Blue) at a rate of 81, 79, 82, 87,

86 and 82 percent respectively. The effect of dye decolorization was also monitored on the growth

of bacteria. It was observed that the dye decolorization process did not affect the bacterial growth

as bacteria prosper to grow.

Dye degrading potential of bacteria was also evaluated in the form of consortium. The rate

of decolorization was increased upto 8-15% when aerobic consortium was used in comparison to

aerobic bacteria while an increase of 10-14% was observed in decolorization rate when consortium

of anaerobic strains was utilized for degradation. When aerobic and anaerobic consortia were used

in combination an increase of 8-10% was observed in decolorization rate in comparison to

individual aerobic and anaerobic consortium.

The analysis of dye degrading products was done by using various techniques. TLC was

done in order to confirm that the dye was degraded into its products. The results showed that the

Rf value of bands of obtained in the dye degraded products (0.89, 0.90, 0.94, 0.95, 0.98) were

different to that of control dye (0.94)

HPLC was also performed to confirm the degradation of dye into its metabolites. Peaks of

different retention time were observed in the control and dye degraded samples. The retention time

of peaks in control sample were 1.80, 2.88 and 5.00 while the retention time of peaks given by

degraded sample of S. lentus 1b (1.97 and 2.92), S. lentus 1m (3.01), B. subtilis (1.92, 2.89 and

4.86), S. lentus 4v (1.94, 2.91, 4.92 and 6.37), A. aquatilis (1.99, 2.30, 2.95 and 3.76) and A.

faecalis (1.93, 2.09 and 2.95).

ix

The comparison of FTIR spectrum showed that the peaks of wavelength 1612 and 1532

cm-1 which are due to the presence of -N=N- stretching were present in the control dye were not

observed in the spectrum of degraded sample. The presence of amines were confirmed in the

degraded samples by the peaks of wavelength 1104, 1108 and 1037 cm-1. This confirmed that the

degradation of dye into dye into amines.

The identification of dye degraded products was done with GC-MS. The results showed

that the predominant compounds found in the final dye treated sample were 3-Aminobutanoic acid,

pyrrolo pyrazine-1, 4-dione and palmitic acid. The compounds could become the part of the energy

yielding metabolic processes.

The dye degraded wastewater was checked for its microbial toxicity as well as

phytotoxicity. It was noted that bacterial-treated wastewater was not toxic for the growth of

microorganisms. Thus, this purified water could possibly be used for irrigation of agricultural

crops. Multiple dyes degradation potential of the bacterial strains makes them ideal candidates to

be utilized for the bioremediation of azo-dyes contaminated sites to reclaim the environment, thus

laying an impending foundation for green chemistry to exterminate the toxic levels of

environmental azo-dyes.

1

INTRODUCTION

Industrialization acts as a backbone for the growth and development of a country. This

rapid increase in industrialization enhances the production and utilization of various chemicals in

day to day life (Sathiya et al., 2007; Saharan and Ranga, 2011). It increases the standard of living

but despite of all its beneficial effects it imposes hazardous effects on the environment as well.

The discharge of toxic effluents from the industries becomes a source of causing environmental

pollution. It contaminates the aquatic as well as terrestrial ecosystem.

The industrial pollutants mainly include organochloride-based water pesticides, heavy

metals (lead, mercury, cadmium, chromium etc.) insecticides, herbicides, oil, cosmetics, fertilizers,

chemical wastes, chlorinated solvents, hydrocarbons, colors and dyes. Synthetic dyes are

extensively used in textile, paper production, leather tanning, agriculture, food technology, light

harvesting array, pharmaceuticals and coloring which make them an important class of industrial

chemicals (Dafale et al., 2010). The annual production of synthetic dyes is 0.7 million tonnes

worldwide and 10,000 various pigments and dyes are used in industries. (Rafi et al., 1990).

The wastewater that is discharged from textile industries becomes a source of pollution due

to presence of dye content in it. About 10-15% of dyestuff is lost directly into wastewater due to

inefficiency in dyeing process. (Spadarry et al., 1994). The wastewater is appeared colored even

when the concentration of dye is 1 mg / L, while the concentration of dye is 10 – 200 mg/ L in

textile processing wastewater which results in decrease in water quality. (O ’Neil et al., 1999). The

removal of dyes from textile effluent is very difficult and thus it becomes a major source of

pollution (Oh et al., 2004; Meerbergen et al., 2018).

The most widely used classes of synthetic dyes in industry include azo, anthraquinone and

triarylmethane dyes. Azo dyes are the most widely used group among these classes and their usage

2

is about 70% of all the organic dyes (Stolz, 2001). They make up the vast majority of the dyes

discharged by textile processing industries. Azo dyes are xenobiotic in nature and are resistant to

process of biodegradation. They pose toxicity, genotoxicity, mutagenecity and carcinogenicity to

aquatic organisms (algae, fish, and bacteria) as well as for animals (Ollgaard et al., 1999; Stolz,

2001). Elevated levels of azo dyes accumulated in the environment and this is because of resistant

nature of azo bond present in these dyes. They have the potential to stabilize in both acidic and

alkaline environments as well as in extreme temperature conditions.

Most of the dyes are toxic in nature and carcinogenic as well (Acuner and Dilek, 2004).

Their disposal into water bodies poses severe damage to hydrophytes by decreasing their

photosynthetic activity as light is unable to pass through from colored content (Aksu et al., 2007)

and their breakdown products poses a serious threat to aquatic organisms (Hao et al., 2000).

Physical and chemical methods such as adsorption, coagulation- flocculation, oxidation and

electrochemical methods can be used for the elimination of dyes from wastewater (Fig. 1) (Lin

and Peng, 1994, 1996). But these processes are very expensive and produce by products which

need further processing. These defects can be removed by using bio processing techniques as they

are cost effective and environmental friendly as well.

Many microorganisms have ability to degrade dyes such as fungi, algae, yeast,

actinomycetes and bacteria (Chen et al., 2003). Bacteria can degrade dyes both under aerobic and

anaerobic conditions (Banat et al., 1996). Many bacteria and fungi such as Aeromonas,

Pseudomonas, Bacillus, Rhodococcus, Shigella, Klebsiella, Rhizopus oryzae, Penicillium

oxalicum and Phanerochaete chrysosporium have the ability to biodegrade azo dyes efficiently

(Chung et al., 1993; Banat et al., 1996; Wong and Yuen, 1996; Zissi et al., 1997; Chang et al.,

3

2001). Bacteria biodegrade azo dyes by producing enzymes such as azo reductase and laccase

while fungi tolerate dyes by absorption.

Figure 1: Schematic diagram showing various treatment methods for textile effluents.

Dyes

The organic compounds which are used for giving color to different products such as paper,

textile, cosmetics, waxes, plastics, pharmaceuticals, and so on are called dyes. Colorants are of

two types i.e., dyes and pigments. The differ in two properties i.e., solubility and interaction with

the substrate. Dyes are soluble in water while pigments are not. Dyes diffuse into the material

while pigments do not interact with the substrate. Dyes are usually prepared by the addition of

different functional groups such as nitro, chloro, amino, bromo etc. to basic organic chemicals e.g.,

benzene, anthracene (Fig. 2 and 3) (Ramanath, 2005).

Dyes have been used by mankind for centuries and the earliest evidence of dye utilization

is thought to be about 1,80,000 years ago by Neanderthal man (Christie, 2001). All the dyes were

more or less natural, till the late nineteenth century, with chief sources like leaves, vegetable

extracts, twigs, berries, roots, flowers of several plants, mollusks insects. Natural dyes did not have

a wide range of colors were usually produced on a small scale (Gupta, 2009).

4

Nevertheless, in 1856 mauve dye being the world’s first commercially effective synthetic

dye was discovered by W.H. Perkin coincidentally (Hunger, 2003). We can define such dyes as

colored substances that give a permanent color to fibers when applied and resist to disappearing

upon revelation to light, perspiration, water, various chemicals, plus oxidizing agents as well as

microbes’ attack (Rai et al., 2005). Synthetic dyes, due to their cost effectiveness and vast variety

of novel colors, have substituted the natural dyes (Gabriel and Hong, 2008).

Electromagnetic energy is absorbed by all aromatic compounds, but those compounds

appear colored which absorb energy in visible wavelength region i.e., 350-700nm. There are two

groups in dyes. First one is chromophoric group in which conjugated double bonds are present and

it is decolorized electron system. Second group is called auxochromes which is responsible to

enhance the color of chromophoric group. Chromophores include functional group –C=C–, –

C=N–, –C=O, –N=N–, –NO2 and quinonoid rings while in auxochromes –NH2, –SO3H, –COOH

and –OH (Christie, 2001). Classification of dye is mainly based on the type of chromophore

present in them or on their use. There are 20-30 different types of dyes depending on their structure.

The production of dyes is in million tonnnes used for dyeing and printing.

Figure 2: Structure of Synozol Red HF-6BN (CI Reactive Red 195)

5

Figure 3: Chemical structure and some characteristics features of major textile dyes (Suteu et al.,

2011).

6

Classification of dye(s)

The basis of classification of dyes includes their chemical structure and color index (CI).

The mono-azo, di-azo, tri-azo, anthroquinone, triarylmethane, and phthalocyanines are the

common classes of dyes. They may be anionic, cationic, and non-ionic on the base of their use in

industry (Fig. 4). They can also be subclassified on the base of color into yellow, orange, red,

violet, blue, green, and black (Ramanath, 2005). Kirk-Othmer (1979), classified dyes into

following categories:

Acid dyes: These are anionic dyes which are soluble in water. They have one or more sulphonic

and carboxylic acid group in their structure and chemically constituted by compound azo,

anthraquinones and triarylmethanes, iminoacetone, nitro, nitrous and quinoline. They are widely

used to dye nylon, silk, modified acrylic, wool, paper, food and cosmetics.

Basic dyes: They are cationic dyes which are also soluble in water and produce cationic

compounds. These are chemically constituted by compounds azo, anthraquinone, triarylmethane,

methane, thiazine, oxazine, acridine and quinoline. They are used to dye modified acrylic,

modified nylon, modified polyesters and papers. They have some biological activity as well as

they are used in medicines and antiseptics.

Direct dyes: They are anionic compounds which are soluble in water when electrolytes are present

in the solution. These are chemically composed of azo compounds, with thiazoles, phtalocyanines

and oxazines. They are usually used in dyeing of cotton, regeneration of cellulose, for dyeing

paper, leather, and nylon.

Fluorescent dyes (group of the xanthenes): They are actually colorless compounds and appeared

fluorescent due to absorbtion of incident UV light and re-emittance in the visible region (blue) of

7

the spectrum. They are not actually dyes in nature but due to their wide use in fabrics and other

materials they are classified under the heading of dyes by Color Index.

Reactive dyes: They have a simple chemical structure. They are constituted chemically by azo

compounds, anthraquinones and phtalocyanines. They have high fixing property due to the

formation of covalent bridges with the hydroxyl group of cellulose within the fiber.

Sulphurous dyes: This is usually a small group of dyes with low cost and good fixing nature.

They are usually used to dye cotton after alkaline reduction bath with sodium sulphite.

Vat dyes: They are usually insoluble in water and mostly used to dye cellulosic fibers after the

treatment of alkaline bath with sodium hydrosulphite. They are chemically anthraquinones and

indigo in nature.

Dye precursors: These are the dyes which are obtained from raw materials. They are cyclic

aromatic compounds having benzene ring in their structure and develop color by chemical

reactions.

Figure 4: Classification scheme of dyes (Uday et al., 2016).

8

Azo dyes

The simple synthesis method of azo dyes makes them the biggest and most widely used

group of dyes in industry. A German scientist P. Gries started their production in 1858 when the

reaction mechanism of diazotization for the production of azo compounds was discovered by him.

Out of the total global production of dyes, which was nearly 1,000,000 tons in 1994 estimations,

over fifty percent were azo dyes (Ollgaard et al., 1999; Stolz, 2001). Based on the azo dyes’

chemical configuration, which is described via one to many (–N=N–) azo groups, in the visible

spectrum light absorption occur (Chang and Kuo, 2000). Azo compounds establish the major as

well as the utmost miscellaneous collection of synthetic dyes which have been extensively utilized

in several industries like food, textile, paper printing and cosmetics (Pandey et al., 2007). Recently

Rajaguru et al. (2002) as well as Umbuzeiro et al. (2005) in their studies publicized that the

addition of azo colorants to surface and ground water through contaminated textile effluents made

them potent mutagens. Their expulsion into surface water harmfully distress the natural assets, soil

fecundity, aquatic organisms as well as disrupt ecosystem legitimacy (Mester and Tien 2000;

Puvaneswari et al., 2006; Zhao et al., 2011) via altering pH, swelling the biochemical oxygen

demand plus chemical oxygen demand, as well as critically upset water quality. Roughly 280,000

tonnes of textile dyes, in a year, are dispensed into the atmosphere globally (Jin et al., 2007)

maximum of which finish up into the aquatic environment. So, the deduction of color from textile

effluents has been a chief anxiety.

Large work has been done on the utilizing physical plus chemical approaches for color

removal of from dyes holding effluents (Swaminathan et al., 2003; Behnajady et al., 2004; Chen

et al., 2010; Malik et al., 2017). Though, employment of physical/chemical approaches have the

innate hitches of being sparingly impractical (demanding extra energy plus chemicals), being

9

incapable to fully eliminate the recalcitrant and obstinate azo colorants either or both their organic

metabolites, making a considerable quantity of sludge that might produce secondary contamination

difficulties, as well as comprising complex processes (Forgacs et al., 2004; Zhang et al., 2004).

Nevertheless, biodegradation is a cost-competitive eco-friendly substitute to chemical decay

procedure which may aid diminish water intake likened with physiochemical approaches i.e.,

adsorption, precipitation, photocatalysis, reduction, reverse osmosis, membrane filtration and

desalination (Verma and Madamwar 2003; Rai et al., 2005).

Bioremediation is a process which is mostly in use these days to either degrade the

environmental pollutants completely or to transform them into less toxic forms. This process is

done by using living organisms or microorganisms. Degraded organic components are mostly

converted into carbon dioxide, water and other inorganic compounds which are usually taken up

by other microorganisms as their source of nutrition. Bioremediation usually involves the role of

microbial enzymes to transform the pollutants into less toxic forms. Therefore, it is very important

that environmental conditions should be optimized to run this process efficiently.

The large amount of dyes used in textile industry lost in wastewater as they do not

completely bind with the fabric. The loss of dye in the wastewater depends upon the class of the

dye and application. It may range from 2% in case of basic dyes while it could be upto 50% for

reactive dyes. This loss of dyes becomes a cause of contaminating surface as well as ground waters

in the surrounding of industries (Ganesh et al., 1994; O’Neill et al., 1999). Many dyes are visible

in water even at a low concentration of 1 mg/L. The wastewater from textile industries is highly

colored having dye concentration between 10–200 mg/L. The discharge of wastewater into water

bodies affects aquatic industry by causing aesthetic problems, causes obstruction of light and

10

oxygen in water bodies (Vandevivere et al., 1998; Weisburger 2002; Öztürk and Abdullah 2006).

It is being reported that colored discharge of textile and other industries where dye is used is more

toxic in nature as compared to discharge of other industries (Houk 1992). The effluent of textile

industry contaminated with azo dyes is responsible for causing mutagenic activity in surface and

ground waters (Rajaguru et al., 2002; Umbuzeiro et al., 2005). Thus, the removal of color from

textile effluents is critical issue these days.

Dyes are stable to degrade in presence of light and therefore can survive in environment.

A dye called Reactive Blue (RB 19) has half-life of 46 years at 25◦C and pH 7 (Hao et al., 2000).

Moreover, they have xenobiotic nature as well for example 4–40 dihydroxy azo benzene (Gill and

Strauch, 1984) so they are resistant to biodegradation. The use of conventional methods of their

treatment or their dilution with domestic wastewater is not sufficient to degrade dyes. There is a

need of alternative methods to degrade these dyes in wastewater.

Mechanisms of azo dye decolorization by microorganisms

Microorganisms degrade azo dyes by two different mechanism i.e., biosorption or

enzymatic degradation or by both (Phugare et al., 2010; Wu et al., 2012).

1. Biosorption

The biomass of various microorganisms such as algae, yeast, filamentous fungi and

bacteria is used in this mechanism to degrade azo dyes (Bhatnagar and Sillanpaa, 2010). Different

functional groups such as amino, carboxyl, hydroxyl, phosphate and other charged groups present

in the heteropolysaccharide and lipid component of cell wall of microorganisms are responsible

for causing strong attractive forces between azo dyes and cell wall (Aksu and Donmez, 2003; Vitor

and Corso, 2008; Srinivasan and Viraraghavan, 2010; Charumathi and Das, 2012). Several

11

pretreatment processes are responsible for increasing adsorption capacity of cell biomass. This

includes autoclaving which ruptures the cell wall and results in an increase surface area (Fu and

Viraraghavan, 2001), treatment with different chemicals such as acids (Srinivasan and

Viraraghavan, 2010), formaldehyde, NaOH (Ambrosio et al., 2012), NaHCO3 and CaCl2 (Fu and

Viraghavan, 2000; Vijayraghavan and Yun, 2007) causes a change in the surface and alters the

binding capacity of various sites. The use of dead cells for biosorption has various advantages over

use of living cells as they have no nutritional requirement, their storage time is greater and can be

regenerated by the use of organic solvents and surfactants (Fu and Viraraghavan, 2001). The

efficiency of biosorption depends on various factors such as temperature, pH, time of contact, ionic

strength, dye concentration, adsorbent and dye concentration, and type of microorganism (Fu and

Viraghavan, 2000; Bakshi et al., 2006; Iqbal and Saeed, 2007; Renganathan et al., 2007;

Vijayraghavan and Yun, 2007; Kaushik and Malik, 2009; Latif et al., 2010; Erden et al., 2011;

Ambrosio et al., 2012).

2. Enzymatic Degradation

The presence of azo linkage (-N=N-) in azo dyes make them an electron deficient

component. The presence of sulphonic (SO3-) or other electron withdrawing groups in their

structure decreases their susceptibility of degradation by microorganisms (Barragan et al., 2007;

Hsueh et al., 2009; Enayatizamir et al., 2011; Kuberan et al., 2011; Kurade et al., 2011).

Microorganisms have reductases for degradation of azo dyes under favorable conditions (Stolz,

2001; Majeau et al., 2010; Chacko and Subramaniam, 2011; Misal et al., 2011, Saratale et al.,

2011; Sugumar and Thangam, 2012). Azo bond is cleaved under anaerobic conditions by anaerobic

azoreductase followed by the transfer of electron between dye molecule and intracellular enzyme

(Fig. 5). As a result, corresponding amines are formed (McMullan et al., 2001; Ramalho et al.,

12

2002; Kodam et al., 2005; Chacko and Subramanian, 2011). Azoreducatases require redox

mediators such as FADH, NADH and NADPH for their proper functioning (dos Santos et al.,

2007; Van der Zee and Cervantes, 2009; Chacko and Subramanian, 2011). The reduction of dye

does not depend on its intracellular uptake due to presence of high molecular weight sulphonate

substituent groups (Robinson et al., 2001; Pearce et al., 2003; Pandey et al., 2007; Chacko and

Subramanian, 2011)

Azo linkages have been reduced by different enzymes which include FMN-dependent

reductases (dos Santos et al., 2007; Burger and Stolz, 2010), FMN-independent reductases (Burger

and Stolz, 2010), NADH-dependent reductases (Ooi et al., 2007; Matsumoto et al., 2010; Misal et

al., 2011), NADPH-dependent reductases (Liu et al., 2007; Misal et al., 2011; Mendes et al., 2011)

and NADH-DCIP reductases (Dhanve et al., 2008; Jadhav et al., 2010; Phugare et al., 2010;

Saratale et al., 2011; Waghmode et al., 2011). The microorganisms which have azoreductases

include bacteria (Vitor and Corso, 2008), algae (El-Sheek et al., 2009) and yeast (Jadhav et al.,

2007).

The azo dyes are degraded aerobically by peroxidases and phenoloxidases which includes

manganese peroxidase (MnP), lignin peroxidase (LiP), laccase (Lac), tyrosinase (Tyr), N-

demethylase (Duran and Esposito, 2000; Baldrian, 2006; Husain, 2010; Jadhav et al., 2011;

Oturkar et al., 2011; Saratale et al., 2011) dye-decolorising peroxidases (Liers et al., 2010) and

cellobiose dehydrogenase (Tilli et al., 2011). These oxidative enzymes are generally present in

filamentous fungi (Majeau et al., 2010), bacteria (Waghmode et al., 2011) and yeast (Martorell et

al., 2012). The non-specific nature of laccases, their effectiveness without cofactor and

dependence on oxygen as electron acceptor make them best option for bioremediation (Telke et

al., 2011; Kalyani et al., 2012). Laccases are able to degrade several complex compounds for

13

example non-phenolic compounds and dye pollutants in the presence of specific mediators such

as 1-hydroxybenzotriazol (Khlifi et al., 2010) or 2, 2ʹ-azinobis (3-ethylthiazoline-6-sulphonic acid)

(ABTS) (Fang et al., 2011). Some reactive radicals such as ABTS are formed when oxidation of

mediators is done by Lac. These radicals are used for oxidation of some dyes like indigo blue,

brilliant blue G, Coomassie blue and Orange 7 without involving enzymes (Solis -Oba et al., 2007).

The Lac oxidation of phenolic azo dyes also results in the production of redox mediators (Soares

et al., 2001; Camarero et al., 2005).

Figure 5: Proposed mechanism for degradation of azo dyes by azo reductase (Keck et al., 1997).

Parameters influencing dye decolorization

The growth of microorganisms is affected by various factors such as presence of chemical

substances, such as dyes, variations in pH, high salinity, and high content of organic compounds

(Dua and Singh, 2002; Ang et al., 2005; Megharaj et al., 2011). Most suitable microorganisms for

14

dye decolorization are those which are isolated from the vicinity of dye contaminated

environments such as soil, wastewater and sludge obtained from treatment of wastewater (Yang et

al., 2009; Ola et al., 2010; Ayed et al., 2011). The following factors affects the biodegradation of

azo dyes.

1. Structure of azo dyes

The structure of azo dyes highly influences their biodegradation (Pasti-Grigsby et al., 1992;

Sawhney et al., 2011; Rajee and Patterson, 2011). The presence of electron withdrawing groups

such as SO3- in their structure aids in their biodegradation as compared to electron releasing groups

such as -NH- triazine. The presence of these electron withdrawing group at ortho and para position

to the azo bond further aids in biodecolorization as compared to meta position because of increased

resonance effect. This makes the dye molecule a strong electrophile which results in an increased

rate of decolorization (Pricelius et al., 2007; Tauber et al., 2008). Azo dyes molecules having less

steric hindrance decolorize easily by microorganisms (Chen et al., 2011). The induction of enzyme

is done by increasing concentration of dye and this induction is dye specific which is maximum at

the end of biotransformation process (Khehra et al., 2005; Moosvi et al., 2007; Ayed et al., 2010).

For example, MnP activity is induced in Debaryomyces polymorphus (Yang et al., 2005) and

Trichosporon akiyoshidainum in the presence of Reactive Black 5 while azoreductase activity in

Chlorella vulgaris and Nostoc linckia in the presence of G-Red and Methyl Red respectively (El-

Sheekh et al., 2009).

The biotransformation of azo dyes is done by both oxidases and reductases. The induction of

different enzymes can take place in the same microorganism for the different dyes depending upon

the structure of azo dye. The activity of Lac is induced in Galactomyces geotricum in the presence

15

of GYHER (Golden Yellow HER) while activity of both Lac (Laccase) and azoreductase is

observed in the presence of Remazol Red (Waghmode et al., 2012).

2. Carbon and nitrogen source

There is a deficiency of carbon in molecule of dyes, so presence of carbon source is essential

for decolorization process (Khelifi et al., 2009; Tony et al., 2009). The addition of carbon source

plays two important role firstly it provides energy for the growth and survival of microorganisms

and secondly, they provide electrons for the reduction of azo bond (Perumal et al., 2007; Gonzalez-

Gutierrez and Escamilla-Silva, 2009). Microorganisms behave differently to different carbon

sources. A consortium of Paenibacillus polymyxa, Micrococcus luteus and Micrococcus sp. has

decolorized Reactive Voilet 5 more efficiently in the presence of glucose (Moosvi et al., 2007)

whereas a combination of different carbon sources is required by some microorganisms to enhance

rate of decolorization for example Staphylococcus arlettae show greater than 90% decolorization

of Reactive Yellow 107 and Reactive Red when glucose and yeast extract is added while when

these sources are substituted with sodium pyruvate less than 50% decolorization is observed

(Elisangela et al., 2009). Microorganisms which use starch as a carbon source render the process

of biodegradation more economical as it is used as an additive in textile finishing process (Babu

et al., 2007) for example, consortium of Paenibacillus polymyxa, M.luteus and Micrococcus sp.

decolorize Reactive Voilet 5 to a greater extent in the presence of starch (Moosvi et al., 2007). The

use of several cheaper carbon sources such molasses, sucrose (Gonen and Aksu, 2009) and

sugarcane baggase (Charumathi and Nilanjan, 2010; Das et al., 2010; Das et al., 2011) for dye

removal renders this process more economical.

On the contaray, the rate of decolorization is decreased by the presence of carbon source for

example Brevibacillus laterosporus does not decolorize dye in the presence of carbon source while

16

decolorzation is increased in the presence of nitrogen source like peptone and yeast (Kurade et al.,

2011). The amount of carbon source is also crucial. It should not be too much high that

microorganisms totally rely on it and ignore dye and also to reduce biomass (Mohana et al., 2008).

Researches have been done to make it possible that microorganisms completely rely on

dyes as carbon and nitrogen source. This is done through the process of acclimatization which

involves a slow increase and decrease in concentration of dye and carbon source respectively until

microorganisms completely rely on dye alone. The example includes the growth of Saccharomyces

cerevisiae (Jadhav et al., 2007) Sphingomonas paucimobilis (Ayed et al., 2011) and consortium

of Pseudomonas, Arthrobacter and Rhizobium (Ruiz-Arias et al., 2010) in the presence of Methyl

Red and Acid Orange 7.

The presence of nitrogen source also plays a key role in the process of decolorization as

their metabolism regenerates NADH (Saratale et al., 2009a). Microorganisms rely on different

nitrogen sources to increase rate of decolorization. A consortium of P. polymyxa, M. luteus and

Micrococcus sp. utilize yeast extract as nitrogen source to decolorize Reactive Voilet 5 (Moosvi

et al., 2007). Consortium of Proteus vulgaris and M. glutamicus utilized agricultural by-products

such as rice husk, rice straw in place of pure substrates like peptone and beef extract to enhance

decolorization rate of Scarlet R (Saratale et al., 2009a).

3. Salinity, dye concentration, pH, temperature and oxygen

The decolorization ability of microorganisms decreases under saline environments as they do

not grow well above 3000 mg/L concentration of sodium (Anjaneya et al., 2011). Some

halotolerant bacteria are reported who degrade dye under high salt concentrations (Meng et al.,

2012). Decolorization of Reactive Black 5 is done by Exiguobacterium acetylicum,

17

Exiguobacterium indicum and Staphylococcus gallinarum in a culture with 60,000 mg/L

concentration of NaCl (Chen et al., 2012).

The decolorization of dye is also affected by the concentration of dye. A decrease in the

decolorization rate of Metanil Yellow by Lysinibacillus sp. from 100% to 62% occurs with an

increase in dye concentration from 200 to 1000 mg/L (Anjaneya et al., 2011).

Enzymatic activity and adsorption is affected by change in pH. It also affects the solubility

of the dye and color of the solution. A variation in the adsorption capacity of Candida tropicalis

of Basic Violet 3 occurred from 45%, 85%, and 33% at pH 3, 4, and 9 respectively (Das et al.,

2011).

Microbial growth and enzyme production is affected by temperature and consequently the

rate of decolorization. A variation in the decolorization rate of 300 mg/L Orange MR by

Micrococcus sp. was observed from 60%, 80%, and 42% at 30, 35, and 45◦C respectively (Rajee

and Patterson, 2011).

The biodegradation of azo dyes is either favored or inhibited by the presence of oxygen.

Mass and oxygen transfer is increased between the cell and medium because of shaking and for

aerobic mechanism of degradation, enzymatic activity is dependent on the presence of oxygen. A

decrease in the decolorization time of Methyl Red by Micrococcus sp from 24 to 6 hours occurred

due to presence of oxygen (Olukanni et al., 2009).

Biological systems involved in the decolorization of azo dyes

A variety of microorganisms have been used for decolorization and mineralization of azo

dyes which includes algae, yeast, filamentous fungi, and bacteria. Consortia of microorganisms

has also been utilized for this purpose.

18

Azo dyes degradation by algae

Azo dyes present in water bodies do not affect algal growth and algae grow efficiently over

industrial effluents (Acuner and Dilek, 2004; Dubey et al., 2011). Thus, they can be used to

degrade azo dyes. Algae do not need additional carbon source like bacteria and fungi (Omar, 2008)

as they use sunlight as energy source and obtain carbon and nitrogen from air. Algae degrade dyes

either by enzymatic degradation, adsorption or both. They can also degrade dyes by induced

azoreductase resulting in the breakage of azo bond and aromatic amines are produced (Omar, 2008;

El-Sheek et al., 2009; Priya et al., 2011). Decolorization process sometimes may also involves

oxidative enzymes (Priya et al., 2011).

The effectiveness of adsorption mainly relies on structure of dye, the specie of algae (Omar,

2008) and pH (Mohan, 2008). Both living and non-viable form of algae can be used for

discoloration of azo dyes (Sivarajasekar et al., 2009; Lim et al., 2010). High uptake of dye

molecules occurs at low pH because of electrostatic forces of attraction between positively charged

biomass and negatively charged dye anions (Srinivasan and Viraraghavan, 2010). Scenedesmus

quadricoda (Ergene et al., 2009), Chlorella vulgaris (Aksu and Tezer, 2005) and non-viable

Spirogyra sp. (Sivarajasekar et al., 2009) showed high adsorption capacity at pH 2. Immobilized

microalgae can also be used for decolorization of azo dyes. In comparison to suspended algae, C.

vulgaris (Chu et al., 2009) and S. quadricoda (Ergene et al., 2009) immobilized on alginate can

remove high percentage of dyes from textile effluent (Ergene et al., 2009).

19

Azo dyes degradation by yeast

Yeast are more effective in decolorization of azo dyes because of their greater potential to

accumulate dyes, fast growth and to with stand unfavorable conditions (Martorell et al., 2012).

Yeast strains also decolorize dyes by adsorption (Yu and Wen, 2005; Aksu and Donmez, 2005),

enzymatic degradation or both. Dyes readily adsorbed on yeast biomass at low pH. For example,

Candida albicans accumulate Direct Violet 51 at maximum level at pH 2.5 (Vitor and Corso,

2008) while C. tropicalis accumulates maximum amount of Violet 3 at pH 4.0 (Charumathi and

Nilanjana, 2010). The extent of adsorption is different for different azo dyes on same strain of

yeast for example C. tropicalis adsorbed 94% of Remazol Blue and 44% of Reactive Red

(Donmez, 2002) while decolorization rate for Reactive Blue and Reactive Red was 63% and 90%,

respectively by Trichosporon akiyoshidanum (Pajot et al., 2007).

The process of dye degradation by yeast mainly depends on the growth process (Yang et

al., 2008; Lucas et al., 2006) and its primary metabolism (Martorell et al., 2012). Glucose is

essential for the growth of yeast strains. The process of degradation of azo dyes depends on the

availability of carbon source (Omar, 2008; Waghmode et al., 2011). The degradation of azo dyes

in yeast is done by oxidases and reductases which include MnP (Manganese dependent

peroxidase), Tyr (Tyrosinase) (Halaburgi et al., 2011; Pajot et al., 2011; Aghaie-Khouzani et al.,

2012; Martorell et al., 2012;) and NADH-DCIP (Nicotinamide adenine dinucleotide dependent

2,6-dichlorophenolindophenol) reductase (Waghmode et al., 2011).

The mechanism of decolorization is influenced by pH. Increase in pH from 2.0 to 7.0 results

in almost complete decolorization of Reactive Blue 221 and Reactive Red 141 from 63% and 90%

respectively (Pajot et al., 2007). Yeast can degrade dyes under both aerobic and anaerobic

20

conditions. The decolorization of a mixture of various dyes which includes Remazol Red, Golden

Yellow HER, Rubine GFL, Scarlet RR, Methyl Red, Brown 3 REL and Brilliant Blue by

Galactomyces geotrichum was higher under aerobic condition as compared to the anaerobic and

anoxic condition (Waghmode et al., 2011) while decolorization of Reactive Blue by C. tropicalis

was done better under static condition (Liu et al., 2011).

Azo dyes degradation by filamentous fungi

Filamentous fungi found in every habitat and they can easily grow on every carbon and

nitrogen source. Due to low cost and their ability to completely mineralize dye they become an

attractive option to degrade dye (Husain and Husain, 2007; Asgher et al., 2008). They usually

decolorize dye by adsorption or enzymatic degradation. Low pH favors the mechanism of

adsorption (Erden et al., 2011). Increase in temperature increases the adsorption capacity of fungal

cells which is due to increased surface activity and kinetic energy of dye molecules (Bakshi et al.,

2006; Kaushik and Malik, 2009). However, at very high temperature decolorization is decreased

due to deactivation of some active sites or deactivation of absorbent surface (Iqbal and Saeed,

2007; Erden et al., 2011).

The enzymes used by filamentous fungi to degrade azo dyes include peroxidases and

phenoloxidases which avoid the production of amines during reduction of dye (Pazarlioglu et al.,

2005; Svobodova et al., 2007; Erkurt et al., 2007; Husain and Ulber, 2011). They decolorize dye

well under aerobic conditions as compared to the anaerobic condition. Better oxygen transfer takes

place at high shaking speed due to greater transfer of oxygen for example degradation of Drimaren

Brilliant Blue by Tramates villosa and Pleurotus sanguineus (Machado et al., 2006).

21

The addition of carbon and nitrogen affects the process of decolorization. The study of

decolorization process of Phanerochaete chrysosporium (Urek and Pazarligolu, 2005; Karimi et

al., 2006) and Bjerkandera sp (Axelsson et al., 2006) indicated that higher rate of decolorization

was achieved when they grow in media where concentration of nitrogen is limited.

Azo dyes degradation by bacteria

Bacteria are more rapidly used to degrade dye because they are easily cultured, grow well

under both anaerobic and aerobic conditions and facultative in nature as well. They can withstand

extreme conditions of temperature and salinity and have a great variety of oxidoreductases. The

reduction of azo bond by bacteria is generally tough under aerobic conditions (Dafale et al., 2009).

However, some bacterial strains are able to degrade dye under aerobic conditions by using oxygen

insensitive or aerobic azoreductases (Nachiyar and Rajakumar, 2005; Lin and Leu, 2008). A

reduction in the decolorization time of reactive dyes by Micrococcus sp. from 24 hours to 6 hours

was reported when the condition was changed from anaerobic to aerobic respectively (Olukanni

et al., 2009). The synthesis of aromatic amines does take place by the degradation of dyes under

aerobic conditions as compared to the microaerophilic conditions which means the cleavage of azo

bond is followed by complete mineralization of amines into less products by oxidation (Wang et

al., 2012).

Bacteria degrade azo dyes with the help of three oxidative enzymes which include Laccase,

Tyrosinase and veratryl alcohol peroxidases. The isolation of laccases generally takes place from

plants and fungi, but fungal laccases are mostly used for various biotechnological processes. As

fungal laccases show sensitivity towards high pH (Xio et al., 2003) and chlorides (Jimenez-Juarez

et al., 2005) bacterial laccases are mostly used because of their stability at high temperature, pH

and salinity (Van Bloois et al., 2010; Santhanam et al., 2011).

22

The industrial effluents usually have high salt concentrations and temperature as well, so

application of extremophiles is a better option for the process of bioremediation (Ertugrul et al.,

2009; Gao et al., 2010; Amoozegar et al., 2011). The halotolerant strains which are able to degrade

dye under high salt concentrations include Staphylococcus, Exiguobacterium (Chaen et al., 2011)

Bacillus firmus and A. hydrophila (Ogugbue et al., 2011).

Azo dyes degradation by using genetically modified microorganisms and

enzymes

The ecofriendly methodology for the degradation of azo dyes in wastewater i.e.,

bioremediation is more economical but different parameters of wastewater such as temperature,

pH, presence of various salts and organic compounds results in damaging the microbial cells and

their enzymes. There is a need to develop such microorganisms and enzymes which are highly

stable and economical as well. Various techniques of molecular biology such as cloning,

heterologous expression, random mutagenesis, site directed mutagenesis, gene recombination

techniques, directed evolution, rational design and metagenomics are helpful in this regard.

Moreover, advances in the field of genetic engineering and microbial genetics make it possible to

express any gene of interest in a suitable host.

Cloning of Lac gene (lac48424-1) of white rot fungi Trametes sp. 48424 was done in Pichia

pastoris. The purified product (rLAC48424-1) decolorizes Methyl Orange and Bromophenol Blue

with greater efficiency in comparison to other laccases (Fan et al., 2011).

The site directed mutagenesis of bacterial dye decolorizing peroxidase from

cyanobacterium Anabena sp. strain PCC7120 (AnaPX) which is sensitive to H2O2 was done. The

resulting product was more stable in presence of H2O2 and have increased dye degrading ability

which was 96% in case of Reactive Black 5 (Ogola et al., 2011).

23

Azo dyes degradation by using consortia

The degrading product of dye by single microbial strain are aromatic amines which are

more toxic and difficult to degrade as compared to the original dye. Each microbial strain is

specific for a dye. So, by building microbial consortia complete degradation of azo dyes will occur

by the combination of various enzymatic processes (Asgher et al., 2007; Joshi et al., 2010). The

dye can be attacked by different sites when different strains are used, and degradation products

synthesized by one strain can be used by other strain results in the complete mineralization of the

dye (Saratale et al., 2009). The presence of other microorganisms influences the enzymatic activity

of a single strain and degradation ability of consortia is greater than its constituent strains.

A consortium of Pseudomonas sp. SUK1, Pseudomonas sp. LBC2, and LBC3, decolorizes

dyes efficiently in comparison to Pseudomonas sp. SUK1 alone while Pseudomonas sp. LBC2 and

LBC3 have no biodegrading ability alone (Jadhav et al., 2010).

In order to obtain an effective system to decolorize dyes the proportion of every

microorganism is very important in the construction of consortium. For example, a consortium

comprising of 0.023% S. paucimobilis, 0.459% Bacillus sp. and 0.516% S. epidermidis was able

to decolorize Congo Red completely and with 95% COD removal (Ayed et al., 2010).

The combination of aerobic and anaerobic systems in the form of a consortium makes a

promising system for decolorization of azo dyes. The anaerobic strains cleave the azo bond by

reduction which is shown by decolorization (Yemashova et al., 2009) while aerobic strains play

their role in the complete mineralization of aromatic amines (Forss and Welandar, 2011; Jonstrup

et al., 2011; Koupaie et al., 2011). A consortium comprising of Enterococcus casseliflavus,

Enterobacter clocae (NAR-1) degrade Orange II under microaerophilic conditions into sulphanilic

acid while individual strains are not able to degrade acid even after 5 days (Chan et al., 2011).

24

Condition of dye pollution in Pakistan

Textile industry is the major industry in Pakistan and we earn major revenue by exporting

the products of this industry. But apart from its major contribution to our growing economy it

becomes a source of causing water pollution. Wastewater that is generated at various steps in

textile industry have high pH, temperature, detergents, suspended and dissolved solids, leveling

agents, toxic colors and alkalinity. The major pollutants produced by the textile industry includes

recalcitrant organics, toxic colors, toxicants, surfactants and chlorinated compounds (AOX). In

Pakistan, this problem of water pollution is severely high due to lack of wastewater management

system by industries as well as the ignorant behavior of concerning industries i.e., environment

and sanitation (Noreen et al., 2019).

Untreated textile effluent, is being discharged into rivers and canals which pose toxic

effects to humans, plants and animals. Hepatitis is major problem caused by water pollution since

last two decades. According to recent reports every third individual is suffered from this fatal

disease in Pakistan. Moreover, a major problem that arise these days is the use of this wastewater

for the cultivation of crops which has serious impact on human health (Azizullah et al., 2010).

As the various physiochemical methodologies used to treat textile effluent are costly and

less efficient and produced secondary waste which is toxic as well so there is need of time to treat

such effluents with bioprocessing which employs the use of various microorganisms to degrade

pollutants. These processes are cost effective and environment friendly as well.

25

Aims and objectives

The main aims of this study are

• To isolate and characterize azo dye degrading bacteria.

• To culture and maintain the microorganisms in laboratory conditions.

• To determine the optimal growth conditions of the isolated bacteria.

• To identify the isolated bacteria.

• To determine the optimal conditions for degradation of azo dyes.

• To ascertain the potential of isolated bacteria to degrade multiple dyes.

• To study the effect of decolorization on the growth of bacteria.

• To study how consortium of bacteria affects decolorization rate.

• To analyze dye degrading products through TLC, HPLC, FTIR and GC-MS.

• To study biodecolorization at large scale.

• To determine the microbial toxicity and phytotoxicity of biodecolorized effluent.

26

MATERIALS AND METHODS

Sample collection

Industrial wastewater samples were collected from Kot Lakhpat industrial estate, Lahore.

The samples were collected in autoclaved screw capped bottles. Physiochemical parameters such

as temperature, pH, as well as color of samples were also noted at the time of sample collection.

Sample enrichment

Wastewater samples were enriched by adding 1 ml of each sample to 100 ml of L-broth

prepared in 250 ml Erlenmeyer flasks and then incubated at 37oC in the shaker at 150 rpm for 24

hours. L-broth was prepared by dissolving tryptone (10g), yeast extract (5g) and NaCl (5g) in 1000

ml of distilled water. The pH of the medium was adjusted to 7. The autoclaving was done at 121oC

for 15 minutes at 15 psi.

Dyes

The dyes used in the recent research worked were named Synazol Red HF-6BN and

Sumifex Tourqi blue obtained from a textile industry and decolorization was measured by

measuring the absorbance of decolorized medium at 465 nm (Ilyas and Rehman, 2013).

Isolation of azo dyes degrading bacteria

In order to isolate azo dye degrading bacteria serial dilutions of the enriched samples were

made i.e., 10-1, 10-2, 10-3 and 10-4. The bacteria were isolated by plating 50 µl of each dilution on

the L-agar plates. Each dilution was plated twice, and plates were then incubated under aerobic

and anaerobic (by using anaerobic jar along with anaerobic sachet) conditions to isolate respective

27

bacteria. L-agar medium was prepared by dissolving tryptone (10g), yeast extract (5g), NaCl (5g)

and agar (15g) in 1000 ml of distilled water. The pH of the medium was adjusted to 7.

Evaluation of dye degrading potential

The bacterial isolates were cultured in 100 ml of mineral salt medium (MSM) prepared in

250 ml Erlenmeyer flask and incubated for 4 days at 37 oC. The composition of MSM is (g/L):

(NH4

)SO4

, 0.28; NH4

Cl, 0.23; KH2

PO4

, 0.067; MgSO4

·7H2 O, 0.04; CaCl2

·2H2

O, 0.022;

FeCl3

·6H2 O, 0.005; yeast extract, 0.2; NaCl, 0.15; NaHCO3

, 1.0 and 1ml/L of a trace element

solution containing (g/L): ZnSO4

·7H2

O, 0.01; MnCl2

·4H2 O, 0.1; CuSO4

·5H2 O, 0.392;

CoCl2

·6H2

O, 0.248; NaB4

O7

·10 H2

O, 0.177; and NiCl2

·6H2

O, 0.02 (Parshetti et al., 2006) with

glucose and yeast extract (1% each) used as carbon and nitrogen source respectively. The medium

was supplemented with dye at a concentration of 50 mg/L.

Aliquot was taken at the time of

incubation and after 4 days of incubation, centrifuged at 3000 rpm and optical density was

measured at 465nm to record initial and final absorbance respectively. The percentage

decolorization of azo dyes in the sample was calculated by the following formula

Decolorization (%) = Initial absorbance - Final absorbance/Initial absorbance


The identification of bacterial isolates was done on the basis of different morphological

parameters such as colony shape, colony size, margin, elevation, and light transparency and

biochemical tests such as Gram staining, spore staining, catalase test etc (Cappuccino et al., 1996).

28

Determination of optimum growth conditions of bacterial isolates

The optimum growth conditions i.e., temperature and pH of the selected bacterial isolates

were determined.

Optimum temperature

To determine the optimal temperature, bacterial isolates were grown on four different

temperatures. They were cultured in 10 ml L-broth taken in a test tube which was then autoclaved

and cooled at room temperature. An inoculation of the medium was done with 50 µl of bacterial

cells which were in their log phase. The tubes were incubated at different temperature i.e., 20oC,

30oC, 37oC, and 45oC after inoculation. After 24 hours of incubation optical density was taken at

600 nm by using spectrophotometer. To plot a graph temperature was taken on X- axis with their

respective cell densities on Y- axis.

Optimum pH

To determine the optimal pH, bacterial isolates were grown on six different pH. They were

cultured in 10 ml L-broth taken in a test tube whose pH was adjusted to 5, 6, 7, 8, 9 and 10 and

was then autoclaved and cooled at room temperature. An inoculation of the medium was done with

50 µl of bacterial cells which were in their log phase. The tubes were incubated at respective

optimal temperature of bacterial isolate after inoculation. After 24 hours of incubation optical

density was taken at 600 nm by using spectrophotometer. To plot a graph pH was taken on X- axis

with their respective optical densities on Y- axis.

29

Growth curves

To determine the growth curves of bacterial isolates they were grown in L-broth. It was

prepared in 250 ml Erlenmeyer flasks for this purpose. The autoclaving of the medium was done,

and it was cooled at room temperature. About 1 ml log phase bacterial culture was used to inoculate

this medium. The optical density was measured at the time of inoculation and after every 4 hours

up to 28 hours at 600 nm by using spectrophotometer. To plot a graph, time interval was taken at

X-axis with respective optical densities on Y-axis.

Molecular characterization of bacterial isolates

DNA isolation (Masneuf-Pomarade et al., 2007)

The following protocol was followed for DNA extraction of bacterial isolates. The growth

of bacterial cultures was made in 50 ml L-broth at 37oC on a rotary shaker at 130 rpm for about

24 hours. The pellet was obtained by centrifugation of the entire medium at 6,000 rpm for 10

minutes. The washing of the pellet was done with 1.5 ml of sterile distilled water and was again

centrifuged on the same speed. The pellet was then dissolved in 300 µl of lysis buffer (1% SDS

and 0.1% mercaptoethanol) and incubated at 37oC for 1 hour. After that, disruption of the cells

was made with vigorous vortexing. Then centrifugation was done at 14,000 rpm for 10 minutes to

separate the disrupted cells and supernatant. The supernatant was then taken in a fresh eppendorf,

mixed with 500 µl phenol-chloroform-isoamylalcohol (25:24:1) by vortexing of 3 minutes and

again centrifuged at 14,000 rpm for 5 minutes. About 500 µl of chloroform-isoamylalcohol (24:1)

was then added, vortexed and centrifuged at 14,000 rpm for 5 minutes. The supernatant was then

taken in a new eppendorf and double amount of absolute ethanol (100%) was added to precipitate

30

out DNA. The pellet of DNA was then washed with 70% ethanol after a centrifugation of 10

minutes at 14,000 rpm. The pellet obtained was then dried at room temperature, then suspended in

50 µl sterile distilled and stored at -20oC till further use.

Agarose gel electrophoresis

To make 1% agarose gel solution, 1 g of agarose was dissolved in 20 ml of 5X tris-borate-

EDTA buffer (10.0g tris base, 1.48g boric acid, 1.08 ml of 0.5 M EDTA and distilled water making

volume up to 200 ml, pH 8.0) and make up the volume up to 100 ml by using distilled water. The

solution was then boiled to make it clear. An addition of 2 µl of ethidium bromide (10 mg/ml) was

made to the gel and then it was poured into gel casting tray and comb was inserted to make wells.

The comb was removed when the gel was solidified. The gel tray was then placed in gel tank and

tris-EDTA buffer (160 ml of tris borate-EDTA buffer in 40 ml distilled water) was added in it.

About 5 µl of isolated genomic DNA was then mixed with 3 µl of gel loading dye (0.25%

bromophenol blue and 40% w/v sucrose dissolved in distilled water) and was loaded into the wells

made on the gel. The DNA marker (GeneRuler 100bp DNA ladder #SM0323 Fermentas) was also

loaded to estimate the size of DNA. A voltage of 100 volts for the gel to run. The power supply

was switched off after tracking dye reached the end of gel. The gel was then visualized by using

UV transilluminator (WAEL TEC UV transilluminator) and DNA appeared as pink fluorescent

bands. A Gel Doc System (WAEL TEC Dolphin-Doc) was used to make picture of DNA bands.

31

PCR amplification

The amplification 16S ribosomal DNA region was done by using universal bacterial

primers 8F (5-AGAGTTTGATCCTGGCTCAG-3) and 1492R (5-

GGTTACCTTGTTACGACTT-3) (Turner et al., 1999). A reaction mixture of 25 µl of distilled

water containing 2.5 µl of each primer (20 pmol), 6 µl of genomic DNA (5 µg/ml) (Amersham

Pharmacia, Piscataway, NJ, USA) was used to perform amplification reactions. PCR was

performed by initial denaturation at 95oC for 5 minutes, followed by 35 cycles of denaturation at

95oC for 1 minute, annealing at 55oC for 1 minute and extension at 72oC for 1 minute, with a final

extension at 72oC for 10 minutes.

Gene cleaning

Gene cleaning of PCR products was done with Fermentas Gene Jet Gel Extraction

kit (#K0691) according to manufacturer’s instructions (Khan et al., 2015).

Sequencing and submission of sequences to GenBank

The amplified and gene clean products of DNA were submitted at 1st Base, Malaysia for

sequencing. Nucleotide BLAST search program at the NCBI website

http://ncbi.nlm.nih.gov/BLAST/ was used to analyze 16S rRNA gene sequence. The submission

of sequenced data was done to GenBank to obtain accession numbers of the respective bacterial

isolates.

http://ncbi.nlm.nih.gov/BLAST/

32

Construction of phylogenetic tree

The sequences showing similarity were aligned and phylogenetic tree were constructed

with the help of neighbor joining method in MEGA 7 software (Kumar et al., 2016) taking 1000

as bootstrap value.

Optimization of biodegradation conditions

The ability of selected bacterial strains to biodegrade azo dye was checked at different

temperatures i.e., 20, 30, 37, and 45° C and pH i.e., 5, 6, 7, 8, 9, and 10 in MSM broth. Moreover,

the effect of incubation condition (static and shaking), carbon (saw dust, sugarcane bagasse and

wheat bran) and nitrogen sources (yeast extract, beef extract and peptone), inoculum percentage

(2%, 4%, 6%, 8% and 10%) and dye concentration (3, 5, 7, 10, 20, and 50 mg/L) was also checked.

In order to observe the effect of temperature and pH, three sets of 100 ml Erlenmeyer flasks

were prepared for each bacterial strain in which 50 ml of MSM was dispensed and labelled for

respective temperature and pH. Flasks were then inoculated with almost equal number of cells of

respective bacterial strain and then incubated at labelled temperature and pH for 4 days. The

percentage decolorization of the dye was calculated after 4 days with the formula mentioned above.

To find the optimum incubation condition for maximum decolorization the cultured flasks

were incubated at static and shaking conditions and after 4 days decolorization percentage was

calculated. The MSM medium was supplemented with 1% of given carbon and nitrogen source to

find out which carbon and nitrogen source gives the maximum percentage of decolorization.

33

To check the effect of inoculum percentage on dye decolorization the inoculum percentage

was slightly increased in comparison to the volume of the medium. The effect of dye concentration

was also monitored to find out the minimum inhibitory concentration of the dye for bacterial

strains. The various concentration of the dye used were 3, 5, 7, 10, 20, and 50 mg/L.

Decolorization of multiple dyes

The bacterial strains were checked for their ability to degrade a mixture of dyes as textile

effluents contain various dyes. For this purpose, bacterial strains were grown in 100 ml of MSM

medium in 250 ml Erlenmeyer flask containing a mixture of various dyes and incubated at

respective optimum dye decolorizing temperature. The mixture of dyes contains Congo Red,

Methyl Red, Phenol Red, Brilliant Black and Navy Blue at a concentration of 50 mg/L. The

decolorization percentage was calculated after 5 days of incubation.

Growth Decolorization relationship

The effect of decolorization was also monitored on the growth of bacterial strains. For this

purpose, bacterial strains were grown in 100 ml of MSM medium in 250 ml Erlenmeyer flask and

incubated at respective optimum dye decolorizing temperature. The dye concentration was kept

constant at 50 mg/L. An aliquot of sample was taken after every day till 5 days and optical density

was calculated at 600 nm to find out a growth ratio. Decolorization was calculated by measuring

the optical density of supernatant at 465 nm.

34

Consortium studies

Development of consortium

Each bacterial isolate was inoculated in 10 ml of LB broth in a test tube and incubated at

their respective optimum temperature for 24 hours. The optical density of each culture was

adjusted to 0.5 after 24 hours with autoclaved water. Then 100 ml of LB broth prepared in 250 ml

Erlenmeyer flasks was inoculated with 0.5 ml of optical density adjusted culture of those bacteria

whose consortium was desired to be made and then incubated at 37 oC.

Aerobic consortium

The consortium of aerobic isolates was developed as mentioned above and 1 ml of this

consortium was used to inoculate 100 ml of MSM medium which was supplemented with dye at

a concentration of 50 mg/L. The medium was then incubated at 37 oC for 5 days. Aerobic strains

were also grown individually to compare the dye decolorization rate.

Anaerobic consortium

The consortium of anaerobic isolates was also developed as mentioned above and 1 ml of

this consortium was used to inoculate 100 ml of MSM medium which was supplemented with dye

at a concentration of 50 mg/L. The medium was then incubated at 37 oC for 5 days under anaerobic

conditions. Anaerobic strains were also grown individually to compare the dye decolorization rate.

Mixed consortium

The consortia of aerobic and anaerobic isolates were developed as mentioned above.

Firstly, 100 ml of MSM medium which was supplemented with dye at a concentration of 50 mg/L

35

was inoculated with 1 ml of culture from anaerobic consortium incubated at 37oC for 2.5 days

under anaerobic conditions. After this period, the medium was further inoculated with 1 ml of

culture from aerobic consortium and then incubated at 37oC for further 2.5 days under aerobic

conditions. Dye decolorization rate of mixed consortium was compared with that of aerobic and

anaerobic consortium.

Analysis of dye degraded products

Sample preparation

In order to extract metabolites for analysis, 100 ml of dye degraded medium was mixed

with an equal volume of ethyl acetate and organic phase was dried over anhydrous rated Na2SO4

and concentrated in rotary evaporator at 4°C. The concentrated sample was then mixed with 1ml

to 2 ml of HPLC grade methanol.

Thin layer chromatography (TLC)

In order to analyze the biodegraded products of dye, TLC was performed. Silica gel F-254

TLC aluminum sheet (MERCK) which was 0.2 mm gel thick was used in this experiment. The

solvent system comprising of n-propanol, methanol, ethyl acetate, water, and glacial acetic acid in

the ratio 3:2:2:1:0.5 (Kalyani et al., 2008) was used to perform TLC. The results were visualized

under UV illuminator at wavelength of 254 and 366 nm.

High performance liquid chromatography (HPLC)

HPLC was performed to analyze the dye and its biodegraded products. HPLC (Waters

model no 2690) was performed with C18 column having symmetry 250 x 4.6 mm using methanol

36

as mobile phase with a flow rate of 1.0 ml/minute for 15 min and UV detector at 254 nm (Telke et

al., 2009).

Fourier transform infrared spectroscopy (FTIR)

In order to observe the difference in the structure of dye before and after decolorization

FTIR was performed. The FTIR analysis was done in the mid IRF region of 400-4000 /cm. Prior

to analysis sample was mixed with pure KBr in the ratio of 5:95 and pellet were then fixed in the

holder for analysis (Saratale et al., 2009b).

Gass Chromatography/Mass Spectrometry (GC/MS)

Dye degraded products were extracted with ethyl acetate from inoculated and un-

inoculated dye augmented MS broth for GC-MS analysis. A gas chromatograph system (GCMS-

QP2010 Ultra, Shimadzu) equipped with capillary column (DB-5ms) was used to analyze products

before and after bacterial treatment. The samples were diluted 1:100 and injected 1.0 µl sample

into GC-MS in split mode with injector temperature of 290°C. Helium was used as a carrier gas

with flow rate of 1.02 ml/min under 54.9 KPa inlet pressure. The column temperature was set to

50°C and hold for 1 min with subsequent increase to 280°C with ramp rate of 30°C/min without

holding and finally to 310°C with ramp rate of 15°C/min by holding it for 8 min. The peaks were

identified by comparing with NIST27.LIB mass spectra library of GC-MS database.

Dye degradation on large scale

An experiment was designed to check the degradation of azo dyes on large scale. For this

purpose, three plastic containers were used. In the first container there was 10 L of tap water along

with dye used as negative control, second container have 10 L of industrial wastewater along with

dye used as positive control while in the third container there was 10 L of industrial wastewater

37

along with 1.5 L of bacterial culture and dye. The experiment was conducted at room temperature

and concentration of the dye was maintained at 50mg/L. The samples were taken from these

containers with an interval of 2 days up to 8 days and percentage decolorization of dye was

calculated with the mentioned formula.

Microbial toxicity

The decolorized dye sample was checked for its toxic effects on soil microflora which have

importance in agriculture i.e., B. subtilis, B. megaterium and B. cereus (Mali et al., 2000). For this

purpose, L-agar plates were prepared on which these bacteria were swabbed. A well of 2 mm was

made in the center of plates which was then filled with dye decolorized broth. An incubation of 24

hours was given to plates at 37°C. The index of toxicity of decolorized sample was measured by

the zone surrounding the well.

Phytotoxicity

To analyze the effect of treated wastewater on the growth of plants, the six seeds of Vigna

radiata (mung beans) were cultivated in autoclaved soil on lab scale. The control pots (n=2) were

those which were watered with tap water and untreated wastewater while experimental pots (n=2)

were those which were watered with bacterially treated wastewater. The plants were grown for

about a period of 10 days at room temperature under 1:1 light and dark period. The comparison of

control and experimental plants were made after 10 days to record the effect of dye on plant

growth.

Statistical analysis

Every experiment was repeated three times for result authentication. The results were subjected

to standard deviation, mean and analysis of variance (ANOVA).

38

RESULTS

Physiochemical characteristics of wastewater samples

The physiochemical characteristics of wastewater collected from industrial area of Sheikhupura

and Lahore have been summarized in Table 1.

Table 1: Physiochemical parameters (temperature, pH) of wastewater collected from industrial

area of Sheikhupura and Lahore.

Sample Location Temperature °C pH Color Smell

CPL-1 ICI Industrial

effluents (Lahore)

32 8 Light yellow No smell

CPL-2 Sewage sample-A

Sheikhupura road

36 7 Black muddy Foul smell

CPL-3 Ghee industry

Sheikhupura road

34 6 Blackish No smell

CPL-4 Paper Mills

Sheikhupura road

28 6 Transparent,

Black particles

No smell

CPL-5 Bata leather

processing plant

Sheikhupura road

30 9 Reddish pink Rotten egg

smell

CPL-6 Sewage sample-B

Sheikhupura road

34 7 Muddy Muddy smell

CPL-7 Steel industry

(Lahore)

28 10 Black No smell

CPL-8 Paint industry

(Lahore)

26 10 Blue No smell

CPL-9 Textile industry

(Lahore)

36 8 Maroon No smell

39

Isolation and evaluation of azo dye degrading potential of bacteria

For the isolation of dye degrading bacteria, the samples collected from industrial area of

Sheikhupura and Lahore were enriched, serially diluted and 50µl of each sample were spread on

L-agar plates in duplicate. After incubation at 37oC for 24-48 hours under aerobic and anaerobic

conditions, different bacterial colonies obtained were then grown in MSM at dye concentration of

50mg/L to evaluate their dye degrading potential. A total of 7 isolates i.e., 4 anaerobic and 3

aerobic were selected for further studies on the basis of their dye degrading potential (Table 2)

(Fig. 6).


The different morphological characteristics of the bacterial isolates e.g., size, shape and

color of the colonies along with elevation, margin and texture were noted on L-agar plates. The

morphological and biochemical characteristics of bacterial isolates are mentioned in Table 3.

40

Table 2: Decolorization (%) of bacterial isolates, isolated from 9 samples.

S. No. Bacterial isolate Decolorization (%)

Anareobic

1. 1b 71

2. 1m 70

3. 1p 42

4. 4e 49

5. 1f 57

6. 2p 73

7. 3h 45

8. 3i 34

9. 3j 47

10. 4m 76

11. 3g 55

12. 4v 79

13. 1c 21

14. 2i 62

Aerobic

1. 1g 23

2. 2f 32

3. 3c 82

4. 3d 78

5. 4k 59

6. CBL-2 83

7. CBL-4 64

41

Figure 6: Change in color of Synazol Red HF-6BN and Sumifex Tourqi blue as compared to the

control in flask containing culture of bacterial strain (a) 1b (b) 1m (c) 2p (d) 4v (e) 3c (f) 3d (g)

CBL-2 isolated from the dye contaminated wastewater.

(a) (b)

(c)

(g) (e)

(e) (d)

(f)

(c)

42

Table 3: Morphological and biochemical characteristics of bacterial isolates. Isolate Size Shape Color Elevatio

-n

Edges Texture Gram

staining

Spore

staining

Cat Ma-

n

Pig Fru Ind VP MR Cit Nit

Ure

1b 1

mm

Round Cream Flat Smooth Mucoid + cocci NA + - - + NA NA NA NA NA NA

1m 1

mm


2p 4mm Round Off

white

Elevated Smooth Dry + rods + NA + NA NA NA + NA NA NA NA

4v 3

mm


3c 1

mm

Round Pale

yellow

Flat Smooth Mucoid - rods NA NA NA NA NA - - - + - -

3d 1

mm

Round Pale

yellow

Flat Smooth Mucoid - rods NA NA NA NA NA - - - + - -

CBL-2 1

mm

Round Yellow Raised Smooth Mucoid - rods NA NA NA NA NA - - - - - -

Cat=Catalase, Man=Mannitol, Pig= Pigment ptoduction, Fru=Fructose fermentation, Ind=Indole production, VP = Voges Proskauer,

MR=Methyl Red, Cit= Citrate utilization, Nit=Nitrate reduction, Ure=Urease, += Positive, - = Negative, + No result, NA=not applicable

43

Determination of optimum growth conditions of bacterial isolates

The optimum temperature, pH, and growth pattern of bacterial isolates were determined

to find out the conditions at which organisms grow at their full potential.

Optimum temperature

The selected bacterial isolates were grown at various temperatures and pH of was adjusted

at 7.0 for the optimization of growth conditions. The growth of all the six bacterial isolates i.e., 1b,

1m, 2p, 4v, 3c, 3d and CBL-2 was determined at different temperature i.e., 20oC, 30oC, 37oC and

45oC. A fair growth of bacterial isolated was observed on all temperatures but they grow optimally

at 37oC (Fig. 7 a and b).

Optimum pH

The isolates 1b, 1m, 2p, 4v, 3c, 3d and CBL-2 were grown on their optimum temperature

with different pH’s to determine the optimal pH. The pH of L-broth medium was adjusted to 5, 6,

7, 8, 9 and 10. All isolates grow optimally on neutral pH i.e., 7 (Fig. 8 a and b).

Growth pattern of bacterial isolates

In order to study the growth pattern of bacterial isolates i.e., 1b, 1m, 2p,4v, 3c, 3d and

CBL-2 L-broth medium was used, and optical density was recorded after every 4 hours till 28

hours (Fig 9 a to f.).

The growth of isolates 1b, 1m and 3d had a rise till 24 hours after that it became stable. The isolates

2p ,4v and CBL-2 clearly showed rise in growth until 20 hours and after that fall in growth pattern

44

was observed. The bacterial isolate 3c showed rise in growth till 16 hours before adopting a

stability in growth pattern.

Figure 7: Growth of bacterial strains at various temperatures after incubation of 16 hours (a) static

and (b) agitated conditions.

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0.9

1

20 30 37 45

Op

tica

l d

ensi

ty a

t 6

00

nm

Temperature (◦C)

(a)1b

1m

2p

4v

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

20 30 37 45

Op

tica

l d

ensi

ty a

t 6

00

nm

Temperature (◦C)

(b) 3c

3d

CBL

-2

45

Figure 8: Growth of bacterial isolates at various pH after incubation of 16 hours in (a) static and

(b) agitated conditions.

0

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0.9

5 6 7 8 9 10

Op

tica

l d

ensi

ty a

t 6

00

nm

pH

(a) 1b

1m

2p

4v

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

1.8

5 6 7 8 9 10

Op

tica

l d

ensi

ty a

t 6

00

nm

pH

(b)3c

3d

CBL-

2

46

0

0.05

0.1

0.15

0.2

0.25

0.3

0.35

4 8 12 16 20 24 28

Op

tica

l d

ensi

ty a

t 6

00

nm

Time (hours)

(a)

0

0.05

0.1

0.15

0.2

0.25

4 8 12 16 20 24 28

Op

tica

l d

ensi

ty a

t 6

00

nm

Time (hours)

(b)

47

0

0.02

0.04

0.06

0.08

0.1

0.12

4 8 12 16 20 24 28

Op

tica

l d

ensi

ty a

t 6

00

nm

Time (hours)

(c)

0

0.05

0.1

0.15

0.2

0.25

4 8 12 16 20 24 28

Op

tica

l d

ensi

ty a

t 6

00

nm

Time (hours)

(d)

48

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

1.8

2

4 8 12 16 20 24 28

Op

tica

l d

ensi

ty a

t 6

00

nm

Time (hours)

(f)

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

1.8

4 8 12 16 20 24 28

Op

tica

l d

ensi

ty a

t 6

00

nm

Time (hours)

(e)

49

Figure 9: Growth curves of bacterial isolates (a) 1b, (b) 1m, (c) 2p, (d) 4v, (e) 3c, (f) 3d and (g)

CBL-2 in LB medium at their respective optimum temperatures. Optical densities were taken at

600 nm after regular time interval.

Molecular characterization of bacterial isolates

The genomic DNA of the bacteria was isolated (Fig. 10) and amplified by using PCR (Fig.

11) with the help of general bacterial primers for molecular characterization. All the dye degrading

strains 1b, 1m, 2p, 4v, 3c, and 3d were identified with 16S rRNA gene sequencing. Three isolates

showed close homology with genus of Staphylococcus. The highest homology had been observed

with Staphylococcus lentus (1b, 1m and 4v) whereas another isolate was found to be homologous

to Bacillus subtilis (2p). Two strain showed close homology with genus Alcaligenes and they are

Alcaligenes aquatilis (3c) and Alcaligenes faecalis (3d). One specie showed homology with

Alishewanella sp. (CBL-2). The nucleotide sequences coding for the 16S rRNA gene of all the

isolates have been submitted to the GenBank database to obtain accession numbers which are

given in the Table 4 and phylogenetic tree were also constructed (Fig. 12 a to g and Fig. 13).

-0.2

0

0.2

0.4

0.6

0.8

1

1.2

1.4

1.6

1.8

4 8 12 16 20 24 28

Op

tica

l d

ensi

ty a

t 6

00

nm

Time (hours)

(g)

50

Figure 10: Lane M, DNA marker (GeneRuler 100bp DNA ladder #SM0323 Fermentas); Lanes

1b, 1m, 2p, 4v, 3c, 3d, and CBL-2 represent genomic DNA of bacterial isolates. The gel is 1%

agarose and stained with ethidium bromide.

Figure 11: Lane M, DNA marker (GeneRuler 100bp DNA ladder #SM0323 Fermentas); Lanes

1b, 1m, 2p, 4v, 3c, 3d, and CBL-2 represent PCR product of 16S rRNA gene of bacterial

isolates. The gel is 1% agarose and stained with ethidium bromide.

1b 1m 2p 4v 3c 3d CBL-2 M

bp

3000

2000

1500

1200

1000

900

800

700

600

500

400

300

200

100

1b M 1m 2p 4v 3c 3d CBL-2

51

Table 4: The nearest homolog and their percentage homology with the azo dye degrading bacterial

strains after 16S rRNA sequencing.

Bacterial isolate Nearest homolog % Homology Accession

numbers

1b Staphylococcus lentus 99% KX950707

1m Staphylococcus lentus 99% KX950708

2p Bacillus subtilis 97% KX950709

4v Staphylococcus lentus 95% KX950710

3c Alcaligenes aquatilis 98% KY009932

3d Alcaligenes faecalis 97% KY009933

CBL-2 Alishewanella sp. 88% JN546609

KC951997.1 Staphylococcus sp. SC1 16S ribosomal RNA gene partial sequence

LC383925.1 Staphylococcus lentus JCM 2426 gene for 16S ribosomal RNA partial sequence

KX525724.1 Staphylococcus sp. strain 82584 16S ribosomal RNA gene partial sequence

MH569456.1 Bacterium strain GIm45 16S ribosomal RNA gene partial sequence

KM378611.1 Staphylococcus lentus strain BGZ-7 16S ribosomal RNA gene partial sequence

KM010146.1 Staphylococcus lentus strain muzyellow 16S ribosomal RNA gene partial sequence

HQ437165.1 Staphylococcus sp. CSA7(2011) 16S ribosomal RNA gene partial sequence

KX950708.1 Staphylococcus lentus strain 1b 16S ribosomal RNA gene partial sequence11

10

10

9

1

(a)

52

KM010139.1 Staphylococcus lentus strain muzorange 16S ribosomal RNA gene partial sequence

MH491955.1 Staphylococcus lentus strain 3-4 16S ribosomal RNA gene partial sequence

KT260750.1 Staphylococcus lentus strain RCB538 16S ribosomal RNA gene partial sequence


KP789167.1 Staphylococcus sp. MRSA222B1_13_3E 16S ribosomal RNA gene partial sequence

MF948914.1 Staphylococcus lentus strain PSB295 16S ribosomal RNA gene partial sequence

LN999936.1 Staphylococcus lentus partial 16S rRNA gene isolate SLKr1

KX950707.1 Staphylococcus lentus strain 1m 16S ribosomal RNA gene partial sequence12

12

11

9

2

MG651202.1 Bacillus halotolerans strain FJAT-47799 16S ribosomal RNA gene partial sequence

KY575587.1 Bacillus subtilis strain Bs_Pg1 16S ribosomal RNA gene partial sequence

KT986121.1 Bacillus tequilensis strain Lmb056 16S ribosomal RNA gene partial sequence

KT986166.1 Bacillus amyloliquefaciens strain Xmb038 16S ribosomal RNA gene partial sequence

KT986156.1 Bacillus axarquiensis strain Xmb027 16S ribosomal RNA gene partial sequence

KT986083.1 Geobacillus stearothermophilus strain Lmb010 16S ribosomal RNA gene partial sequence

KT986187.1 Bacillus subtilis strain Xmb062 16S ribosomal RNA gene partial sequence

KX950709.1 Bacillus subtilis strain 2p 16S ribosomal RNA gene partial sequence12

11

10

9

2

KC951997.1 Staphylococcus sp. SC1 16S ribosomal RNA gene partial sequence

KY653102.1 Staphylococcus lentus strain FJAT-hcl-24 16S ribosomal RNA gene partial sequence

KR364789.1 Staphylococcus sp. WCA-CC-3-1 16S ribosomal RNA gene partial sequence

NR_043418.1 Staphylococcus lentus strain MAFF 911385 16S ribosomal RNA gene partial sequence

AY161045.1 Staphylococcus lentus strain PLC-6 16S ribosomal RNA gene partial sequence

DQ819132.1 Uncultured bacterium clone aab26c04 16S ribosomal RNA gene partial sequence

KX950710.1 Staphylococcus lentus strain 4v 16S ribosomal RNA gene partial sequence


72

17

13

7

6

KT748639.1 Alcaligenes aquatilis strain C_6 16S ribosomal RNA gene partial sequence(2)

KJ513379.1 Alcaligenes faecalis strain E5.Zn 4 16S ribosomal RNA gene partial sequence

AB694007.1 Rhodobacter sphaeroides gene for 16S rRNA partial sequence strain: AMT-08

KT748636.1 Alcaligenes aquatilis strain C_11 16S ribosomal RNA gene partial sequence

KP274847.1 Alcaligenes sp. SAD2 16S ribosomal RNA gene partial sequence(2)

KU860464.1 Alcaligenes sp. AG3 16S ribosomal RNA gene partial sequence

KT808882.1 Alcaligenes aquatilis strain C_7 16S ribosomal RNA gene partial sequence

KY009932.1 Alcaligenes aquatilis strain 3c 16S ribosomal RNA gene partial sequence

11

10

10

9

2

(b)

(e)

(d)

(c)

53

Figure 12: Phylogenetic tree of all the selected bacterial isolates based on neighbor joining method

(a) S. lentus 1b, (b) S. lentus 1m, (c) B. subtilis, (d) S. lentus 4v, (e) A. aquatilis and (f) A. faecalis

(g) Alishwanella sp.

Figure 13: Phylogenetic tree of all the selected bacterial isolates based on neighbor joining method

in combined form.

AY994311.1 Alcaligenes sp. PAOAC171 16S ribosomal RNA gene partial sequence

KF054926.1 Rhodobacter sphaeroides strain IARI-NIAW1-7 16S ribosomal RNA gene partial sequence

AY346138.1 Alcaligenes sp. IS-17 16S ribosomal RNA gene complete sequence

KF383272.1 Alcaligenes faecalis strain Fa1.3 16S ribosomal RNA gene partial sequence

JX843599.1 Uncultured bacterium clone B171 16S ribosomal RNA gene partial sequence

KX118706.1 Alcaligenes faecalis strain O1R4 16S ribosomal RNA gene partial sequence

KX118704.1 Alcaligenes faecalis strain DEP8 16S ribosomal RNA gene partial sequence

KY009933.1 Alcaligenes faecalis strain 3d 16S ribosomal RNA gene complete sequence73

100

53

25

1

JQ012976.1 Alishewanella sp. 199 16S ribosomal RNA gene partial sequence

NR_125524.1 Alishewanella solinquinati strain NCIM 5295 16S ribosomal RNA gene partial sequence

GQ280101.1 Alishewanella sp. JS-31 16S ribosomal RNA gene partial sequence

HM104373.1 Alishewanella sp. AMP-7 16S ribosomal RNA gene partial sequence

JQ012977.1 Alishewanella sp. 197 16S ribosomal RNA gene partial sequence

JQ245619.1 Uncultured bacterium clone SYNH02_C3-18B-154 16S ribosomal RNA gene partial sequence

KM362999.1 Rheinheimera sp. AE24 16S ribosomal RNA gene partial sequence

JN546609.1 Alishewanella sp. SE-R 16S ribosomal RNA gene partial sequence45

45

8

8

7

KY009933.1 Alcaligenes faecalis strain 3d 16S ribosomal RNA gene complete sequence

KY009932.1 Alcaligenes aquatilis strain 3c 16S ribosomal RNA gene partial sequence

JN546609.1 Alishewanella sp. SE-R 16S ribosomal RNA gene partial sequence

KX950709.1 Bacillus subtilis strain 2p 16S ribosomal RNA gene partial sequence

KX950707.1 Staphylococcus lentus strain 1m 16S ribosomal RNA gene partial sequence

KX950710.1 Staphylococcus lentus strain 4v 16S ribosomal RNA gene partial sequence

KX950708.1 Staphylococcus lentus strain 1b 16S ribosomal RNA gene partial sequence

100

93

100

64

(g)

(f)

54

Optimization of bio decolorization conditions

The biodecolorization of azo dye by bacterial strains was checked under various conditions

i.e., temperature, pH, incubation condition, carbon and nitrogen source, inoculum size and dye

concentration to find out optimum conditions for dye decolorization.

S. lentus 1b showed maximum decolorization at 37 oC, pH 7 and under static condition. It

efficiently utilized wheat bran and yeast extract as carbon and nitrogen source respectively to show

optimum decolorization. The percentage of decolorization was maximum when inoculum size was

6% and dye concentration was 10 mg/L (Fig.14 a to g).

S. lentus 1m showed maximum decolorization percentage when grow statically at 37 oC

and pH 7. The rate of decolorization was maximum when wheat bran and yeast extract were used

as carbon and nitrogen source respectively. It efficiently decolorized dye when inoculum size was

4% and concentration of dye was 10 mg/L (Fig. 15 to g).

B. subtilis decolorized dye efficiently at 37 oC, pH 7 and under static condition. The rate of

decolorization was quite high when sugarcane bagasse and yeast extract were used as carbon and

nitrogen source respectively for its growth. The maximum rate of decolorization was determined

when inoculum size was 6% and dye concentration was 10 mg/L (Fig. 16 a to g).

S. lentus 4v decolorized dye efficiently at 37°C, pH 7 and under static condition. The rate

of decolorization was high when wheat bran and yeast extract were used as carbon and nitrogen

source respectively. It showed maximum decolorization when inoculum size was 4% and dye

concentration was 20 mg/L (Fig. 17 a to g).

A. aquatilis showed maximum decolorization at 37°C, pH 7 and at static condition. The

bacterium efficiently utilized saw dust and yeast extract as carbon and nitrogen source,

respectively to show maximum percentage decolorization ability. Moreover, efficient

55

decolorization was determined when inoculum percentage was 6 and dye concentration was 10

mg/L (Fig.18 a to g).

A. faecalis showed maximum decolorization rate at 37°C, pH 7 and at static condition. The

strain efficiently utilized saw dust and yeast extract as carbon and nitrogen source to show

optimum rate of decolorization. The percentage of decolorization was maximized when inoculum

size was 6% and dye concentration was 10mg/L (Fig. 19 a to g).


The bacterial strains were grown in a medium containing multiple azo dye i.e., Congo Red,

Methyl Red, Phenol Red, Brilliant Black and Navy Blue in order to check their potential to degrade

multiple dyes at the same time.

S. lentus 1b decolorized 81% of these dyes within a period of 5 days (Fig. 21a) while S.

lentus 1m showed a decolorization percentage of 79 (Fig. 21b). B. subtilis decolorized 82% percent

of the dye mixture (Fig. 21c) while S. lentus 4v showed a decolorization percentage of 87

(Fig.21d). A. aquatilis was able to decolorize mixture of dyes to a percentage of 86 (Fig. 21 e)

while A. faecalis was able to decolorize upto 82% (Fig.21 f).

Growth Decolorization relationship

It was observed that decolorization had a postive effect on the growth of the bacteria.

Inspite of the fact that decolorization suppressed the growth of bacteria it played its role in

increasing cell growth which is evident from the figure (Fig. 22 a to f ).

56

Figure 14: Optimization of decolorization condition for S. lentus 1b (a) temperature, (b) pH, (c)

incubation condition, (d) inoculum percentage, (e) carbon source, (f) nitrogen source and (g) dye

concentration.

0

20

40

60

80

25 30 37 45 50

Dec

olo

riza

tio

n (

%)

Temperature (◦C)

(a)

0

20

40

60

80

5 6 7 8 9 10

Dec

olo

riza

tio

n(%

)

pH

(b)

0

20

40

60

80

Static Shaking

Dec

olo

riza

tio

n (

%)

Incubation condition

(c)

0

20

40

60

80

2 4 6 8 10

Dec

olo

riza

tio

n (

%)

Innoculum size (%)

(d)

0

20

40

60

80

S.B S.D W.B

Dec

olo

riza

tio

n (

%)

Carbon source

(e)

0

20

40

60

80

B.extract Peptone Y.extract

Dec

olo

riza

tio

n (

%)

Nitrogen source

(f)

0

20

40

60

80

3 5 7 10 20 50

Dec

olo

riza

tio

n (

%)

Dye concentration (mg/L)

(g)

57

Figure 15: Optimization of decolorization condition for S. lentus 1m (a) temperature, (b) pH, (c)


concentration.

0

20

40

60

80

5 6 7 8 9 10

Dec

olo

riza

tio

n (

%)

pH

(b)

0

20

40

60

80

Static Shaking

Dec

olo

riza

tio

n (

%)


(c)

0

20

40

60

80

2 4 6 8 10

Dec

olo

riza

tio

n (

%)

Innoculum size (%)

(d)

0

20

40

60

80

S.B S.D W.B

Dec

olo

riza

tio

n (

%)

Carbon source

(e)

0

20

40

60

80


Dec

olo

riza

tio

n (

%)

Nitrogen source

(f)

0

20

40

60

80

25 30 37 45 50

Dec

olo

riza

tio

n (

%)

Temperature (◦C)

(a)

0

20

40

60

80

3 5 7 10 20 50

Dec

olo

riza

tio

n (

%)


(g)

58

Figure 16: Optimization of decolorization condition for B. subtilis (a) temperature, (b) pH, (c)


concentration.

0

20

40

60

80

Static Shaking

Dec

olo

riza

tio

n (

%)


(c)

0

20

40

60

80

S.B S.D W.B

Dec

olo

riza

tio

n (

%)

Carbon source

(e)

0

20

40

60

80


Dec

olo

riza

tio

n (

%)

Nitrogen source

(f)

0

20

40

60

80

3 5 7 10 20 50

Dec

olo

riza

tio

n (

%)


(g)

0

20

40

60

80

5 6 7 8 9 10

Dec

olo

riza

tio

n (

%)

pH

(b)

0

20

40

60

80

25 30 37 45 50

Dec

olo

riza

tio

n (

%)

Temperature (◦C)

(a)

0

20

40

60

80

2 4 6 8 10

Dec

olo

riza

tio

n (

%)

Innoculum size (%)

(d)

59

Figure 17: Optimization of decolorization condition for S. lentus 4v (a) temperature, (b) pH, (c)


concentration.

0

20

40

60

80

5 6 7 8 9 10

Dec

olo

riza

tio

n (

%)

pH

(b)

0

20

40

60

80

Static Shaking

Dec

olo

riza

tio

n (

%)


(c)

0

20

40

60

80

2 4 6 8 10

Dec

olo

riza

tio

n (

%)

Innoculum size (%)

(d)

0

50

100

S.B S.D W.B

Dec

olo

riza

tio

n (

%)

Carbon source

(e)

0

20

40

60

80

3 5 7 10 20 50

Dec

olo

riza

tio

n (

%)


(g)

0

20

40

60

80

25 30 37 45 50

Dec

olo

riza

tio

n (

%)

Temperature (◦C)

(a)

0

20

40

60

80


Dec

olo

riza

tio

n (

%)

Nitrogen source

(f)

60

Figure 18: Optimization of decolorization condition for A. aquatilis (a) temperature, (b) pH, (c)


concentration.

0

20

40

60

80

25 30 37 45 50

Dec

olo

riza

tio

n (

%)

Temperature (◦C)

(a)

0

20

40

60

80

5 6 7 8 9 10

Dec

olo

riza

tio

n (

%)

pH

(b)

0

50

100

Static Shaking

Dec

olo

riza

tio

n (

%)


(c)

0

50

100

2 4 6 8 10D

eco

lori

zati

on

(%

)

Innoculum size (%)

(d)

0

20

40

60

80

S.B S.D W.B

Dec

olo

riza

tio

n (

%)

Carbon source

(e)

0

20

40

60

80


Dec

olo

riza

tio

n (

%)

Nitrogen source

(f)

0

20

40

60

80

3 5 7 10 20 50

Dec

olo

riza

tio

n (

%)


(g)

61

Figure 19: Optimization of decolorization condition for A. faecalis (a) temperature, (b) pH, (c)


concentration.

0

20

40

60

80

25 30 37 45 50

Dec

olo

riza

tio

n (

%)

Temperature (◦C)

(a)

0

20

40

60

80

5 6 7 8 9 10

Dec

olo

riza

tio

n (

%)

pH

(b)

0

20

40

60

80

Static Shaking

Dec

olo

riza

tio

n (

%)


(c)

0

20

40

60

80

2 4 6 8 10D

eco

lori

zati

on

(%

)

Innoculum size (%)

(d)

0

20

40

60

80

S.B S.D W.B

Dec

olo

riza

tio

n (

%)

Carbon source

(e)

0

20

40

60

80

B.extract Peptone Y.extractDec

olo

riza

tio

n (

%)

Nitrogen source

(f)

0

20

40

60

80

3 5 7 10 20 50

Dec

olo

riza

tio

n (

%)


(g)

62

0

20

40

60

80

100

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Time (Days)

(a)

0

20

40

60

80

100

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Time (Days)

(b)

0102030405060708090

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Time (Days)

(c)

63

Figure 20: Potential of (a) S. lentus 1b, (b) S. lentus 1m, (c) B. subtilis, (d) S. lentus 4v, (e) A.

aquatilis and (f) A. faecalis to decolorize multiple dyes incubated at 37°C after suitable time

interval.

0

20

40

60

80

100

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Time (Days)

(d)

0

20

40

60

80

100

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Time (Days)

(e)

0

20

40

60

80

100

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Time (Days)

(f)

64

0

20

40

60

80

100

0

0.5

1

1.5

2

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Op

tica

l d

ensi

ty a

t 6

00

nm

Days

(a)

Growth

Decolorization

0

20

40

60

80

100

0

0.5

1

1.5

2

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Op

tica

l d

ensi

ty a

t 6

00

nm

Days

(b)

Growth

Decolorization

0

10

20

30

40

50

60

70

80

0

0.5

1

1.5

2

2.5

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Op

tica

l d

ensi

ty a

t 6

00

nm

Days

(c)

Growth

Decolorization

65

Figure 21: Effect of decolorization on growth of (a) S. lentus 1b, (b) S. lentus 1m, (c) B. subtilis,

(d) S. lentus 4v, (e) A. aquatilis and (f) A. faecalis under optimum growth conditions.

0

20

40

60

80

100

0

0.5

1

1.5

2

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Op

tica

l d

ensi

ty a

t 6

00

nm

Days

(d)

Growth

Decolorization

0

20

40

60

80

100

0

0.5

1

1.5

2

2.5

3

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Op

tica

l d

ensi

ty a

t 6

00

nm

Days

(f)

Growth

Decolorization

0

20

40

60

80

100

0

1

2

3

1 2 3 4 5 Dec

olo

riza

tio

n (

%)

Op

tica

l d

ensi

ty a

t 6

00

nm

Days

(e)

Growth

Decolorization

66

Consortium studies

Aerobic and anaerobic isolates were grown together, in the form of consortium, to study

their effect on dye degradation. Figure 22 a and b is showing that rate of decolorization was

increased by 8-15% (aerobic) and 10-14% (anaerobic) when dye containing sample was inoculated

with the consortium of bacterial isolates. Moreover, the mixed consortium (anaerobic and aerobic)

was more effective in decolorization of dye i.e., 8-10% as compared to the aerobic and anaerobic

consortium (Fig. 22c).

Figure 22: Effect of (a) Aerobic, (b) Anaerobic and (c) Mixed consortium on degradation of

dye.

0

50

100

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Time (Days)

(a)A. aquatilis

A. faecalis

Consortium

0

20

40

60

80

100

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Time (Days)

(b)S. lentus1b

S. lentus1m

B. subtilis

S. lentus4v

Consortium

0

50

100

1 2 3 4 5

Dec

olo

riza

tio

n (

%)

Time (Days)

(c)Aerobic

Anaerobic

Mixed

67


TLC

TLC was performed to analyze the dye degraded products. Bands of different Rf values

were observed in dye treated cultures when observed under UV light of 254nm. The control (dye)

band was observed with a Rf value of 0.94. In case of S. lentus 1b, Rf value of two bands were

0.89 and 0.95 (Fig. 23a) while in chromatogram of S. lentus 1m treated dye sample a band of Rf

value of single band was 0.89 (Fig.23b). The chromatogram of B. subtilis treated dye sample

showed two bands of Rf value 0.92 and 0.98 (Fig.23c) while S. lentus 4v showed two bands of Rf

value 0.90 and 0.96 (Fig.23d). The chromatograms of A. aquatilis and A. faecalis treated dye

sample showed bands of Rf value 0.95 (Fig. 23e) and 0.93 (Fig.23f) respectively.

Figure 23: TLC chromatogram of extracted metabolites of (a) S. lentus 1b, (b) S. lentus 1m, (c)

B. subtilis, (d) S. lentus 4v, (e) A. aquatilis and (f) A. faecalis decolorized dye sample visualized

under UV range of 254nm.

(C) (T) (C) (T) (C) (T) (C) (T) (C) (T) (C) (T)

(a) (b) (c) (e) (d) (f)

68

HPLC

The chromatogram of the dye sample showed three detectable peaks at retention time of

1.80, 2.88 and 5.00 min (Fig. 24a) while chromatogram of S.lentus strain 1b has two detectable

peaks of retention time 1.97 and 2.92 min (Fig. 24b) The chromatogram of dye sample treated with

S.lentus strain 1m has a detectable of retention time 3.01 min (Fig. 24c) while that of which treated

with B. subtilis has three peaks of retention time 1.92, 2.89 and 4.86 min (Fig. 24d). The

chromatogram of dye sample which was treated with S.lentus strain 4v showed four prominent

peaks of retention time 1.94, 2.91, 4.92 and 6.37 min (Fig. 24e). The chromatogram of dye sample

treated with A. aquatilis showed peaks at retention time of 1.99, 2.30, 2.95 and 3.76 min (Fig. 24f)

and that treated with A. faecalis has three detectable peaks of retention time 1.93, 2.09 and 2.95

min (Fig. 24g). The dye sample was incubated with these strains for a period of 4 days in order to

obtain maximum decolorization.

FTIR

A comparison between the FTIR spectrum of control dye and extracted metabolites is given in

figure 25 a to f. Two specific peaks of wavelength 1612 and 1532 cm-1 are shown in FTIR spectrum

of control dye which are due to the presence of -N=N- stretching. The peak with wavelength 1395

cm-1 indicates C-O-H bending. The peak with a wavelength 1037 cm-1 is due to C-O stretching.

There is a variation in peaks in the FTIR spectrum of metabolites extracted from bacterial

decolorized sample of dye when compared to the control dye spectrum. The absence of peaks with

wavelength 1612 and 1532 cm-1 indicates the reductive cleavage of azo bond. The peaks with

wavelength 2850 and 2923 cm-1 in S. lentus 1b, B. subtilis, S. lentus 4v and A. aquatlis, 2852 and

2926 cm-1 S. lentus 1m and 2923 cm-1 in A. faecalis is due to stretching of alkanes. The peaks with

1755 and 1717 cm-1 wavelength in above mentioned organisms indicate the presence of C=O

69

group. A peak of wavelength 1289 cm-1 depicts the presence of carboxyl group. The peaks with

wavelength 1104, 1108 and 1037 cm-1 show the presence of amines (C-N). The presence of peak

with wavelength 769 cm-1 indicates the presence of conjugated C=C.

(a)

(b)

)

70

(c)

(d)

71

(e)

(f)

72

Figure 24: HPLC analysis of reactive Synazol red HF-6BN with a mobile phase of profile of

methanol at 1.0 mL/min. The column was C-18 (250×4.6 mm) (a) represents dye chromatogram

while (b) represents S. lentus 1b, (c) S. lentus strain 1m, (d) B. subtilis, (e) S. lentus strain 4v, (f)

A. aquatilis and (g) A. faecalis degraded dye products extracted after 4 days of incubation at 37°C.

(g)

(a)

(g)

73

(b)

(c)

74

(d)

(e)

75

Figure 25: FTIR spectrum of control dye (Synazol red HF-6BN) with (a) S. lentus 1b, (b) S. lentus

1m, (c) B. subtilis, (2p) (d) S. lentus 4v, (e) A. aquatilis (3c) and (f) A. faecalis (3d).

GC-MS

GC-MS was performed to identify the degraded products of the dye. The results

showed that the predominant compounds found in the final dye treated sample were 3-

Aminobutanoic acid, pyrrolo pyrazine-1, 4-dione and palmitic acid. The chromatogram of all the

organisms were given in the figure (Fig. 26 a to g).

(f)

76

(a)

(b)

77

(c)

(d)

78

(e)

(f)

79

Figure 26: GC-MS chromatograms of (a) control dye (b) S. lentus 1b, (c) S. lentus 1m, (d) B.

subtilis, (2p) (e) S. lentus 4v, (f) A. aquatilis (3c) and (g) A. faecalis (3d).

Dye degradation at large scale

The bacterial isolates efficiently degraded dye in industrial effluent when estimated after

2, 4, 6 and 8 days of incubation at room temperature. The dye decolorization was also visible by

change in color of experimental flasks when compared with the control containing no bacterial

culture (Fig. 27 a to f). S. lentus 1b decolorized dye by 77% within a period of 8 days (Fig. 28a)

while S. lentus 1m by 71% (Fig. 28b). B. subtilis efficiently decolorized dye by 70% (Fig.28c) and

S. lentus 4v by 71% (Fig.28d). A. aquatilis and A. faecalis decolorized dye to a percentage of 72

and 68, respectively (Fig. 28 e and f).

(g)

80

aaaaa

Figure 27: Change in color of dye with respect to control in flasks containing culture of bacterial

isolates (a) S. lentus 1b, (b) S. lentus 1m, (c) B. subtilis (2p), (d) S. lentus 4v, (e) A. aquatilis (3c)

and (f) A. faecalis (3d) from the original industrial wastewater.

(a) (b)

(c)

(d)

(c)

(f)

(e)

81

0

20

40

60

80

100

2 4 6 8

Dec

olo

riza

tio

n (

%)

Time (days)

(a)

negative control

positive control

S. lentus1b

0

10

20

30

40

50

60

70

80

2 4 6 8

Dec

olo

riza

tio

n %

Time (days)

(b)

negative control

positive control

S. lentus strain 1m

0

20

40

60

80

2 4 6 8

Dec

olo

riza

tio

n (

%)

Time (days)

(c)

negative control

positive control

B. subtilis

82

Figure 28: Decolorization of Synazol red HF-6BN by bacterial isolates (a) S. lentus 1b (b) S.

lentus strain 1m, (c) B. subtilis (2p), (d) S. lentus strain 4v, (e) A. aquatilis (3c) and (f)

A. faecalis (3d) from industrial wastewater after incubation of 2, 4, 6 and 8 days of

incubation at room temperature.

0

20

40

60

80

100

2 4 6 8

Dec

olo

riza

tio

n (

%)

Time (days)

(d)

negative control

positive control

S. lentus strain 4v

0

20

40

60

80

2 4 6 8

Dec

olo

riza

tio

n (

%)

Time (days)

(e)

negative control

positive control

A. aquatilis

0

20

40

60

80

2 4 6 8

Dec

olo

riza

tio

n (

%)

Time (days)

(f)

negative control

positive control

A. faecalis

83

Microbial toxicity

The bio decolorized dye sample was checked for its effect on microbial flora. There was

no zone of inhibition found around the well having decolorized dye sample which clearly indicated

that the biodegraded wastewater is not harmful for the growth of microbial species (Fig. 29 to 35).

Figure 29: The broth decolorized by S. lentus 1b used for incubation of (a) B. megaterium, (b) B.

subtilis and (c) B. cereus at 37°C for 48 hours. No zone of inhibition was appeared in any case.

Figure 30: The broth decolorized by S. lentus 1m used for incubation of (a) B. megaterium, (b) B.


(a)

B. megaterium

(c)

B. cereus

(b)

B. subtilis

(a)

B. megaterium

(b)

B. subtilis

(c)

B. cereus

84

Figure 31: The broth decolorized by B. subtilis used for incubation of (a) B. megaterium, (b) B.


Figure 32: The broth decolorized by S. lentus 4v used for incubation of (a) B. megaterium, (b) B.


Figure 33: The broth decolorized by A.aquatilis used for incubation of (a) B. megaterium, (b) B.


(a)

B. megaterium

(b)

B. subtilis

(c)

B. cereus

(a)

B. megaterium

(b)

B. subtilis

(c)

B. cereus

(a)

B. megaterium

(b)

B. subtilis

(c)

B. cereus

85

Figure 34: The broth decolorized by A. faecalis used for incubation of (a) B. megaterium, (b) B.


Figure 35: Alishewanella sp. decolorized broth used for incubation of (a) B. megaterium, (b) B.

subtilis and (c) B. cereus at 37°C for 48 h. No zone of inhibition was observed in any case.

Phytoxicity

A clear difference in growth of control and experimental plants was observed (Fig. 36 a

to f). Experimental pots showed fair growth as they watered with wastewater in which dye was

graded by the action of bacteria. Those pots watered with untreated wastewater showed less

growth.

(a)

B. megaterium

(b)

B. subtilis

(c)

B. cereus

(a)

B. megaterium

(b)

B. subtilis

(c)

B. cereus

86

(a)

(b)

87

(c)

(d)

88

Figure 36: Use of treated wastewater by (a) S. lentus 1b, (b) S. lentus strain 1m, (c) B. subtilis

(2p), (d) S. lentus strain 4v, (e) A. aquatilis (3c) and (f) A. faecalis (3d)., tap water and untreated

wastewater for growth of Vigna radiata (mung beans) for a 10 days incubation period under 1:1

light and dark conditions. Two pots were used for each treatment (n=2).

(e)

(f)

89

DISCUSSION

Variations in natural and anthropogenic activities causes global pollution which

contaminates various terrestrial and aquatic ecosystems with heavy metals, inorganic and organic

compounds and radionuclide. The controlled and uncontrolled solid and liquid discharge from

industries, accidental spillages, use of agricultural fertilizers, herbicides, insecticides and

pesticides, sewage disposal mainly contributes to increase pollution at an alarming rate.

The discharge produced by textile industries is a major issue of today’s world as it become

a major source of pollution in water bodies. Several types of dyes are used in textile industries and

among them azo dyes are the most toxic ones because of their complex structure which mainly

involves aromatic rings, azo bonds and amino groups. Moreover, their degradation products are

also toxic in nature. The physiochemical techniques used to degrade these dyes such as adsorption,

coagulation- flocculation, oxidation and electrochemical methods are unable to degrade dyes

completely (Lin and Peng, 1994). Therefore, processing of these dyes using microorganisms is a

modern-day approach to deal with this problem.

The present research deals with the isolation and characterization of dye degrading bacteria

and their potential in the bioremediation of dye contaminated environment. For the isolation of

dye degrading microorganisms, wastewater samples were collected from the industrial area of

Sheikhupura road and Lahore. Microorganisms which efficiently decolorize dyes are the ones

which are isolated from dye contaminated sites such as textile effluents, soil nearby the

contaminated sites and sludge obtained after the treatment of wastewater (Yang et al., 2009; Ola

et al., 2010; Ayed et al., 2011).

90

A total of 7 isolates which degrade dye efficiently were finalized for further study. Four of

them are anaerobic while the rest of three are aerobic. Both aerobic and anaerobic bacterial strains

can decolorize dyes, but dyes are resistant to degradation under aerobic environment as the

presence of oxygen inhibits the reduction of azo bond (Solis et al., 2012). The isolate 1b, 1m and

4v were Gram positive cocci, 2p was Gram positive rod while 3c, 3d and CBL-2 were Gram

negative rods. Various groups of bacteria are studied to investigate their dye degrading ability.

Both Gram positive and negative bacteria are equally efficient in degrading dyes (Sani and

Banerjee 1999; Kodam et al., 2005; Moosvi et al., 2005).

Optimization of bio decolorization conditions

The process of degradation of azo dyes is a complex one which relies on numerous factors.

The limitation of these factors greatly influences the rate of decolorization. In the present study

numerous factors were evaluated to optimize the rate of decolorization by the isolated bacteria.

These factors include temperature, pH, incubation condition, carbon and nitrogen source, inoculum

percentage and dye concentration.

The growth as well as enzymatic activity of microorganisms is influenced by temperature.

Therefore, biodegradation of azo dyes is also affected by temperature. All the bacterial isolates

used in this study decolorized dye at optimum rate at 37oC. Guadie et al. (2017) reported a Bacillus

sp. strain CH12 which decolorized dye at optimum rate between 25 to 35oC. Micrococccus strain

R3 was able to decolorize methyl red at 37oC (Olukanni et al., 2009).

The extent of decolorization is also influenced by the pH of the medium. It also effects the

solubility and color of the dye. All the bacterial isolates used in this study decolorized dye at

optimum rate at pH 7. An increase in the rate of decolorization was noted from pH 5-7 while it

http://www.sciencedirect.com/science/article/pii/S0960852406004184#bib22




91

was decreased with further increase in pH from 8-10. The decolorization rate of methyl red by

Micrococcus strain R3 within a pH range of 6-8 was 84.6%, 94.19%, and 93.94%, respectively

(Olukanni et al., 2009). Pseudomonas sp. SUK 1 was able to decolorize a mixture of azo dyes

(Remazol Orange 3R, Scarlet RR, Brown 3REL, Golden yellow HER, Remazol Red, Cotton Blue,

Amaranth, Orange 2RX) efficiently at pH 7 (Chougule et al., 2014).

The incubation condition i.e., static or shaking greatly effects the rate of decolorization as

the presence of oxygen competes with nitrogen for the process of reduction. In the present case,

aerobic strains i.e., A. aquatilis and A. faecalis showed optimum decolorization under shaking

conditions while anaerobic strains i.e., S. lentus 1b, S. lentus 1m, B. subtilis and S lentus 4v

decolorized well under static condition. Pseudomonas sp. SUK 1 was able to decolorize a mixture

of azo dyes (Remazol Orange 3R, Scarlet RR, Brown 3REL, Golden yellow HER, Remazol Red,

Cotton Blue, Amaranth, Orange 2RX) efficiently under static conditions (Chougule et al., 2014).

Carbon and nitrogen source also plays a vital role in the process of decolorization. They

not only provide the energy but also serves as an electron donor for the reduction of azo bond.

Furthermore, the size of the inoculum also effects the rate of decolorization. The studied bacteria

showed increased rate of decolorization by the addition of carbon and nitrogen source.

Shah et al. (2013) reported that Bacillus species i.e., B. cereus and B. megaterium having

95% and 98% dye decolorizing potential respectively under optimum conditions. Optimal

condition for B. cereus was found to be 37 oC, pH 7, 1% sucrose, 0.25% peptone and 8% inoculum

and that for B. megaterium was found to be 37°C, pH 6, glucose 1%, 0.25% yeast extract and 10%

inoculum.

The concentration of dyes in the textile effluent ranges from 16-20 mg/L (Banat et al.,

1996). The dye concentration influences the rate of decolorization as well. The bacteria isolated

92

in the present study can decolorize dye within the range of 20 -50 mg/L. Bacillus sp. strain CH12

had the ability to decolorize lower dye concentrations (50-100mg/L) with ≥ 96% decolorization

rate while higher concentrations (150-250mg/L) with 84-95% decolorization rate (Guadie et al.,

2017). Saha et al. (2017) reported two Alcaligenes faecalis species namely A. faecalis E5.Cd and

A. faecalis Fal.3 which decolorize upto 93% of Blue H/C and Red 3B dye at pH 7 and 94% of

yellow 3R at pH 8 within a period of 96 h. Both the strains decolorize upto 91% of dye at 35°C,

92% with 50 mg/L initial dye concentration and 93% with 20% inoculum size and supplementation

of 1% co-substrate, respectively.


The current study revealed that the anaerobic strains have the potential to degrade mixture

of multiple dyes uptil 87% while that of aerobic strains is 86%. Chougule et al. (2014) reported a

strain named Pseudomonas sp. SUK1 which was able to decolorize the mixture of various dyes

(Remazol Orange 3R, Scarlet RR, Brown 3REL, Golden yellow HER, Remazol Red, Cotton Blue,

Amaranth, Orange 2RX) by 94.3%. Shah (2014) reported three bacteria namely P. putida, P.

aeruginosa and B. subtilis capable of degrading multiple dyes i.e, Blue RR, Black B, Red RR,

Yellow RR and Navy blue. P. putida showed maximum decolorization of Blue RR (95%), P.

aeruginosa of Black B (93%) and Navy Blue (70.58%) and B. subtilis of Yellow RR (65%) and

Red RR (91%). A bacterium Clostridium biofermentans which was able to decolorize different

dyes such as Reactive red 3B-A, Reactive black 5, and Reactive yellow 3B-A, by over 90% within

a period of 36 hours (Joe et al., 2008).

Growth decolorization relationship

The azo dye molecules are deficient in carbon (Levin et al., 2010; Asgher et al., 2008)

which serves as source of energy for growth of microorganisms as well as an electron donor for

93

the reduction of azo bond (Yemendzhiev et al., 2009; Yang et al., 2009). However, some

researches have been done to make it possible that microorganisms would rely on dye as a sole

source of carbon to make this process of biodegradation more economical. In the present study we

tried the same thing and it was observed that the bacterial growth flourish as the biodegradation

process proceeded. Saratale et al., (2009a) demonstrated that M. glutamicus was able to grow by

using Navy Blue HE2R and its reaction intermediates as sole source of carbon.

Consortium studies

The azo dyes are generally degraded into aromatic amines by the action of microorganisms.

These amines are more toxic and difficult to degrade as compared to their parent molecule.

Moreover, a single microorganism is specific for a single dye. There is a need of complex

enzymatic system to degrade the by-products of dyes. So there arises the need of development of

microbial consortium to completely mineralize the dye (Phugare et al., 2011).

In the present study consortium of aerobic as well as anaerobic strains was created in order

to observe the difference in decolorization efficiency of individual strains and their consortium.

About 8-15% increase in decolorization rate was observed by aerobic consortium as compared to

the individual aerobic strains. Anaerobic consortium showed 10-14% increase in decolorization

rate in comparison to individual anaerobic strains.

Bose and Rajasekaran (2016) developed a bacterial consortium comprising of Klebsiella

sp1and sp2, S. aureus, B. cereus and P. fluorescence which was able to decolorize various dyes at

different rates. The degraded yellow, green, orange blue and black dye by 63%, 92%, 40%, 65%

and 96% respectively. The increased decolorization rate was due to the synergistic effect of

94

microorganisms. The biocatalytic activity of microorganisms is enhanced when they are present

in the form of a consortium.

The combination of anaerobic and aerobic consortium makes a perfect system for the

degradation of azo dyes as reductive cleavage of azo bone is done under anaerobic conditions

(Yemashova et al., 2009) and resulting aromatic amines are completely mineralized under aerobic

conditions (Forss and Welandar, 2011; Koupaie et al., 2011; Jonstrup et al., 2011).

In the present study anaerobic and aerobic consortium were simultaneously applied for

complete mineralization of azo dye. An increase of 8-10% in the final decolorization rate was

observed in comparison to individual anaerobic and aerobic consortium. Another consortium

reported by Lade et al., (2015) comprising of Providencia rettgeri strain HSL1 and Pseudomonas

sp. SUK1 was able to decolorize structurally different azo dyes under microaerophilic and aerobic

conditions. It was able to decolorize 98-99% of various dyes including Reactive Black 5, Reactive

Orange 16, Disperse Red 78 and Direct Red 81 at concentration of 100 mg/L within a period of 12

to 30 hours.

The degradation of Acid Red 18 was done by using anaerobic system followed by aerobic

system. This results in the 98% decolorization rate and more than 80% COD reduction which

occurred under anaerobic conditions. Upto 80% removal of by-product of anaerobic process (1-

naphthylamine-4-sulphonate) was done under aerobic process confirmed through HPLC (Koupaie

et al., 2011).

95


TLC

The analysis of degraded products of dye was done by using various techniques. Firstly,

this was done by thin layer chromatography. Rf value of the control band and dye band was

different in case of each bacterium. These results predict that original dye was broken down into

some other products when treated with bacterial strains. M. luteus strain SSN2 was used to degrade

Direct Orange 16 and when the degraded sample was analyzed through TLC two bands of Rf value

0.63 and 0.965 were detected under UV light of 254 nm while in control dye there is only one band

with Rf value 0.74 (Singh et al., 2015). The TLC chromatogram of degraded sample of Direct Red

by Enterococcus faecalis YZ66 had a band with Rf value of 0.71 while in control dye the Rf value

of band was 0.97 (Sahasrabudhe et al., 2014). Ghodake et al. (2011) reported three bands of Rf

value 0.93, 0.82 and 0.79 in TLC chromatogram of degraded sample of mono azo dye Amarnath

by Acinetobacter calcoaceticus NCIM 2890 but only a single band of Rf value 0.83 was produced

by control dye. Ajaz et al. (2018) reported the degraded of Sumifex Tourqi blue as new bands of

different Rf value was appeared in chromatogram of treated sample.

HPLC

To further confirm that the dye was degraded into its respective metabolites, HPLC was

performed. HPLC worked on the principal of relative affinity for mobile phase and stationary

phase. The compounds having greater affinity for stationary phase take longer time to pass through

the column or vice versa. The peaks of different retention time were observed in control and dye

sample treated with bacteria. This confirms the breakdown of parent dye into by products.

96

The degraded sample of three of anaerobic strains i.e., S. lentus 1b, B. subtilis and S. lentus

4v gave major peaks with retention time of comparable values i.e., 2.92, 2.89 and 2.91 respectively.

One anaerobic strain which was S. lentus strain 1m, its degraded sample gave the major peak with

retention time of 3.01. The major peaks of degraded sample of two aerobic strains i.e., A. aquatilis

and A. faecalis were of same retention time i.e.,2.95.

Lade et al. (2015) observed three major peaks with retention time 2.521, 3.241 and 3.564

and two minor peaks of retention time 3.123 and 3.910 when extracted metabolites from

Provedencia rettgeri decolorized C.I reactive blue 172 were analyzed through HPLC. The HPLC

analysis of the control dye results in the presence of one major peak with retention time of 2.702

min three minor peaks at retention time of 2.125, 2.801 and 3.394 min. HPLC analysis

demonstrated the presence of two major peaks at retention time 2.45 and 2.68 and two minor peaks

at retention time 3.50 and 6.50 min when B. halodurans decolorized Acid black- 24. The

chromatogram of original dye had only one peak at retention time of 2.81 min (Prasad and Rao.,

2014).

FTIR

In order to detect the presence of various groups in the degraded products of the dye FTIR

was performed. FTIR works on the principle that molecules have the capacity to absorb light at

various frequencies of infra-red region of spectrum of light. Each bond present in the molecule has

the capacity to absorb light of specific frequency. The measure of frequency is done in form of

wave number. The measure of frequency is usually done between the range of 4000 – 600 cm-1.

The spectrum obtained as result of absorption of light in the infra-red region describes the presence

of different functional groups and chemical bonds in the analyzing sample.

97

A comparison between the FTIR spectrum of control dye and extracted metabolites was

given in figure 25. The presence of specific peaks of wavelength 1612 and 1532 cm-1 in the

spectrum of dye which indicates the presence of -N=N- stretching in the control dye are not seen

in the spectrum of degraded metabolites which reflects the breakdown of azo bond.

Devi et al. (2018) reported a bacterium B. cereus isolated from contaminated soil present

near the vicinity of textile effluent. It was involved in the biodegradation of azo dye called Remazol

Red RB. FTIR spectrum before and after the degradation process showed a marked difference.

Stretching of C-N amine and alkene group was confirmed by the presence of peaks at 1040.44 cm-

1 and 954.80 cm-1. Moreover, O-H stretching of acids was confirmed by a peak at 2929.13 cm-1.

The degradation of disperse textile dye Brown 3REL by another Bacillus sp. was further

confirmed by FTIR spectrum (Dawker et al., 2008). The presence of a peak at 1458 cm-1 in the

spectrum degraded metabolites showed the C-H asymmetric deformation of alkane while the peak

at the frequency of 1667 cm-1 represented the C=N stretching which results the release of amine

group.

S. saprophyticus BHUSS X3 a bacterium isolated from the soil found in the vicinity of

carpet industry was efficiently able to decolorize a widely used acidic azo dye in the carpet industry

Navy N5RL1 (Kumari et al., 2015). The absence of peak with frequency of 1598.93 cm-1 which is

due to presence of aromatic nitrogen compound in azo dye spectrum clearly represents the

degradation of azo bond. The O-H stretching was confirmed by the presence of peak at frequency

of 3230 cm-1. The peak with the frequency of 1667 cm-1 represented the C=N stretching. The peak

with the frequency of 1455 cm-1 indicated the asymmetric deformation in the structure of alkanes

which usually results in the formation of oximes and amines or nitroso compounds by the

rearrangement of oximes.

98

The absence of peak of wavelength 1631 cm-1 in FTIR spectrum of Marinobacter sp. strain

HBRA treated Direct Blue-1 sample as compared to the spectrum of control dye clearly

demonstrated the decolorization of azo bond (Prasad et al., 2013). Shyamala et al., 2014 reported

the breakdown of Methyl Orange by halotolerant Bacillus sp. as peaks of wavelength 1567 and

1424 cm-1 which were present in FTIR spectrum of control dye were absent in spectrum of

bacterially treated sample.

GC-MS

It is a combination of two techniques i.e., gas chromatography and mass spectroscopy. It

is used for the identification of different substances within a sample at molecular level. The heating

of a testing substance degrades it into its products which is then mixed with an inert gas (helium).

Then they flow into MS where their identification is done on the basis of their mass.

Hungerer et al. (1996) reported that the azo-dye degraded compound 1,2- benzene dicarboxylic

acid, butyl 8-methylnonyl ester was cleaved to form phthalic acid and palmitic acid. Yang et al.,

(2018) reported the identification of degradation products of Direct Blue 15 dye by GC-MS. The

products were identified as 1- amino-8-napthol, 3, 3ʹ-dimethylbenzidine, anti-pentenoic acid, 2-

amino phenol and phenol.

Our findings are similar with other studies that reported that the predominant compounds found

in the final dye treated sample were 3-Aminobutanoic acid, pyrrolo pyrazine-1, 4-dione and

palmitic acid (Harwood and Parales, 1996; Walker and Van der, 2016; Shanmugam et al., 2017).

In the current research, it is confirmed through GC-MS analysis that azo dye enzymatically

converted into various end products which may be involved in the synthesis of energy yielding

processes.

99

The proposed metabolic pathway demonstrating the degradation of Synazol Red HF-6BN into

metabolites by A. faecalis and S. lentus 1m is given in the figure 37and 38 respectively.

Microbial Toxicity

The textile effluent is discharged into water bodies, so it also become a cause of damaging

agriculturally important microbial flora. The biodegraded products of azo dyes were checked for

their toxicity for agriculturally important microbial flora. The results showed that the biodegraded

products were not harmful for various tested agriculturally important microorganisms.

Ilyas and Rehman (2013) reported two fungal species i.e., Aspergillus niger and Nigrospora

sp. whose decolorized sample was proved safe for the microbial species. The microbial toxicity of

dye degraded products was also checked by Gomaa (2016). The degraded products of various dyes

produced by B. subtilis, B. cereus, B. licheniformis and Pseudomonas sp. were proved safe

for microbial flora as no zone of inhibition was observed in microbial toxicity plate assay.

The degradation products of Sudan 1 by B. circulans BWL1061 and S. putrefaciens CN32 showed

a decreased toxicity to E. coli BL21 and B. subtilis 168 (Liu et al., 2018).

Phytotoxicity

We are living in a country which is having depleted water resources. The need of

time is that not only to conserve natural water resources but also make it possible to bring

in use the microbially treated wastewater. For this purpose, the toxicity of treated

wastewater was checked for plants. The plants watered with treated waste water showed

efficient growth as compared to those watered with untreated wastewater (Fig. 39).

Nair et al. (2017) produced the same results as they used the B. megaterium treated

wastewater for growing Vigna radiata. The plants watered with treated wastewater showed fair

growth in comparison to those watered with untreated wastewater.

100

Figure 37: Synazol Red HF 6BN dye enters into the cell (A. aquatilis 3c) by unknown mechanism.

Upon entrance into the cell the dye is enzymatically processed into various end products. Most

probably, azo group (-N=N-) is reduced, followed by cleavage reaction and different end products

are produced. Secondly, desulfonation and oxidative deamination results in synthesis of

pyrrolo[1,2-a] pyrazine-1,4-dione derivative which can be used as substrates in amino acid

metabolism. The amino acid catabolism can synthesize pyruvate (3C compound) which can be

converted into acetyl-CoA. The acetyl-CoA undergoes Krebs cycle to produce NADH2 and

FADH2 (substrates of electron transport chain). Moreover, dye desulfonation, oxidative

deamination and carboxylation lead to produce phthalate derivatives, which can be transformed

into different fatty acids and aldehydes. The phthalate, fatty acids and aldehydes can

directly/indirectly enter into fatty acid oxidation reactions (β-oxidation) to produce acetyl-CoA,

NADH2 and FADH2 (Sarkar et al., 2017).

101

Figure 38: Synazol Red HF 6BN dye enters into the cell (Staphyloccocus lentus 1M) by unknown

mechanism. Upon entrance into the cell the dye is enzymatically processed into various end

products. Most probably, azo group (-N=N-) is reduced, followed by cleavage reaction and

different end products are produced. Secondly, desulfonation and oxidative deamination

eventually results in synthesis of 4-guanidinobutanoic acid derivative which can be used as

substrates in amino acid metabolism. The amino acid catabolism can synthesize pyruvate (3C

compound) which can be converted into acetyl-CoA. The acetyl-CoA undergoes Krebs cycle to

produce NADH2 and FADH2 (substrates of electron transport chain). Moreover, dye desulfonation,

oxidative deamination and carboxylation lead to produce phthalate derivatives, which can be

transformed into different fatty acids and aldehydes. The phthalate, fatty acids and aldehydes can

directly/indirectly enter into fatty acid oxidation reactions (β-oxidation) to produce acetyl-CoA,

NADH2 and FADH2 (Parmar and Shukla, 2018).

102

Figure 39: Flow diagram demonstrating the microbial treatment of textile effluent for dye

degradation and its use of treated for the growth of plants.

Textile

effluent

(Lahore)

103

REFERENCES

Acuner, E. and Dilek, F. B., 2004. Treatment of tectilon yellow 2G by Chlorella vulgaris. Process

aBiochem., 39: 623-631.

Aghaie-Khouzani, M., Forootanfar, H., Moshfegh, M., Khoshayand, M. R. and Faramarzi, M. A.,

2012. Decoloration of some synthetic dyes using optimized culture broth of laccase

producing ascomycete Paraconiothyrium variabile. Biochem. Eng. J., 60: 9–15.

Ajaz, M., Elahi, A. and Rehman, A., 2018. Degradation of azo dye by bacterium, Alishewanella

sp. CBL-2 isolated from industrial effluent and its potential use in decontamination of

wastewater. J. Water Reuse Desalination, 8(4): 507-515.

Aksu, Z. and Donmez, G. A., 2003. Comparative study on the biosorption characteristics of some

yeast for Remazol blue reactive dye. Chemosphere, 50: 1075–1083.

Aksu, Z. and Donmez, G., 2005. Combined effects of molasses sucrose and reactive dye on the

growth and dye bioaccumulation properties of Candida tropicalis. Process Biochem., 40:

2443-2454.

Aksu, Z., Kilic, N. K., Ertugrul, S. and Donmez, G., 2007. Inhibitory effects of chromium (VI)

and dyestuff removal by Trametes versicolor. Enzyme Microbiol. Technol., 40: 1167 -

1174.

Ambrosio, S. T., Vilar, Junior, J. C., da Silva, C. A. A., Okada, K., Nascimento, A. E., Longo, R.

L, Campos-Takaki, G. M., 2012. A biosorption isotherm model for the removal of reactive

azo dyes by inactivated mycelia of Cunninghamella elegans UCP542. Molecules, 17: 452–

462.

104

Amoozegar, M. A., Hajighasemi, M., Hamedi, J., Asad, S. and Ventosa, A., 2011. Azo dye

decolorization by halophilic and halotolerant microorganisms. Ann. Microbiol., 61: 217–

230.

Ang, E. L., Zhao, H. and Obbard, J. P., 2005. Recent advances in the bioremediation of persistent

organic pollutants via biomolecular engineering. Enzyme Microbiol. Technol., 37: 487–

496.

Anjaneya, O., Souche, S. Y., Santoshkumar, M. and Karegoudar, T. B., 2011. Decolorization of

sulfonated azo dye Metanil Yellow by newly isolated bacterial strains: Bacillus sp. strain

AK1 and Lysinibacillus sp. strain AK2. J. Hazard. Mater., 190: 351–358.

Asgher, M., Bhatti, H. N., Ashraf, M. and Legge, R. L., 2008. Recent developments in

biodegadation of industrial pollutants by white rot fungi and their enzyme system.

Biodegrad., 19: 771–783.

Asgher, M., Bhatti, H. N., Shah, S. A. H., Asad, M. J. and Legge, R. L., 2007. Decolorization

potential of mixed microbial consortia for reactive and disperse textile dyestuffs.

Biodegrad., 18: 311–316.

Asgher, M., Kausar, S., Bhatti, H. N., Shah, S. A. H. and Ali, M., 2008. Optimization of medium

for decolorization of Solar golden yellow R direct textile dye by Schizophyllum commune

IBL-06. Int. Biodeter. Biodegrad., 61: 189–193.

Axelsson, J., Nilsson, U., Terrazas, E., Alvarez, T. and Welander, U., 2006. Decolorization of the

Ayed, L., Khelifi, E., Jannet, H. B, Miladi, H., Cheref, A., Achour, S. and Bakhrouf, A., 2010.

Response surface methodology for decolorization of azo dye Methyl Orange by bacterial

consortium: produced enzymes and metabolites characterization. Chem. Eng. J., 165: 200–

208.

105

Ayed, L., Mahdhi, A., Cheref, A. and Bakhrouf, A., 2011. Decolorization and degradation of azo

dye Methyl Red by an isolated Sphingomonas paucimobilis: biotoxicity and metabolites

characterization. Desalination, 274: 272–277.

Azizullah, A., Khattak, M. N. K., Richter, P. and Häder, D., 2010. Water pollution in Pakistan and

its impact on public health — A review. aa 37: 479-497.

Babu, B. R., Parande, A. K., Raghu, S., and Kumar, T. P., 2007. Cotton textile processing: waste

generation and effluent treatment. J. Cotton Sci., 11: 141–153.

Bakshi, D. K., Saha, S., Sindhu, I. and Sharma, P., 2006. Use of Phanerochaete chrysosporium

biomass for the removal of textile dyes from a synthetic effluent. World J. Microbiol.

Biotech., 22: 835–839.

Baldrian, P., 2006. Fungal laccases – occurrence and properties. FEMS Microbiol, Rev., 30: 215–

242.

Banat, I. M., Nigam, P., Singh, D. and Merchant, R., 1996. Microbial decolorization of textile-dye

containing effluent: A review. Bioreour. Technol., 58(3): 217-227.

Barragan, B. E., Costa, C. and Marquez, M. C., 2007. Biodegradation of azo dyes by bacteria

inoculated on solid media. Dyes Pigments, 75: 73–81.

Behnajady, M. A., Modirshahla, N. and Shokri, M., 2004. Photodestruction of acid orange 7 (AO7)

in aqueous solutions by UV/H2O2: influence of operational parameters. Chemosphere, 55:

129-134.

Bhatnagar, A. and Sillanpaa, M., 2010. Utilization of agro-industrial and municipal waste

materials as potential adsorbents for water treatment: a review. Chem. Eng. J., 157: 277–

296.

106

Bose, P. and Rajasekaran, A., 2016. Decolourization of textile dyes using bacterial consortium.

Int. J. Sci. Res. Environ. Sci., 4(1): 17-22.

Burger, S. and Stolz, A., 2010. Characterisation of the flavin-free oxygen-tolerant azoreductase

from Xenophilus azovorans KF46F in comparison to flavin-containing azoreductases.

Appl. Microbiol. Biotechnol., 87: 2067–76.

Camarero, S., Ibarra, D., Martinez, M. J. and Martinez, A. T., 2005. Lignin-derived compounds as

efficient Laccase mediators for decolorization of different types of recalcitrant dyes. Appl.

Environ. Microbiol., 71: 1775–1784.

Cappuccino, J. G. S., Cappuccino, N. J. G. and Sherman, N. 1996. Microbiology: a laboratory

manual.

Chacko, J. T. and Subramaniam, K., 2011. Enzymatic degradation of azo dyes: a review. Int. J.

Environ. Sci., 1: 1250–1260.

Chan, G. F., Rashid, N. A. A., Koay, L. L., Chang, S.Y. and Tan, W. L., 2011. Identification and

optimization of novel NAR-1 bacterial consortium for the biodegradation of Orange II.

Insight Biotechnol., 1: 7–16.

Chang, J. S. and Kuo, T. S., 2000. Kinetics of bacterial decolorization of azo dye with Escherichia

coli NO3. Bioresour. Technol., 75: 107-111.

Chang, J., Chou, Y., Lin, P., Lin, J. and Hu, T. L., 2001. Kinetic characteristics of bacterial azo

dye decolorization by Pseudomonas luteola. Water Res., 35: 2841–2850.

Charumathi, D. and Das, N., 2012. Packed bed column studies for the removal of synthetic dyes

from textile wastewater using immobilised dead C. tropicalis. Desalination, 285: 22–30.

Charumathi, D. and Nilanjana, D., 2010. Bioaccumulation of synthetic dyes by Candida tropicalis

growing in sugarcane bagasse extract medium. Adv. Biol. Res., 4: 233–240.

107

Chen, B.Y., Hsueh, C. C, Chen, W. M. and Li, W. D., 2011. Exploring decolorization and

halotolerance characteristics by indigenous acclimatized bacteria: chemical structure of

azo dyes and dose–response assessment. J. Taiwan Inst. Chem. Eng. 42: 816–825.

Chen, B.Y., Zhang, M. M., Chang, C. T., Ding, Y., Chen, W. M., Hsueh, C. C., 2011. Deciphering

azo dye decolorization characteristics by indigenous Proteus hauseri: chemical structure.

J. Taiwan Inst. Chem. Eng., 42: 327–333.

Chen, K. C., Wu, J. Y., Liou, D. J. and Hwang, S. C. J., 2003. Decolorization of the textile dyes

by newly isolated bacterial strains. J. Biotechnol., 101: 57-68.

Chen, T., Gao, B. and Yue, Q., 2010. Effect of dosing method and pH on color removal

performance and floc aggregation of polyferric chloride–polyamine dual-coagulant in

synthetic dyeing wastewater treatment. Colloids Surf A: Physicochem. Eng. Asp., 355:

121-129.

Chougule, A. S., Jadhav, S. B. and Jadhav, J. P., 2014. Microbial degradation and detoxification

of synthetic dye mixture by Pseudomonas sp. SUK 1. Pro. Nation. Acad. Sci. Ind. Sec. B:

Bio. Sci., 84(4): 1059-1068.

Christie, R. M., 2007. Environmental aspects of textile dyeing. Elsevier.

Chung, K.T. and Stevens, S. E., 1993. Decolorization of azo dyes by environmental

microorganisms and helminths. Environ. Toxicol. Chem., 12: 2121-2132.

Dafale N., Wate, S. R., Meshram S. U. and Rao, N. N., 2010. Bioremediation of wastewater

containing azo dyes through sequential anaerobic-aerobic bioreactor system and its

biodiversity. Environ. Rev., 18: 21–36.

108

Dafale, N., Rao, N. N., Meshram, S. U. and Wate, S. R., 2008. Decolorization of azo dyes and

simulated dye bath wastewater using acclimatized microbial consortium: biostimulation

and halo tolerance. Bioresour. Technol., 99: 2552–2558.

Das, D., Charumathi, D. and Das, N., 2010. Combined effects of sugarcane bagasse extract and

synthetic dyes on the growth and bioaccumulation properties of Pichia fermentans MTCC

189. J. Hazard. Mater., 183: 497–505.

Das, D., Charumathi, D. and Das, N., 2011. Bioaccumulation of the synthetic dye Basic Violet 3

and heavy metals in single and binary systems by Candida tropicalis grown in a sugarcane

bagasse extract medium: modelling optimal conditions using response surface

methodology (RSM) and inhibition kinetics. J. Hazard. Mater., 186:1541–1552.

Dawkar, V. V., Jadhav, U. U., Jadhav, S. U. and Govindwar, S. P., 2008. Biodegradation of

disperse textile dye Brown 3REL by newly isolated Bacillus sp. VUS. J. Appl. Microbiol.,

105(1): 14-24.

Devi, P. S., Madhuri, T. and Indrani V., 2018. Analytical biodegradation of azo dye (Remazol Red

RB) by Bacillus cereus. J. Chem. Pharma. Res., 10(4): 74-80.

Dhanve, R. S., Shedbalkar, U. U. and Jadhav, J. P., 2008. Biodegradation of diazo Reactive dye

Navy blue HE2R (Reactive blue 172) by an isolated Exiguobacterium sp. RD3. Biotechnol.

Bioprocess. Eng., 13: 53–60.

dos Santos, A. B., Cervantes, F. J. and Van Lier, J. B., 2007. Review paper on current technologies

for decolourisation of textile wastewaters: perspectives for anaerobic biotechnology.

Bioresour. Technol., 98: 2369–2385.

Dua, M., Singh, A., Sethunathan, N. and Johri, A. K., 2002. Biotechnology and bioremediation:

successes and limitations. Appl. Microbiol. Biotechnol., 59: 143–152.

109

Dubey, S. K., Dubey, J., Viswas, A. J. and Tiwari, P., 2011. Studies on Cyanobacterial biodiversity

in paper mill and pharmaceutical industrial effluents. Br. Biotechnol. J. 1: 61–70.

Duran, N. and Esposito, E., 2000. Potential applications of oxidative enzymes and phenoloxidase-

like compounds in wastewater and soil treatment: a review. Appl. Catal. B: Environ., 28:

83–99.

El-Sheekh, M. M., Gharieb, M. M. and Abou-El-Souod, G. W., 2009. Biodegradation of dyes by

some green algae and cyanobacteria. Int. Biodeter. Biodegrad., 63: 699–704.

Enayatizamir, N., Tabandeh, F., Rodriˇıguez Couto, S., Yakhchali, B., Alikhani, H. A.,

Mohammadi, L., 2011. Biodegradation pathway and detoxification of the diazo dye

Reactive Black 5 by Phanerochaete chrysosporium. Bioresour. Technol., 102: 10359–

10362.

Erden, E., Kaymaz, Y. and Pazarlioglu, N. K., 2011. Biosorption kinetics of a direct azo dye Sirius

Blue K-CFN by Trametes versicolor. Electron J. Biotechnol., 14: 1–10.

Ergene, A., Ada, K., Tan, S. and Katırcıoglu, H., 2009. Removal of Remazol Brilliant Blue R dye

from aqueous solutions by adsorption onto immobilized Scenedesmus quadricauda:

equilibrium and kinetic modeling studies. Desalination, 249: 1308–1314.

Erkurt, A. E., Unyayar, A. and Kumbur, H., 2007. Decolorization of synthetic dyes by white rot

fungi, involving laccase enzyme in the process. Process Biochem., 42: 1429–1435.

Ertu˘grul, S., San, N. O. and Donmez, G., 2009. Treatment of dye (Remazol Blue) and heavy

metals using yeast cells with the purpose of managing polluted textile wastewaters. Ecol.

Eng., 35: 128–134.

Ertugrul, S., Bakır, M. and Donmez, G., 2008. Treatment of dye-rich wastewater by an

immobilized thermophilic cyanobacterial strain: Phormidium sp. Ecol. Eng., 32: 244–248.

110

Fan, F., Zhuo, R., Sun, S., Wan, X., Jiang, M., Zhang, X. and Yang, Y., 2011. Cloning and

functional analysis of a new laccase gene from Trametes sp. 48424 which had the high

yield of laccase and strong ability for decolorizing different dyes. Bioresour. Technol., 102:

3126–3137.

Fang, Z., Li, T., Wang, Q., Zhang, X., Peng, H, Fang, W., Hong, Y., Ge, H. and Xiao, Y., 2011. A

bacterial laccase from marine microbial metagenome exhibiting chloride tolerance and dye

decolorization ability. Appl. Microbiol. Biotechnol., 89: 1103–1110.

Forgacs, E., Cserhati, T. and Oros, G., 2004. Removal of synthetic dyes from wastewaters: a

review. Environ. Int. 30: 953–971.

Forss, J. and Welander, U., 2011. Biodegradation of azo and anthraquinone dyes in continuous

systems. Int. Biodeter. Biodegrad., 65: 227–237.

Fu, F. and Viraraghavan, T., 2001. Fungal decolorization of dye wastewaters: a review. Bioresour.

Technol., 79: 251–262.

Fu, Y. and Viraraghavan, T., 2000. Removal dye from an aqueous solution by fungus Aspergillus

niger. Water Qual. Res. J. Can., 35: 95–111.

Gabriel, H. and Hong, J., 2008. Decolouration of reactive blue dye KN-R. Res. J. Appl. Sci., 3:

216-224.

Ganesh, R., Boardman, G. D. and Michelson, D., 1994. Fate of azo dyes in sludges. Water Res.,

28: 1367–1376.

Gao, S., Tao, F., Wu, L., Huang, M. and Chen, C., 2010. Evaluation of the damage by dyestuffs

and NaCl on Phanerochaete chrysosporium. Environ. Prog. Sustainable Energy, 29: 428–

434.

111

Ghodake, G., Jadhav, U., Tamboli, D., Kagalkar, A. and Govindwar, S., 2011. Decolorization of

textile dyes and degradation of mono-azo dye Amaranth by Acinetobacter

calcoaceticus NCIM 2890. Ind. J. Microbiol., 51: 501–508.

Gill, M. and Strauch, R. J., 1984.Constituents of Agaricus xanthodermus Genevier: the first

naturally endogenous azo compound and toxic phenolic metabolites. Zeitschrift fur

Naturforschung C: J. Biosci., 39c: 1027 -1029.

Gomaa, E. Z., 2016. Biodegradation and detoxification of azo dyes by some bacterial strains. J.

Microbiol., 6(1-2): 15-24.

Gonen, F. and Aksu, Z.,2009. Predictive expressions of growth and Remazol Turquoise Blue-G

reactive dye bioaccumulation properties of Candida utilis. Enzyme Microbiol. Technol.,

45: 15-21.

Gonzalez-Gutierrez, L. V. and Escamilla-Silva, E.M., 2009. Reactive red azo dye degradation in

a UASB bioreactor: mechanism and kinetics. Eng. Life Sci., 9: 311–316.

Guadie, A., Tizazua, S., Meleseb, M., Guoc, W., Ngo, H. H., Xiad, S., 2017. Biodecolorization of

textile azo dye using Bacillus sp. strain CH12 isolated from alkaline lake. Biotechnol.

Reports, 15: 92-100.

Gupta, V. K., 2009. Application of low-cost adsorbents for dye removal-A review. J Environ.

Manage., 90: 2313-2342.

Halaburgi, V., Sharma, S., Sinha, M., Singh, T, P. and Karegoudar, T. B., 2011. Purification and

characterization of a thermostable laccase from the ascomycetes Cladosporium

cladosporioides and its applications. Process Biochem., 46: 1146–1152.

Hao, O. J., Kim, H. C and Hiang, P. C., 2000. Decolorization of wastewater. Critical reviews.

Environ. Sci. Technol., 30: 449 -505.

https://www.ncbi.nlm.nih.gov/pubmed/?term=Ghodake%20G%5BAuthor%5D&cauthor=true&cauthor_uid=23024414

https://www.ncbi.nlm.nih.gov/pubmed/?term=Jadhav%20U%5BAuthor%5D&cauthor=true&cauthor_uid=23024414

https://www.ncbi.nlm.nih.gov/pubmed/?term=Tamboli%20D%5BAuthor%5D&cauthor=true&cauthor_uid=23024414

https://www.ncbi.nlm.nih.gov/pubmed/?term=Kagalkar%20A%5BAuthor%5D&cauthor=true&cauthor_uid=23024414

https://www.ncbi.nlm.nih.gov/pubmed/?term=Govindwar%20S%5BAuthor%5D&cauthor=true&cauthor_uid=23024414

https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3209937/

112

Hao, O. J., Kim, H. C. and Hiang, P. C., 2000. Decolorization of wastewater. Critical reviews.

Environ. Sci. Technol., 30: 449 -505.

Harwood, C. S., Parales, R. E., 1996. The beta-ketoadipate pathway and the biology of self

identity. Annu. Rev. Microbiol., 50: 553-590.

Houk, V. S., 1992. The genotoxicity of industrial wastes and effluents: A review. Mutat. Res., 277:

91–138.

Hsueh, C. C., Chen, B. Y. and Yen, C. Y., 2009. Understanding effects of chemical structure on

azo dye decolorization characteristics by Aeromonas hydrophila. J. Hazard. Mater., 167:

995–1001.

Hunger, K., (Ed.), 2003. Industrial dyes: chemistry, properties, applications. John Wiley & Sons.

Hungerer, C., Troup, B., Römling, U. and Jahn, U. D., 1996. Regulation of the hemA gene during

5-aminolevulinic acid formation in Pseudomonas aeruginosa. J. Bacteriol. 177: 1435-43.

Husain, M. and Husain, Q., 2007. Applications of redox mediators in the treatment of organic

pollutants by using oxidoreductive enzymes: a review. Crit. Rev. Env. Sci. Technol., 38:

1–42.

Husain, Q. and Ulber, R., 2011. Immobilized peroxidase as a valuable tool in the remediation of

aromatic pollutants and xenobiotic compounds: a review. Crit. Rev. Env. Sci. Technol., 41:

770-804.

Husain, Q., 2010. Peroxidase mediated decolorization and remediation of wastewater containing

industrial dyes: a review. Rev. Environ. Sci. Biotechnol., 9: 117–140.

Ilyas, S. and Rehman, A., 2013. Decolorization and detoxification of Synozol red HF-6BN azo

dye, by Aspergillus niger and Nigrospora sp. Iranian J. Environ. Health Sci. Eng., 10: 12.

https://www.ncbi.nlm.nih.gov/pubmed/?term=Ilyas%20S%5BAuthor%5D&cauthor=true&cauthor_uid=23369298

https://www.ncbi.nlm.nih.gov/pubmed/?term=Rehman%20A%5BAuthor%5D&cauthor=true&cauthor_uid=23369298

https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3605312/

113

Iqbal, M. and Saeed, A., 2007. Biosorption of reactive dye by loofa sponge-immobilized fungal

biomass of Phanerochaete chrysosporium. Process Biochem., 42: 1160–1164.

Jadhav, J. P., Kalyani, D. C., Telke, A. A., Phugare, S. S. and Govindwar, S. P., 2010. Evaluation

of the efficacy of a bacterial consortium for the removal of color, reduction of heavy metals

and toxicity from textile dye effluent. Bioresour. Technol., 101: 165–173.

Jadhav, J. P., Parshetti, G. K., Kalme, S. D. and Govindwar, S. P., 2007. Decolourization of azo

dye Methyl Red by Saccharomyces cerevisiae MTCC 463. Chemosphere, 68: 394–400.

Jadhav, J. P., Phugare, S. S., Dhanve, R. S. and Jadhav, S. B., 2010. Rapid biodegradation and

decolorization of Direct Orange 39 (Orange TGLL) by an isolated bacterium strain BCH.

Biodegrad., 21: 453–463.

Jadhav, S. B., Phugare, S. S., Patil, P. S. and Jadhav, J. P., 2011. Biochemical degradation pathway

of textile dye Remazol Red and subsequent toxicological evaluation by cytotoxicity,

genotoxicity and oxidative stress studies. Int. Biodeter. Biodegrad., 65: 733–743.

Jimenez-Juarez, N., Roman-Miranda, R., Baeza, A., Sanchez-Amat, A., Vazquez-Duhalt, R. and

Valderrama, B., 2005. Alkali and halide-resistant catalysis by the multipotent oxidase from

Marinomonas mediterranea. J. Biotechnol., 117: 73–82.

Jin, X. C., Liu, G. Q., Xu, Z. H., Tao, W. Y., 2007. Decolorization of a dye industry effluent by

Aspergillus fumigatus XC6. Appl. Microbiol. Biotechnol., 74: 239-243.

Joe, M., Lim, S. Y., Kim, D. H. and Lee, I. S., 2008. Decolorization of textile dyes by Clostridium

bifermentans SL186 isolated from contaminated soil. World J. Microbiol. Biotechnol., 24:

2221–2226.

114

Joshi, S. M., Inamdar, S. A., Telke, A. A., Tamboli, D. P. and Govindwar, S. P., 2010. Exploring

the potential of natural bacterial consortium to degrade mixture of dyes and textile effluent.

Int. Biodeter. Biodegrad., 64: 622–628.

Kalyani, D. C., Telke, A. A., Dhanve, R. S. and Jadhav, J. P., 2008. Ecofriendly biodecolorization

and detoxification of reactive red 2 textile dye by newly isolated Pseudomonas sp. SUK1.

J. Hazard. Mater., 163: 735.

Kalyani, D., Dhiman, S. S., Kim, H., Jeya, M., Kim, I. W. and Lee, J. K., 2012. Characterization

of a novel laccase from the isolated Coltricia perennis and its application to detoxification

of biomass. Process Biochem., 47(4): 671-678.

Karimi, A., Vahabzadeh, F. and Bonakdarpour, B., 2006. Use of Phanerochaete chrysosporium

immobilized on kissiris for synthetic dye decolourization: involvement of manganese

peroxidase. World J. Microbiol. Biotech., 22: 1251–1257.

Kaushik, P. and Malik, A., 2009. Fungal dye decolourization: recent advances and future potential

review article. Environ. Int., 35: 127–141.

Keck, A., Klein, J., Kudlich, M., Stolz, A., Knackmuss, H. J., Mattes, R., 1997. Reduction of azo

dyes by redox mediators originating in the naphthalenesulfonic acid degradation pathway

of Sphingomonas sp. strain BN6. Appl. Enviro. Microbiol., 63(9): 3684–3690.

Khan, Z., Nisar, M. A., Hussain, Z. H., Arshad, M. N. and Rehman, A., 2015. Cadmium resistance

mechanism in Escherichia coli P4 and its potential use to bioremediate environmental

cadmium. Appl. Microbiol. Biotechnol., 99(24): 10745-10757.

Khelifi, E., Ayed, L., Bouallagui, H., Touhami, Y. and Hamdi, M., 2009. Effect of nitrogen and

carbon sources on Indigo and Congo red decolourization by Aspergillus alliaceus strain

121C. J. Hazard. Mater., 163: 1056–1062.

115

Khlifi, R., Belbahri, L., Woodward, S., Ellouz, M., Dhouib, A., Sayadi, S. and Mechichi, T., 2010.

Decolorization and detoxification of textile industry wastewater by the laccase-mediator

system. J. Hazard. Mater., 175: 802–808.

Kirk-Othmer, 1979. Encyclopedia of Chemical Technology. 3rd Ed, pp. 1-172, New York: Wiley-

Inter Science.

Kodam, K. M., Soojhawon, I., Lohande, P.D. and Gawai, K.R., 2005. Microbial decolorization of

reactive azo dyes under aerobic conditions. World J. Microbiol. Biotechnol., 21: 367–370.

Kumari, L., Verma, A. K., Tiwary, D., Giri, D. D., Nath, G. and Mishra, P. K., 2015.

Biodegradation of Navy N5RL1 carpet dye by Staphylococcus saprophyticus strain

BHUSS X3. Biotech., 5: 775–782.

Kurade, M. B., Waghmode, T. R. and Govindwar, S. P., 2011. Preferential biodegradation of

structurally dissimilar dyes from a mixture by Brevibacillus laterosporus. J. Hazard.

Mater., 192: 1746–1755.

Lade, H. S., Kadam, A. A., Paul, D. and Govindwar, S., 2015. Biodegradation and detoxification

of textile azo dyes by bacterial consortium under sequential microaerophilic/aerobic

processes. EXCLI J., 14: 158-174.

Lade, H., Govindwar, S. and Paul, D., 2015. Low-cost biodegradation and detoxification of textile

azo dye C.I. Reactive blue 172 by Providencia rettgeri strain HSL1. J. Chem., 2015: 10.

Latif, A., Noor, S., Sharif, Q. M. and Najeebullah, M., 2010. Different techniques recently used

for the treatment of textile dyeing effluents: a review. J. Chem. Soc. Pak., 32: 115–124.

Levin, L., Melignani, E. and Ramos, A. M., 2010. Effect of nitrogen sources and vitamins on

ligninolytic enzyme production by some white-rot fungi. Dye decolorization by selected

culture filtrates. Bioresour. Technol., 101: 4554–4563.

116

Liers, C., Bobeth, C., Pecyna, M., Ullrich, R. and Hofrichter, M., 2010. DyP-like peroxidases of

the jelly fungus Auricularia auricula-judae oxidize nonphenolic lignin model compounds

and high-redox potential dyes. Appl. Microbiol Biotechnol., 85:1869–1879.

Lim, S. L., Chu, W. L. and Phang, S. M., 2010. Use of Chlorella vulgaris for bioremediation of

textile wastewater. Bioresour. Technol., 101: 7314–7322.

Lin, S. H. and Peng, C. F., 1994. Treatment of textile wastewater by electrochemical method.

Water Res., 28: 277-282.

Lin, S. H. and Peng, C. F., 1996. Continuous treatment of textile wastewater by combined

coagulation, electrochemical oxidation and activated sludge. Water Res., 30: 587 -592.

Lin, Y. H. and Leu, J.Y., 2008. Kinetics of reactive azo-dye decolorization by Pseudomonas

luteola in a biological activated carbon process. Biochem. Eng. J., 39: 457–467.

Liu, G., Zhou, J., Lv, H., Xiang, X., Wang, J. and Qv, Y., 2007. Azoreductase from Rhodobacter

sphaeroides AS1.1737 is a flavodoxin that also functions as nitroreductase and flavin

mononucleotide reductase. Appl. Microbiol. Biotechnol., 76: 1271–1279.

Liu, W., You, Y., Sun, D. and Wang, S., 2018. Decolorization and detoxification of water insoluble

Sudan dye by Shewanella putrefaciens CN32 co-cultured with Bacillus circulans

BWL1061. Ecotoxicol. Environ. Safety, 166: 11-17.

Liu, X., Zhang, J., Jiang, J., Li, R., Xie, Z. and Li, S., 2011. Biochemical degradation pathway of

reactive blue 13 by Candida rugopelliculosa HXL-2. Int. Biodeter. Biodegrad., 65: 135–

141.

Lucas, M. S., Amaral, C., Sampaio, A., Peres, J. A. and Dias, A. A., 2006. Biodegradation of the

diazo dye Reactive Black 5 by a wild isolate of Candida oleophila. Enzyme Microbiol.

Technol., 39: 51–55.

117

Machado, K. M. G., Compart, L. C. A., Morais, R. O., Rosa, L. H. and Santos, M. H., 2006.

Biodegradation of reactive textile dyes by basidiomycetous fungi from Brazilian

ecosystems. Braz. J. Microbiol., 37: 481–487.

Majeau, J. A., Brar, S. K. and Tyagi, R. D., 2010. Laccases for removal of recalcitrant and

emerging pollutants. Bioresour. Technol., 101: 2331–2350.

Mali, P. L., Mahajan, M. M., Patil, D. P. and Kulkarni, M. V., 2000. Bio-decolorization of

members of triphenyl methane and azo group of dyes. J. Sci. Ind. Res., 59: 221-224.

Malik, S. N., Ghosh, P. C., Vaidya, A. N., Waindeskar, V., Das, S. and Mudliar, S. N., 2017.

Comparison of coagulation, ozone and ferrate treatment processes for color, COD and

toxicity removal from complex textile wastewater. Water Sci. Technol., 76: 1001-1010.

Martorell, M. M., Pajot, H. F. and de Figueroa, L. I. C., 2012. Dye-decolourizing yeasts isolated

from Las Yungas rainforest Dye assimilation and removal used as selection criteria. Int.

Biodeter. Biodegrad., 66: 25–32.

Masneuf-Pomarade, I., Jeuneb, C. L., Durrensc, P., Lollierb, M., Aigled, M. and Dubourdieu, D.,

2007. Molecular typing of wine yeast strains Saccharomyces bayanus var. uvarum using

microsatellite markers. Sys. Appl. Microbiol., 30: 75-82.

Matsumoto, K., Mukai, Y., Ogata, D., Shozui, F., Nduko, J. M. and Taguchi, S. and Ooi, T., 2010.

Characterization of thermostable FMN-dependent NADH azoreductase from the moderate

thermophile Geobacillus stearothermophilus. Appl. Microbiol. Biotechnol., 86: 1431–

1438.

McMullan, G., Meehan, C., Conneely, A., Kirby, N., Robinson, T. and Nigam, P., Banat, I.,

Merchant, R. and Smyth, W., 2001. Microbial decolourisation and degradation of textile

dyes. Appl. Microbiol. Biotechnol., 56: 81–87.

118

Meerbergen, K., Willims, K. A., Dewil, F., Impe, J. V., Appels, L. and Lievens, B., 2018. Isolation

and screening of bacterial isolates from wastewater treatment plants to decolorize azo dyes.

J. Biosci. Bioengineer., 125(4): 448-456.

Megharaj, M., Ramakrishnan, B., Venkateswarlu, K., Sethunathan, N. and Naidu, R., 2011.

Bioremediation approaches for organic pollutants: a critical perspective. Environ. Int., 37:

1362–1375.

Mendes, S., Pereira, L., Batista, C. and Martins, L. O., 2011. Molecular determinants of

azoreduction activity in the strain Pseudomonas putida MET94. Appl. Microbiol.

Biotechnol., 92: 393–405.

Meng, X., Liu, G., Zhou, J., Fu, Q. S. and Wanga, G., 2012. Azo dye decolorization by Shewanella

aquimarina under saline conditions. Bioresour, Technol., 114: 95–101.

Mester, T. and Tien, M., 2000. Oxidation mechanism of ligninolytic enzymes involved in the

degradation of environmental pollutants. Int. Biodeter. Biodegrad., 46: 51-59.

Misal, S. A., Lingojwar, D. P., Shinde, R. M. and Gawai, K. R., 2011. Purification and

characterization of azoreductase from alkaliphilic strain Bacillus badius. Process

Biochem., 46: 1264–1269.

Mohan, S. V., Ramanaiah, S.V. and Sarma, P. N., 2008. Biosorption of direct azo dye from

aqueous phase onto Spirogyra sp I02: evaluation of kinetics and mechanistic aspects.

Biochem. Eng. J. 38: 61–69.

Mohana, S.V., Shrivastav, S., Divecha, J. and Madamwar, D., 2008. Response surface

methodology for optimization of medium for decolorization of textile dye Direct Black 22

by a novel bacterial consortium. Bioresour. Technol., 99: 562–569.

119

Moosvi, S., Keharia, H. and Madamwar, D., 2005. Decolorization of textile dye Reactive Violet 5

by a newly isolated bacterial consortium RVM 11.1. World J. Microbiol. Biotechnol., 21:

667–672.

Moosvi, S., Kher, X. and Madamwar, D., 2007. Isolation, characterization and decolorization of

textile dyes by a mixed bacterial consortium JW-2. Dyes Pigments, 74: 723–729.

Nachiyar, C. V. and Rajakumar, G. S., 2005. Purification and characterization of an oxygen

insensitive azoreductase from Pseudomonas aeruginosa. Enzyme Microbiol. Technol., 36:

503 -509.

Nair, L. K., Begum, M. and Ragunathan, R., 2017. Biodegradation of azo dyes using Bacillus

megaterium and its phytotoxicity study. IOSR J. Environ, Sci, Toxicol, Food Technol.,

11(7): 12-20.

Noreen, U., Ahmed, Z., Khalid, A., Diserafino, A., Ume-Habiba, Ali, F. and Hussain, F., 2019.

Water pollution and occupational health hazards caused by the marble industries in district

Mardan, Pakistan. Environ. Technol. Innov., 16: 100470.

O’Neill, C., Hawkes, F. R., Hawkes, D. L., Lourenco, N. D., Pinheiro, H. M. and Delee, W., 1999.

Color in textile effluents sources, measurement, discharge consents and simulation: a

review. J. Chem. Technol. Biotechnol., 53: 249–254.

Ogola, H. J. O., Hashimoto, N., Miyabe, S., Ishikawa, T., Shibata, H. and Sawa, Y., 2010.

Enhancement of hydrogen peroxide stability of a novel Anabaena sp. DyP-type peroxidase

by site directed mutagenesis of methionine residues. Appl. Microbiol. Biotechnol., 87:

1727–1736.

Oh, Y. K., Kim, Y. J., Ahn, Y., Song, S. K. and Park, S., 2004. Color removal of real textile

wastewater by sequential anaerobic aerobic reactors. Biotechnol. Bioprocess. Eng., 9: 419.

120

Ola, I. O., Akintokun, A. K., Akpan, I., Omomowo, I. O. and Areo, V. O., 2010. Aerobic

decolourization of two reactive azo dyes under varying carbon and nitrogen source by

Bacillus cereus. Afr. J. Biotechnol., 9: 672–677.

Ollgaard, H., Frost, L., Galster, J. and Hensen, O. C., 1999. Survey of azo-colorants on Denmark:

Milgoproject 509, Danish Environmental Protection Agency.

Olukanni, O. D., Osuntoki, A. A. and Gbenle, G. O., 2009. Decolorization of azo dyed by a starin

of Micrococcus isolated from refuse dump soil. Biotechnol., 8(4): 442-448.

Olukanni, O. D., Osuntoki, A. A. and Gbenle, G. O., 2009. Decolourizatioin of azo dyes by a strain

of Micrococcus isolated from a refuse dump soil. Biotechnol., 8: 442–448.

O'Neil, C., Hawkes, F. R., Hawkes, D. L., Lourenco, N. D., Pinheiro, H. M. and Delee, W., 1999.

Color in textile effluents sources, measurements, discharge contents and simulation: a

review. J. Chem. Technol. Biotechnol., 74: 1009-1018.

Ooi, T., Shibata, T., Sato, R., Ohno, H., Kinoshita, S., Thuoc, T. L. and Taguchi, S., 2007. An

azoreductase, aerobic NADH-dependent flavoprotein discovered from Bacillus sp.:

functional expression and enzymatic characterization. Appl. Microbiol. Biotechnol., 75:

377–386.

Oturkar, C. C., Nemade, H. N., Mulik, P. M., Patole, M. S., Hawaldar, R. R. and Gawai, K. R.,

2011. Mechanistic investigation of decolorization and degradation of Reactive Red 120 by

Bacillus lentus BI377. Bioresour. Technol., 102: 758–764.

Öztürk, A. and Abdullah, M. I., 2006. Toxicological effect of indole and its azo dye derivatives on

some microorganisms under aerobic conditions. Sci. Total Environ., 358: 137–142

121

Pajot, H. F., de Figueroa, L. I. C., Farina, J. I., 2007. Dye-decolorizing activity in isolated yeasts

from the ecoregion of Las Yungas (Tucuman Argentina). Enzyme Microbiol. Technol., 40:

1503-1511.

Pandey, A., Singh, P. and Iyengar, L., 2007. Bacterial decolorization and degradation of azo dyes.

Int. Biodeter. Biodegrad., 59: 73-84.

Pandey, A., Singh, P. and Lyengar, L., 2007. Review Bacterial decolorization and degradation of

azo dyes. Int. Biodeter. Biodegrad., 59: 73–84.

Parmar, N. D. and Shukla, S. R., 2018. Biodegradation of anthraquinone based dye using an

isolated strain Staphylococcus hominis subsp. hominis DSM 20328. Environ. Progress

Sustainable Energy, 37(1): 203-214.

Parshetti, G., Kalme, S., Saratale, G., and Govindwar, S., 2006. Biodecolorization of malachite

green by Kocuria rosea MTCC 1532. Acta. Chim. Slov., 53: 492-498.

Pasti-Grigsby, M. B., Paszczynski, A., Goszczynski, S., Crawford, D. L. and Crawford, R. L.,

1992. Influence of aromatic substitution patterns on azo dye degradability by Streptomyces

sp. and Phanerochaete chrysosporium. Appl. Environ. Microbiol., 58: 3605–3613.

Pazarlioglu, N. K., Urek, R. O. and Ergun, F., 2005. Biodecolourization of Direct Blue 15 by

immobilized Phanerochaete chrysosporium. Process Biochem., 40: 1923–1929.

Pearce, C. I., Lloyd, J. R. and Guthrie, J. T., 2003. The removal of colour from textile wastewater

using whole bacterial cells: a review. Dyes Pigments, 58: 179–196.

Perumal, S. M., Munuswamy, D., Sellamuthu, P. S., Kandasamy, M. and Thangavelu, K. P., 2007.

Biosorption of textile dyes and effluents by Pleurotus florida and Trametes hirsuta with

evaluation of their laccase activity. Iran J Biotechnol. 5:114–118.

122

Phugare, S. S., Kalyani, D. C., Patil, A. V. and Jadhav, J. P., 2011. Textile dye degradation by

bacterial consortium and subsequent toxicological analysis of dye and dye metabolites

using cytotoxicity, genotoxicity and oxidative stress studies. J. Hazard. Mater., 186: 713–

723.

Phugare, S., Patil, P., Govindwar, S. and Jadhav, J., 2010. Exploitation of yeast biomass generated

as a waste product of distillery industry for remediation of textile industry effluent. Int.

Biodeter.Biodegrad., 64: 716–726.

Prasad, A. S. A. and Rao, K. V. B., 2014. Aerobic biodegradation of azo dye Acid Black-24 by

Bacillus halodurans. J. Environ. Bio., 35: 549-554.

Prasad, A. S. A., Satyanarayana, V. S. V. and Rao, K.V. B., 2013. Biotransformation of Direct

blue 1 by a moderately halophilic bacterium Marinobacter sp. strain HBRA and toxicity

assessment of degraded metabolites. J. Hazard. Mater., 262: 674-684.

Pricelius, S., Held, C., Sollner, S., Deller, S., Murkovic, M., Ullrich, R., et al 2007. Enzymatic

reduction and oxidation of fibre-bound azo-dyes. Enzyme Microbiol. Technol., 40: 1732–

1738.

Puvaneswari, N., Muthukrishnan, J. and Gunasekaran, P., 2006. Toxicity assessment and microbial

degradation of azo dyes. Ind. J. Exp. Biol., 44: 618-26.

Rafii, F., Franklin, W. and Cerniglia, C. E., 1990. Azoreductase activity of anaerobic bacteria

isolated from human intestinal microflora. Appl. Environ. Microbiol., 56: 2146-2151.

Rai, H. S., Bhattacharyya, M. S., Singh, J., Bansal, T. K., Vats, P. and Banerjee, U. C., 2005.

Removal of dyes from the effluent of textile and dyestuff manufacturing industry: a review

of emerging techniques with reference to biological treatment. Crit. Rev. Environ. Sci.

Technol., 35: 219-238.

https://www.ncbi.nlm.nih.gov/pubmed/16924831

123

Rai, H. S., Bhattacharyya, M. S., Singh, J., Bansal, T. K., Vats, P. and Banerjee, U. C., 2005.

Removal of dyes from the effluent of textile and dyestuff manufacturing industry: a review

of emerging techniques with reference to biological treatment. Crit. Rev. Environ. Sci.

Technol., 35: 219-238.

Rajaguru, P., Vidya, L., Baskarasethupathi, B., Kumar, P. A., Palanivel, M. and Kalaiselvi, K.,

2002. Genotoxicity evaluation of polluted ground water in human peripheral blood

lymphocytes using the comet assay. Mutat. Res.Genet. Toxicol. Environ. Mutagen. 517:

29-37

Rajaguru, P., Vidya, L., Baskarasethupathi, B., Kumar, P. A., Palanivel, M. and Kalaiselvi, K.,

2002. Genotoxicity evaluation of polluted ground water in human peripheral blood

lymphocytes using the comet assay. Mutat. Res., 517: 29–37.

Rajee, O. and Patterson, J., 2011. Decolorization of azo dye (Orange MR) by an autochthonous

bacterium Micrococcus sp. DBS 2. Ind. J. Microbiol., 51: 159–163.

Ramalho, P. A., Scholze, H., Cardoso, M. H. and Ramalho, M. T. and Oliveira-Campos, A. M.,

2002. Improved conditions for the aerobic reductive decolourisation of azo dyes by

Candida zeylanoides. Enzyme Microbiol. Technol., 31: 848–854.

Ramanath, R., 2005. Color: An Introduction to Practice and Principles. Wiley Online Library,

USA.

Renganathan, S., Thilagaraj, W. R., Miranda, L. R., Gautam, P. and Velan, M. 2006. Accumulation

of Acid Orange 7 Acid Red 18 and Reactive Black 5 by growing Schizophyllum commune.


124

Robinson, T., McMullan, G., Marchant, R. and Nigam, P., 2001. Remediation of dyes in textile

effluent: a critical review on current treatment technologies with a proposed alternative.


Ruiz-Arias, A., Juarez-Ramirez, C., de los Cobos-Vasconcelos, D., Ruiz-Ordaz, N., Salmeron-

Alcocer, A., Ahuatzi, C. D. and Galindz-Mayer, J., 2010. Aerobic biodegradation of a

sulfonated phenylazonaphthol dye by a bacterial community immobilized in a multistage

packed-bed BAC reactor. Appl. Biochem. Biotechnol., 162: 1689–1707.

Saha, A. K., Sultana, N., Mohanta, M. K., Mandald, A. and Haquee, M., 2017. Identification and

characterization of azo dye decolourizing bacterial strains, Alcaligenes faecalis E5.Cd and

A. faecalis Fal.3 isolated from textile effluents. Am. Sci. Res. J. Eng. Technol. Sci., 3: 163-

175.

Saharan, B. S. and Ranga, P., 2011. Optimization of cultural conditions for decolourization of

textile azo dyes by Bacillus subtilis spr42 under submerged fermentation, Int. J. Adv.

Biotechnol. Res., 2: 148-153.

Sahasrabudhe, M. M., Saratale, R. G., Saratale, G. D. and Pathade, G. R., 2014. Decolorization

and detoxification of sulfonated toxic diazo dye C.I. Direct red 81 by Enterococcus

faecalis YZ 66. J. Environ. Health Sci. Eng., 12: 151.

Sani, K. R. and Banerjee, U. C., 1999. Decolorization of triphenylmethane dyes and textile and

dyestuff effluent by Kurthia sp. Enzyme Microbiol. Technol. 24: 433–437.

Santhanam, N., Vivanco, J. M., Decker, S. R. and Reardon, K. F., 2011. Expression of industrially

relevant laccases: prokaryotic style. Trends Biotechnol., 29: 480–489.

125

Saratale, G. D., Saratale, R. G., Chang, J. S. and Govindwar, S. P., 2011. Fixed-bed decolorization

of Reactive Blue 172 NCIM-2027 immobilized on Luffa cylindrica sponge. Int. Biodeter.

Biodegrad., 65: 494–503.

Saratale, R. G., Saratale, G. D., Chang, J. S. and Govindwar, S. P., 2009a. Ecofriendly degradation

of sulfonated diazo dye C.I Reactive Green 19A using Micrococcus glutamicus NCIM-

2168. Bioresour. Technol., 100: 3897–3905.

Saratale, R. G., Saratale, G. D., Chang, J. S. and Govindwar, S. P., 2009b. Decolorization and

biodecolorization of textile dye Navy Blue HER by Trichosporon beigelii NCIM-3326. J.

Hazard. Mater., 166: 1421.

Sarkar, S., Banerjee, A., Halder, U., Biswas, R. and Bandopadhyay, R., 2017. Degradation of

synthetic azo dyes of textile industry: a sustainable approach using microbial enzymes.

Water Conserv. Sci. Eng., 2: 121–131.

Sathiya, M., Periyar, S., Sasikalaveni, A., Murugesan, K. and Kalaichelvan, P., T., 2007.

Decolorization of textile dyes and their effluents using white rot fungi. Afr. J. Biotechnol.,

6: 424-429.

Sawhney, R. and Kumar, A., 2011. Congo Red (azo dye) decolorization by local isolate VTII

inhabiting dye effluent exposed soil. Int. J. Environ. Sci., 1: 1261–1267.

Shah, M. P., 2014. Biodegradation of azo dyes by three isolated bacterial strains: an environmental

bioremedial approach. J. Microbial. Biochem. Technol., S3, 007.

Shah, M. P., Patel, K. A., Nair, S. S and Darji, A. M., 2103. Potential effect of two Bacillus sp.

on decolorization of azo dye. J. Bioremed. Biodegrad., 4: 199.

126

Shanmugam, B. K., Easwaran, S. N, Lakra, R., Deepa, P. R. and Mahadevan, S., 2017. Metabolic

pathway and role of individual species in the bacterial consortium for biodegradation of

azo dye: A biocalorimetric investigation. Chemosphere, 188: 81-89.

Shyamala, A., Hemapriya, J., Vadakkan. K. and Vijayanand, S., 2014. Bioremediation of Methyl

orange, a synthetic textile azo dye by a halotolerant bacterial strain. Int. J. Curr. Res. Acad.

Rev., 2: 373-381.

Singh, S., Chatterji, S., Nandini, P. T., Prasad, A. S. A. and Rao, K. V. B., 2015. Biodegradation

of azo dye Direct Orange 16 by Micrococcus luteus strain SSN2. Int. J. Environ. Sci.

Technol., 12: 2161-2168.

Sivarajasekar, N., Baskar, R. and Balakrishnan, V., 2009. Biosorption of an azo dye from aqueous

solutions onto Spirogyra. J. Univ. Chem. Technol. Metall., 44: 157–164.

Soares, G. M. B., Costa-Ferreira, M. and de Amorim, M. T. P., 2001. Decolorization of an

anthraquinone-type dye using a laccase formulation. Bioresour. Technol., 79: 171–177.

Solis, M., Solis, A., Perez, H. I., Manjarrez, N. and Flores, M., 2012. Microbial decolouration of

azo dyes: a review. Process Biochem., 47: 1723-1748.

Solis-Oba, M., Barzana, E., Garcia-Garibay, M. and Viniegra-Gonzalez, G., 2007. The ABTS: an

oxidant agent of different chemical compounds and its recycling process between laccase

and substrate. Rev. Mex. Ing. Quim., 6: 275–281.

Spadarry, J. T., Isabelle, L. and Renganathan, V., 1994. Hydroxyl radical mediated degradation of

azo dyes: evidence for benzene generation. Environ. Sci. Technol., 28: 1389-1393

Srinivasan, A. and Viraraghavan, T., 2010. Decolorization of dye wastewaters by biosorbents: a

review. J. Environ. Manage., 91: 1915–1929.

https://www.ncbi.nlm.nih.gov/pubmed/?term=Shanmugam%20BK%5BAuthor%5D&cauthor=true&cauthor_uid=28869849

https://www.ncbi.nlm.nih.gov/pubmed/?term=Easwaran%20SN%5BAuthor%5D&cauthor=true&cauthor_uid=28869849

https://www.ncbi.nlm.nih.gov/pubmed/?term=Lakra%20R%5BAuthor%5D&cauthor=true&cauthor_uid=28869849

https://www.ncbi.nlm.nih.gov/pubmed/?term=Deepa%20PR%5BAuthor%5D&cauthor=true&cauthor_uid=28869849

https://www.ncbi.nlm.nih.gov/pubmed/?term=Mahadevan%20S%5BAuthor%5D&cauthor=true&cauthor_uid=28869849

127

Stolz, A., 2001. Basic and applied aspects in the microbial degradation of azo dyes. Appl.

Microbiol. Biotechnol., 56: 69 –80.

Sugumar, S. and Thangam, B., 2012. BiodEnz: a database of biodegrading enzymes.

Bioinformation, 8: 40–42.

Sumathi, S. and Phatak, V., 1999. Fungal treatment of bagasse based pulp and paper mill wastes.

Environ. Technol., 19: 93–98.

Suteu, D., Zaharia, C. and Malutan, T., 2011. Biosorbents Based On Lignin Used. In Biosorption

Processes From Wastewater Treatment (chapter 7). In: Lignin: Properties and Applications

in Biotechnology and Bioenergy, Ryan J. Paterson (Ed.), pp. 27, Nova Science Publishers,

ISBN 978-1-61122-907-3, New York, U.S.A.

Svobodova, K., Senholdt, M., Novotny, C. and Rehorek, A., 2007. Mechanism of Reactive Orange

16 degradation with the white rot fungus Irpex lacteus. Process Biochem., 42: 1279–1284.

Swaminathan, K., Sandhya, S., Sophia, A. C., Pachhade, K., Subrahmanyam, Y. V., 2003.

Decolorization and degradation of H-acid and other dyes using ferrous-hydrogen peroxide

system. Chemosphere, 50: 619-625.

Tauber, M. M., Gubitz, G. M. and Rehorek, A., 2008. Degradation of azo dyes by oxidative

processes: Laccase and ultrasound treatment. Bioresour. Technol., 99: 4213–4220.

Telke, A. A., Ghodake, G. S., Kalyani, D. C., Dhanve, R. S and Govindwar, S. P., 2011.

Biochemical characteristics of a textile dye degrading extracellular laccase from a Bacillus

sp. ADR. Bioresour. Technol., 102: 1752–1756.

Telke, A. A., Kalyani, D., Dawkar, V. and Govindwar, S., 2009. Influence of organic and inorganic

compounds on oxidoreductive decolourization of sulfonated azo dye C.I.Reactive Orange

16. J. Hazard. Mater., 172: 298-309.

128

textile dyes Reactive Red 2 and Reactive Blue 4 using Bjerkandera sp Strain BOL 13 in a

continuous rotating biological contactor reactor. Enzyme Microbiol. Technol., 9: 32–37.

Tilli, S., Ciullini, I., Scozzafava, A. and Briganti, F., 2011. Differential decolorization of textile

dyes in mixtures and the joint effect of laccase and cellobiose dehydrogenase activities

present in extracellular extracts from Funalia trogii. Enzyme Microbiol, Technol., 49: 465–

471.

Tony, B. D., Goyal, D. and Khanna, S., 2009. Decolorization of textile azo dyes by aerobic

bacterial consortium. Int. Biodeter. Biodegrad., 63: 462–469.

Uday, U. S., Choudhury, P., Bandyopadhyay, T. K. and Bhunia, B., 2016. Classification, mode of

action and production strategy of xylanase and its application for biofuel production from

water hyacinth. Int. J. Bio. Macromolecules, 82: 1041–1054.

Umbuzeiro, G. A., Freeman, H., Warren, S. H., Oliveira, D, P., Terao, Y., Watanabe, T., Claxton,

L. D., 2005. The contribution of azo dyes to the mutagenic activity of the Cristais River.

Chemosphere., 60: 55–64.

Umbuzeiro, G. A., Freeman, H., Warren, S. H., Oliveira, D. P., Terao, Y., Watanabe, T. and

Claxton, L. D., 2005. The contribution of azo dyes to the mutagenic activity of the Cristais

River. Chemosphere, 60: 55–64.

Urek, R. O. and Pazarlioglu, N. K., 2005. Production and stimulation of manganese peroxidase by

immobilized Phanerochaete chrysosporium. Process Biochem., 40: 83–87.

Van Bloois, E., Torres Pazmino, D. E., Winter, R. T. and Fraaije, M. W., 2010. A robust and

extracellular heme-containing peroxidase from Thermobifida fusca as prototype of a

bacterial peroxidase superfamily. Appl. Microbiol. Biotechnol., 86: 1419–1430.

129

Van der Zee, F. P. and Cervantes, F. J., 2009. Impact and application of electron shuttles on the

redox (bio) transformation of contaminants: a review. Biotechnol. Adv., 27: 256–277.

Vandevivere, P. C., Bianchi, R. and Verstraete, W., 1998. Treatment and reuse of wastewater from

the textile wet-processing industry. Rev. Emerging Technologies. J. Chem. Technol.

Biotechnol., 72: 289-302.

Verma, P. and Madamwar, D., 2003. Decolorization of synthetic dyes by a newly isolated strain

of Serratia maerascens. World J. Microbiol. Biotechnol., 19: 615-618.

Vijayaraghavan, K. and Yun, Y. S., 2007. Utilization of fermentation waste (Corynebacterium

glutamicum) for biosorption of Reactive Black 5 from aqueous solution. J. Hazard. Mater.,

141: 45–52.

Vitor, V. and Corso, C. R., 2008. Decolorization of textile dye by Candida albicans isolated from

industrial effluents. J. Ind. Microbiol. Biotechnol., 35: 1353–1357.

Waghmode, T. R., Kurade, M, B., Kabra, A. N. and Govindwar, S. P., 2012. Degradation of

Remazol Red Dye by Galactomyces geotrichum MTCC 1360 leading to increased iron

uptake in Sorghum vulgare and Phaseolus mungo from soil. Biotechnol. Bioeng., 17: 117–

126.

Waghmode, T. R., Kurade, M. B., Khandare, R. V. and Govindwar, S. P., 2011. A sequential

aerobic/microaerophilic decolorization of sulfonated mono azo dye Golden Yellow HER

by microbial consortium GG–BL. Int. Biodeter. Biodegrad., 65: 1024–1034.

Walker, M. C., Van der Donk, W. A., 2016. The many roles of glutamate in metabolism. J. Ind.

Microbiol. Biotechnol., 43: 419–430.

130

Wang, J., Liu, G. F., Lu, H., Jin, R. F., Zhou, J. T. and Lei, T. M., 2012. Biodegradation of Acid

Orange 7 and its auto-oxidative decolorization product in membrane-aerated biofilm

reaction. Int. Biodeter. Biodegrad., 67: 73–77.

Weisburger, J. H., 2002. Comments on the history and importance of aromatic and heterocyclic

amines in public health. Mutat. Res., 30: 506–507.

Wong, P. K. and Yuen, P. Y., 1996. Decolorization and biodegradation of Methyl Red by

Klebsiella pneumoniae RS-13. Water Res., 30: 1736-1744.

Wu, Y., Li, T. and Yang, L., 2012. Mechanisms of removing pollutants from aqueous solutions

by microorganisms and their aggregates: a review. Bioresour. Technol., 107: 10–18.

Xiao, Y. Z., Tu, X. M., Wang, J., Zhang, M., Cheng, Q., Zeng, W. Y. and Shi, Y. Y., 2003.

Purification, molecular characterization and reactivity with aromatic compounds of a

laccase from basidiomycete Trametes sp. strain AH28-2. Appl. Microbiol. Biotechnol., 60:

700–707.

Yang, B., Gao, Y., Yan, Y., Xu, H., and Wang, J., 2018. Degradation characteristics of color index

direct blue 15 dye using iron-carbon micro-electrolysis coupled with H2O2. Int. J. Environ.

Res. Public Health, 15(7): 1523.

Yang, Q., Li, C., Li, H., Li, Y. and Yu, N., 2009. Degradation of synthetic reactive azo dyes and

treatment of textile wastewater by a fungi consortium reactor. Biochem. Eng. J., 43: 225–

230.

Yang, Q., Tao, L., Yang, M. and Zhang, H., 2008. Effects of glucose on the decolorization of

Reactive Black 5 by yeast isolates. J. Environ. Sci., 20: 105–108.

131

Yang, Q., Yediler, A., Yang, M. and Kettrup, A., 2005. Decolorization of an azo dye Reactive

Black 5 and MnP production by yeast isolate: Debaryomyces polymorphus. Biochem. Eng.

J., 24: 249–253.

Yemashova, N. A., Kotova, I. B., Netrusov, A. I. and Kalyuzhnyi, S. V., 2009. Special traits of

decomposition of azo dyes by anaerobic microbial communities. Appl. Biochem.

Microbiol., 45: 176–181.

Yemendzhiev, H., Alexieva, Z. and Krastanov, A., 2009. Decolorization of synthetic dye Reactive

Blue 4 by mycelial culture of white-rot fungi Trametes versicolor 1. Biotechnol. Biotech.

Eq., 23: 1337–1339.

Yu, Z. and Wen, X., 2005. Screening and identification of yeasts for decolorizing synthetic dyes

in industrial wastewater. Int. Biodeter. Biodegrad., 56: 109–114.

Zhang, F., Yediler, A., Liang, X., Kettrup, A., 2004. Effects of dye additives on the ozonation

process and oxidation by products: a comparative study using hydrolysed CI Reactive red

120. Dyes Pigments, 60: 1–7.

Zhao, G., Jiang, L., He, Y., Li, J., Dong, H., Wang, X., Hu, W., 2011. Sulfonated graphene for

persistent aromatic pollutant management. Adv. Mater., 23: 3959–3963.

Zissi, U., Lyberatos, G. and Pavlou, S., 1997. Biodegradation of p-amino benzene by Bacillus

subtilus under aerobic conditions. J. Ind. Microbiol. Biotechnol., 19: 49 -55.

507 © 2018 The Authors Journal of Water Reuse and Desalination | 08.4 | 2018

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Degradation of azo dye by bacterium, Alishewanella sp.

CBL-2 isolated from industrial effluent and its potential

use in decontamination of wastewater

Mehvish Ajaz, Amina Elahi and Abdul Rehman

ABSTRACT

In the present study, Alishewanella sp. CBL-2 was characterized on the basis of biochemical and 16S

rRNA sequencing. The bacterium was found to decolorize 83% Sumifex Tourqi blue within a period of

6 days at 37 �C and pH 7. The predominant form of the enzyme was found to be extracellular (78%) as

compared to intracellular (16%). The breakdown of azo bond was confirmed through thin layer

chromatography (TLC) analysis as new bands and peaks were observed in chromatograms of

extracted metabolites of decolorized samples. Growth of beneficial microbial fauna in the presence

of bacterial decolorized wastewater indicates that this wastewater is safe for the survival of

microorganisms. High bioremediation potential makes Alishewanella sp. CBL-2 an impending

foundation for green chemistry to remove azo dyes from industrial wastewater.

This is an Open Access article distributed under the terms of the Creative

Commons Attribution Licence (CC BY-NC-ND 4.0), which permits copying

and redistribution for non-commercial purposes with no derivatives,

provided the original work is properly cited (http://creativecommons.org/

licenses/by-nc-nd/4.0/)

doi: 10.2166/wrd.2018.065

s://iwaponline.com/jwrd/article-pdf/8/4/507/482391/jwrd0080507.pdf

Mehvish AjazAmina ElahiAbdul Rehman (corresponding author)Department of Microbiology and MolecularGenetics,

University of the Punjab,New Campus, Lahore 54590,PakistanE-mail: [email protected]

Key words | Alishewanella sp. CBL-2, azo dye, decolorization, TLC

INTRODUCTION

Among industrial wastewaters, dye wastewater from textile

and dyestuff industries is one of the most difficult to treat.

This is because dyes usually have a synthetic origin and com-

plex aromatic molecular structures which make them more

stable and more difficult to biodegrade (Seshadri et al. ).

Azo dyes, which are aromatic compounds with one or more

–N¼N– groups, constitute the largest class of synthetic dyes

used in commercial applications (Zollinger ). It is

reported that there are over 100,000 commercially available

dyes with a production of over 7 × 105 metric tons per year

(Mishra & Tripathy ).

Azo dyes are used in different industries, including tex-

tiles, cosmetics, paper, leather, pharmaceuticals, and food,

with an annual consumption of about 0.7 million ton

(Chen et al. ). The two major sources of release of

dyes into the environment are the effluents from textile

processing units and dyestuff manufacturing industries

(O’Neill et al. ). Discharge of highly colored dye con-

taining effluents from textile and dyestuff industries to

neighboring water bodies is causing significant health con-

cerns (Martins et al. ). During the past two decades,

several physico-chemical decolorization techniques have

been reported; few, however, have been accepted by the tex-

tile industries (Okazaki et al. ; da Silva & Faria ).

Their lack of implementation has been largely due to high

cost, low efficiency, and generation of toxic by-products

(Selvam et al. ).

The ability of microorganisms to carry out dye decolor-

ization has recently received much attention. Microbial

decolorization of dyes is a cost-effective method to remove

them from the environment (Moosvi et al. ; Aftab

mailto:[email protected]

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508 M. Ajaz et al. | Degradation of azo dye by bacterium, Alishewanella sp. CBL-2 Journal of Water Reuse and Desalination | 08.4 | 2018

Downloaded frby gueston 21 October

et al. ). The present day bioremediation relies upon the

pollutant degrading capacities of naturally occurring

microbial consortia in which bacteria play a central role

(O’Neill et al. ). Microbial consortia are usually used

without analyzing the constituent microbial populations

for environmental remediation and complexity of the

microbial consortium enables them to act on a variety of

pollutants (Watanabe & Baker ). Recently, the aerobic

decolorization of azo dyes has been reported by a number

of investigators (Adedayo et al. ; Aftab et al. ).

A wide variety of microorganisms is capable of decolor-

ization of a wide range of dyes including fungi: Aspergillus

niger (Fu & Viraraghavan ), Phanerochaete chrysospor-

ium, Aspergillus terricola (Saikia & Gopal ),

P. chrysosporium (Fournier et al. ); yeasts: Saccharo-

myces cerevisiae, Candida tropicalis, C. lipolytica (Aksu &

Dönmez ); algae: Spirogyra species (Gupta et al. ),

Chlorella vulgaris (Acuner & Dilek ), C. sorokiniana

(de-Bashan et al. ), Lemna minuscula (Valderama

et al. ), Scenedesmus obliquus, C. pyrenoidosa and

Closterium lunula (Yan & Pan ), and bacteria:

Escherichia coli NO3 (Chang & Kuo ), Aeromonas

hydrophila (Chen et al. ), Kurthia species (Sani &

Banerjee ).

The present study was aimed at isolating and character-

izing potent bacterium from industrial wastes able to

decolorize azo dyes. Decolorization products were tested

against beneficial microbial flora and dye degraded products

were also analyzed through thin layer chromatography

(TLC).

EXPERIMENTAL

Wastewater sample collection

Wastewater samples were collected in sterilized (autoclaved

at 121 �C, 15 lb/inch2 pressure for 15 min) screw capped

bottles. Some physiochemical parameters of wastewater,

i.e., temperature, pH, color, and smell were also measured.

About six wastewater samples were collected and brought

to the laboratory and then proceeded to bacterial isolation.

For isolation of azo dyes degrading bacteria, 100 μL of

wastewater sample was spread on Luria-Bertani (LB) agar

om https://iwaponline.com/jwrd/article-pdf/8/4/507/482391/jwrd0080507.pdf

2018

plates containing 100 μL of dye stress in the medium. LB

agar plates were prepared by dissolving 1 g NaCl, 1 g Tryp-

tone and 0.5 g yeast extract in 100 mL distilled water, pH

adjusted to 7.0–7.2 and then agar was added in 250 mL

flasks. The medium was autoclaved at 121 �C and 15 lb/

inch2 for 15 min. The growth of bacterial colonies was

observed after 24 h of incubation at 37 �C.

Isolation of azo dye degrading bacteria

Effect of dye on the growth of bacterial isolates was deter-

mined in minimum salt medium (MSM) which contained

(g L�1): KH2PO4, 0.2 g; MgSO4.7H2O, 0.02 g; Na2CO3 1 g;

yeast extract, 0.5 g; in 100 mL distilled water (pH 7.0), then

agar 1.5 g was added, autoclaved, and supplemented with

azo dyes when the temperature reached 50–55 �C. Medium

was poured in autoclaved dried Petri plates under sterilized

conditions. Plates were left to solidify, inoculated with bac-

terial isolates and then incubated again at 37 �C for 24 h.

This experiment was repeated with successfully higher

concentration of dye until the minimum inhibitory concen-

tration (MIC) of the bacterial isolates was obtained.

Morphological, biochemical, and molecular

characterization

The morphological parameters which were checked for bac-

terial isolates include size, shape, color, margin, elevation,

motility, etc. Gram staining and various biochemical tests

were performed according to the scheme given in Cappu-

cino & Sherman (). For molecular characterization,

DNA was isolated (Masneuf-Pomarade et al. ) and

16S rRNA gene was amplified through polymerase chain

reaction (PCR) by using universal bacterial primers 8F

(5-AGAGTTTGATCCTGGCTCAG-3) and 1492R (5-

GGTTACCTTGTTACGACTT-3) (Turner et al. ). PCR

was performed by initial denaturation at 95 �C for 5 min, fol-

lowed by 35 cycles of denaturation at 95 �C for 1 min,

annealing at 55 �C for 1 min and extension at 72 �C for

1 min, with a final extension at 72 �C for 10 min. Fermentas

Gene Jet Gel Extraction kit (#K0691) was used for gene

cleaning of PCR product and then sequenced from 1st

Base, Malaysia. The sequence was then submitted to Gen-

Bank to obtain accession numbers.



Determinations of optimum growth conditions

For optimum growth of bacterial isolate two parameters, i.e.,

temperature and pH were considered. For determination of

optimum temperature, LB broth was prepared in 100 mL

distilled water. Five milliliters of LB broth was added in

four sets, each of three sets of test tubes autoclaved and

inoculated with 50 μL of freshly prepared culture of bac-

terial isolate. Tubes were incubated at 20 �C, 30 �C, 37 �C,

and 45 �C for overnight incubation of 24 h. Absorbance

was measured at 600 nm using UV/Vic spectrophotometer

(PerkinElmer, USA).

For optimum pH, test tubes having 5 mL LB broth were

prepared in six sets, each containing three test tubes and

their pH was adjusted to 5.0, 6.0, 7.0, 8.0, 9.0, and 10.0 by

using 0.1 M NaOH and 0.1 M HCl and then autoclaved.

The tubes were inoculated with 50 μL freshly prepared cul-

ture of bacterial isolate and were incubated at 37 �C for

24 h and their absorbance was measured at 600 nm using

spectrophotometer.

Effect of dye on bacterial growth

Growth curves of bacterial isolate were determined by using

MSM. The flasks containing 100 μL of dye were designated

as ‘treated’ and flasks without dye were designated as ‘con-

trol’. In this way, 100 mL MSM was taken into 250 mL

flasks, autoclaved and then inoculated with 50 μL of freshly

prepared bacterial culture. The cultures were incubated at

37 �C in a shaker at 100 rpm. An aliquot of culture was

taken out in an oven sterilized test tube at regular intervals

of 0, 4, 8, 12, 16, 20, and 24 h.

Dye decolorization

Decolorization experiment was performed in 250 mL

Erlenmeyer flasks containing 100 mL of MSM and was

inoculated with 100 μL of freshly prepared bacterial culture.

The dye was added to the culturing medium 100 μL and

200 μL from stock solution after 24 h of growth and the

flask containing no bacterial culture was used as control.

The flasks were incubated at 37 �C, 3 ml culture medium

was withdrawn after 1, 2, 3, 4, 5, and 6 days and centrifuged

at 10,000 rpm for 15 min to separate the bacterial cell mass.


Decolorization was determined by measuring the absor-

bance of decolorization medium at 465 nm (Kalyani et al.

). Percentage decolorization was calculated as follows:

Decolorization(%)

¼ Initial absorbance�Observed absorbanceInitial absorbance

× 100

Enzyme assay

To prepare the crude cell-free extract, the bacterial cultures

were grown with azo dyes (100 μL from stock solution of

dye) in 100 mL MSM for 96 h at 37 �C. Cells were harvested

by centrifugation at 6,000 × g for 15 min. For intracellular

enzyme assay, pellet was washed with lysis buffer (Mercap-

toethanol; SDS) and was suspended in 1 mL of the same

buffer. Cells were disrupted by sonication for 15 s with

60 s interval (5 cycles) in cold conditions. The resultant

homogenate was centrifuged at 10,000 × g for 5 min. The

supernatant was used as intracellular crude extract. While

for extracellular crude extract, the supernatant was mixed

with 60% ammonium salt, and the mixture was incubated

overnight in a refrigerator at 4 �C. The overnight chilled mix-

ture was then centrifuged at 10,000 rpm for 15 min. The

aliquot was then used as extracellular crude enzyme. The

activity of azoreductase was determined by a slightly modi-

fied method of Moutaouakkil et al. (). The reaction

mixture consisted of: sodium acetate, 500 μL (pH 5); auto-

claved water, 500 μL; ABTS, 100 μL; and crude enzyme

100 μL. The reaction was allowed to proceed for 30 min at

30 �C and optical density was measured at 465 nm. For con-

trol reaction, distilled water was used instead of crude

enzyme.

Thin layer chromatography

After the enzyme reaction the products were used for TLC.

For this purpose, a silica plate was used. The spotting was

done with the help of a Pasteur pipette. The spots were

dried and then allowed to run into a chromatographic

tank through a mobile phase (chloroform, 30 mL; ethyl acet-

ate, 30 mL; acetic acid, 0.6 mL). The silica plate was then

observed under UV illuminator at wavelength 254 and

366 nm.

Figure 1 | Change in color of Sumifex Tourqi blue as compared to the control in flask

containing culture of Alishewanella sp. CBL-2.



Microbial toxicity

The toxic effect of decolorized dye sample was checked on

agriculturally important soil microflora, i.e., Bacillus subti-

lis (z-66), Pseudomonas aeruginosa (PA1), and Bacillus

cereus (T358-2) (Mali et al. ). L-agar plates were

swabbed with these bacteria and then a well of 2 mm was

made which was filled by centrifuged decolorized broth.

The plates were then incubated at 37 �C for 24 h. The

zone surrounding the well showed the index of toxicity of

the decolorized sample.

Statistical analysis

Observations were made and all the experiments run in tri-

plicate. At least three separate flasks were usually

maintained for one treatment. Each time three readings

were taken, their mean, and standard error of the mean

were calculated.

RESULTS AND DISCUSSION

Sampling and screening of dye decolorizing bacteria

A total of six wastewater samples with physiochemical par-

ameters was collected (Table S1, available with the online

version of this paper). Bacterial cultures were isolated, pur-

ified, and screened for the decolorization of azo dye. Of

all the cultures tested, one bacterial isolate (CBL-2) was

selected on the basis of dye tolerance, i.e., resistance in

MSM containing 1% of reactive azo dye Sumifex Tourqi

Blue (Figure 1).

Bioremediation is becoming important because it is cost-

effective and environmentally friendly, and produces less

sludge (Robinson et al. ; Chen et al. ). Microbial

decolorization of dyes is a cost-effective procedure to

remove them from the environment (Moosvi et al. ).

The biodegradation ability of bacteria is assumed to be

associated with the production of lignolytic enzymes such

as lignin peroxidase and laccase. The presence of laccase-

like enzymes has been reported in bacteria (Givaudan

et al. ). The ability of bacteria to metabolize azo dyes

has been investigated by a number of research groups.


2018

Many microorganisms are capable of decolorizing the azo

dyes, including Gram-positive and Gram-negative bacteria

(Sani & Banerjee ; Kodam et al. ; Moosvi et al.

).

Biochemical and molecular characterization of

bacterial isolate

Initially, the bacterial isolate was identified on the basis

of morphological and biochemical characteristics

(Table 1). The sequence of 16S rRNA gene of bacterial

isolate showed 95% homology with the sequence of 16S

rRNA gene of Alishewanella sp. The sequence was

then submitted to GenBank under accession number

JN546609.

Determination of growth conditions

The most suitable temperature and pH of bacterial isolate

were found to be 37 �C and 7 (Figure S1, available online).

The strain CBL-2 showed continuous increase in growth

rate until the stationary phase was reached, while the con-

trol showed a steady increase in the growth. The control

was without dye and for the treated Turqi blue dye plus

inoculum was used (Figure 2). Asada et al. () isolated

Table 1 | Morphological and biochemical characteristics of Alishewanella sp. CBL-2

Alishewanella sp. CBL-2

Morphological characteristics

Shape Rod

Motility Non-motile

Color Yellowish

Elevation Raised

Size 2 μm

Margin Smooth

Gram staining Gram negative

Biochemical characteristics

Indole production �ve

Methyl red reaction �ve

Voges Proskauer reaction �ve

Citrate use �ve

MacConkey agar �ve

Catalase activity þve

Oxidase activity þve

EMB agar �ve

þve: positive, �ve: negative.



the azo dye decolorizing bacteria which were Gram-nega-

tive and showed maximum growth at 34 �C–37 �C. Banat

et al. () also observed that various textile and other

dye effluents are produced at relatively high temperatures

(50–60 �C), so temperature will be an important factor in

the real application of biosorption by biomass in the

future.

Figure 2 | Effect of Sumifex Tourqi blue on the growth (OD) of Alishewanella sp. CBL-2

incubated at 37 �C for a period of 24 h.


Dye decolorization

The efficient removal of dye is necessary and significant

for the protection of the environment. In the present

study, bacterial strain CBL-2 was able to decolorize 80

and 83% of 100 and 200 μL mL�1 dye stress, respectively

(Figure 3). Sheth & Dave () reported P. aeruginosa

exhibited 91% decolorization of azo dye. Wu et al.

() reported that Shewanella oneidensis WL-7 was

capable of decolorizing the Reactive Black 5 from the

medium containing 100 μM after 12 h. Decolorization of

azo dyes by bacterial isolates has been reported by

many researchers (Hong et al. ; Xu et al. ;

Khalid et al. ).

Enzyme assay

Bacterial degradation of azo dyes is generally considered

a specific reaction by azo reductase under aerobic

condition or a nonspecific reduction process under

anaerobic conditions. In the present study, both extra-

and intracellular enzyme activities were determined by

ABTS method. Alishewanella sp. CBL-2 showed high

extracellular enzyme activity (78%) as opposed to intra-

cellular enzyme activity (16%), so the predominant form

of the enzyme was extracellular (Figure 4). Maier et al.

() reported that a thermoalkalophilic Bacillus sp.

was able to reduce a large structural variety of azo dyes.

Kolekar et al. () reported that Alishewanella sp.

Figure 3 | Alishewanella sp. CBL-2 potential to decolorize Sumifex Tourqi blue incubated

at 37 �C for a period of 6 days.

Figure 4 | Comparison between extra- and intracellular enzyme (azoreductase) activities

of Alishewanella sp. CBL-2 using ABTS as a substrate.



strain KMK6 was able to degrade efficiently a mixture of

textile dyes (0.5–2.0 g L�1) within 8 h. An initial 28%

reduction in COD was observed immediately after deco-

lorization at static anoxic conditions, which on further

incubation at shaking conditions, reduced by 90%.

Figure 5 | TLC chromatogram of extracted metabolites of Alishewanella sp. CBL-2

decolorized dye sample visualizes UV range of 254 and 366 nm.


2018

Partially purified azoreductase was able to utilize different

azo dyes as substrates.

Dye degraded products analysis

The presence of bands with different Rf value in the control

and treated sample clearly revealed that dye was degraded

into its products (Figure 5). Another study from this labora-

tory showed the presence of bands with Rf value of 0.96 and

0.94 when observed under UV range of 254 and 366 nm

while the Rf value of control dye was 0.83 (unpublished

data). Singh et al. () reported that two additional

bands of Rf value 0.63 and 0.965 were observed in the

TLC chromatogram of a decolorized sample of Direct

orange 16 by Micrococcus luteus strain SSN2 when

observed under UV range of 254 nm while the dye had a

band of Rf value 0.74. Similarly, Sahasrabudhe et al. ()

reported a band of Rf value 0.71 in the TLC chromatogram

of extracted metabolites from a decolorized sample of Direct

red by Enterococcus faecalis YZ66 while the original dye

had a band of Rf value 0.97. Likewise, a single band of Rf

value 0.83 was observed in TLC chromatogram of mono

azo dye Amarnath while the extracted metabolites from Aci-

netobacter calcoaceticus NCIM 2890 decolorized sample

had three bands of Rf value 0.93, 0.82, and 0.79 (Ghodake

et al. ).

Microbial toxicity

There was no zone of inhibition found around the well

which was a clear indication that the biodegraded waste-

water is safe for the growth of microbial species (Figure 6).

Ilyas & Rehman () reported two fungal species, i.e.,

Aspergillus niger and Nigrospora sp. whose decolorized

sample was proved safe for the microbial species. Similarly,

the dye degradation products by Alishewanella sp. strain

KMK6 were non-toxic compared to the dye mixture in gen-

otoxicity assessment assay (Kolekar et al. ).

CONCLUSIONS

Alishewanella sp. CBL-2, characterized on the basis of bio-

chemical tests and 16S rRNA sequencing, has promising

Figure 6 | Bacterial decolorized broth used for incubation of (a) B. cereus (T358-2), (b) P. aeruginosa (PA1), and (c) B. subtilis (z-66) at 37 �C for 48 h. No zone of inhibition was observed in

any case.



potential to decolorize Sumifex Tourqi blue (83%) after 6

days of incubation at 37 �C and pH 7. Alishewanella sp.

CBL-2 had shown maximum enzyme activity in extracellu-

lar assay (78%) as compared to the intracellular assay

(16%). This degradation was confirmed through TLC. It

was also proved that microbially treated dye wastewater is

safe for growth of beneficial microbial flora. Alishewanella

sp. CBL-2 can be used to ameliorate wastewater containing

azo dyes by utilizing the decolorization potential ability of

the bacterium.

ACKNOWLEDGEMENTS

This work was supported by the Research Cell, Quaid-e-

Azam Campus, Punjab University, Lahore-54590, Pakistan


(Research grant number, 34/2014-15) which is gratefully

acknowledged. The authors have declared that no

competing interests exist.

REFERENCES

Acuner, E. & Dilek, F. B. Treatment of tectilon yellow 2G byChlorella vulgaris. Process. Biochem., 39, 623–631.

Adedayo, O., Javadpour, S., Taylor, C., Anderson, W. A. & Moo-Young, M. Decolorization and detoxification of methylred by aerobic bacteria from a wastewater treatment plant.World J. Microbiol. Biotechnol. 20, 545–550.

Aftab, U., Khan, M. R., Mahfooz, M., Ali, M., Hassan, S. H. &Rehman, A. Decolorization and degradation of textileazo dyes by Corynebacterium sp. isolated from industrialeffluent. Pakistan J. Zool. 43, 1–8.

http://dx.doi.org/10.1016/S0032-9592(03)00138-9

http://dx.doi.org/10.1016/S0032-9592(03)00138-9

http://dx.doi.org/10.1023/B:WIBI.0000043150.37318.5f

http://dx.doi.org/10.1023/B:WIBI.0000043150.37318.5f



Aksu, Z. & Dönmez, G. A comparative study on thebiosorption characteristics of some yeasts for Remozal Bluereactive dye. Chemosphere 50, 1075–1083.

Asada, S., Amoozegarb, M. A., Pourbabaeec, A. A., Sarboloukic,M. N. & Dastgheib, S. M. M. Decolorization of textileazo dyes by newly isolated halophilic and halotolerantbacteria. Bioresour. Technol. 98 (11), 2082–2088.

Banat, I. M., Nigam, P., Singh, D. & Marchant, R. Microbialdecolorization of textile-dye-containing effluents: a review.Bioresour. Technol. 58, 217–227.

Cappucino, J. G. & Sherman, N. Microbiology. A LaboratoryManual, 6th edn. Pearson Education, Benjamin Cummings,San Francisco.

Chang, J. S. & Kuo, T. S. Kinetics of bacterial decolorizationof azo dye with Escherichia coliNO3. Bioresour. Technol. 75,107–111.

Chen, K. C., Wu, J. Y., Liou, D. J. & Hwang, S. C. J. Decolorization of the textile azo dyes by newly isolatedbacterial strains. J. Biotechnol. 101, 57–68.

da Silva, C. G. & Faria, J. L. Photochemical andphotocatalytic degradation of an azo dye in aqueous solutionby UV irradiation. Photochem. Photobiol. A: Chem. 155,133–143.

de-Bashan, L. E., Moreno Hernandez, J. P. & Bashan, Y. Removal of ammonium and phosphorus ions from syntheticwastewater by the microalgae Chlorella vulgariscoimmobilized in alginate beads with the microalgae growth-promoting bacterium Azospirillum brasilense. Water Res. 36,2941–2948.

Fournier, D., Halasz, A., Thiboutot, S., Ampleman, G., Dominic,M. & Hawari, J. Biodegradation of octahydro-1,3,5,7-tetranitro-1,3,5,7-tetrazocine (HMX) by Phnerochaetechrysosporium. New insight into the degradation pathway.Environ. Sci. Technol. 38, 4130–4133.

Fu, Y. & Viraraghavan, T. Dye biosorption sites in Aspergillusniger. Bioresour. Technol. 82, 139–145.

Ghodake, G., Jadhav, U., Tamboli, D., Kagalkar, A. & Govindwar,S. Decolorization of textile dyes and degradation ofmono-azo dye Amaranth by Acinetobacter calcoaceticusNCIM 2890. Indian J. Microbiol. 51, 501–508.

Givaudan, A., Effosse, A., Faure, D., Potier, P., Bouillant, M. L. &Bally, R. Polyphenol oxidase in Azospirillum lipoferumisolated from rice rhizosphere: evidence for laccase activityin non-motile strains of Azospirillum lipoferum. FEMSMicrobiol. Lett. 108, 205–210.

Gupta, V. K., Rastogi, A., Saini, V. K. & Jain, N. Biosorptionof copper(II) from aqueous solutions by Spirogyra species.J. Colloid Interf. Sci. 296, 59–63.

Hong, Y., Guo, J., Xu, Z., Chen, X. & Sun, G. Respirationand growth Sheanella oneidensis S12 with an azocompound as the sole electron. Appl. Environ. Microbiol. 99,4635–4641.

Ilyas, S. & Rehman, A. Decolorization and detoxification ofSynozol red HF-6BN azo dye, by Aspergillus niger andNigrospora sp. Iranian J. Environ. Health Sci. Eng. 10, 12.


2018

Kalyani, D. C., Patil, P. S., Jadhhav, J. P. & Govindwar, S. P. Biodegradation of reactive textile dye Red BL1 by an isolatedbacterium Pseudomonas sp. SUKI. Bioresour. Technol., 99,4635–4641.

Khalid, A., Arshad, M. & Crowley, D. E. Accelerateddecolourization of structurally different azo dyes by newlyisolated bacterial strains. Appl. Microbiol. Biotechnol. 78,1838–1852.

Kodam, K. M., Soojhawon, I., Lohande, P. D. & Gawai, K. R. Microbial decolorization of reactive azo dyes under aerobicconditions. Word J. Microbiol. Biotechnol. 21, 367–370.

Kolekar, Y. M., Konde, P. D., Markad, V. L., Kulkarni, S. V.,Chaudhari, A. U. & Kodam, K. M. Effective bioremovaland detoxification of textile dye mixture by Alishewanella sp.KMK6. Appl. Microbiol. Biotechnol. 97 (2), 881–889.

Maier, J., Kandelbauer, A., Erlacher, A., Cavaco-Paulo, A. &Gubits, G. A new alkali thermostable azoreductase fromBacillus sp. strain SF. Appl. Environ. Microbiol. 70, 837–844.

Mali, P. L., Mahajan, M. M., Patil, D. P. & Kulkarni, M. V. Biodecolorization of members of triphenyl methane and azogroup of dyes. J. Sci. Ind. Res. 59, 221–224.

Martins, M. A. M., Queiroz, M. J., Silvestre, A. J. D. & Lima, N. Relationship of chemical structure of textile dye on thepreadaptation medium and the potentialities of theirbiodegradation by Phanerochaete chrysosporium. Res.Microbiol. 153, 361–368.

Masneuf-Pomarade, I., Jeuneb, C. L., Durrensc, P., Lollierb, M.,Aigled, M. & Dubourdieu, D. Molecular typing of wineyeast strains Saccharomyces bayanus var. uvarum usingmicrosatellite markers. Syst. Appl. Microbiol. 30, 75–82.

Mishra, G. & Tripathy, M. A critical review of the treatmentsfor decolorization of textile effluent. Colourage 40, 35–38.

Moosvi, S., Keharia, H. & Madamwar, D. Decolorization oftextile dye Reactive Violet 5 by a newly isolated bacterialconsortium RVM 11.1. World J. Microbiol. Biotechnol. 21,667–672.

Moutaouakkil, A., Zeroual, Y., Dzayri, F. Z., Talbi, M., Lee, K. &Blaghen, M. Purification and partial characterization ofazoreductase from Enterobacter agglomerans. Arch.Biochem. Biophys. 413, 139–146.

O’Neill, C., Hawkes, F. R., Hawkes, D. L., Lourenco, N. D.,Pinheiro, H. M. & Delee, W. Color in textile effluents:sources, measurement, discharge consents and simulation: areview. Chem. Technol. Biotechnol. 74, 1009–1018.

O’Neill, C., Lopez, A., Esteves, S., Hawkes, F. R., Hawkes, D. L. &Wilcox, S. Azo-dye degradation in an anaerobic–aerobictreatment system operating on simulated textile effluent.Appl. Microbiol. Biotechnol. 53, 249–254.

Okazaki, S., Nagasawa, S., Goto, M., Furusaki, S., Wariishi, H. &Tanaka, H. Decolorization of azo dye andanthraquinone dye in hydrophobic organic media usingmicroperoxidase-11 entrapped in reverse micelles. Biochem.Engin. 12, 237–241.

Robinson, T., McMullan, G., Marchant, R. & Nigam, P. Remediation of dyes in textile effluent: a critical review on

http://dx.doi.org/10.1016/S0045-6535(02)00623-9

http://dx.doi.org/10.1016/S0045-6535(02)00623-9

http://dx.doi.org/10.1016/S0045-6535(02)00623-9

http://dx.doi.org/10.1016/j.biortech.2006.08.020



http://dx.doi.org/10.1016/S0960-8524(96)00113-7

http://dx.doi.org/10.1016/S0960-8524(96)00113-7

http://dx.doi.org/10.1016/S0960-8524(00)00049-3

http://dx.doi.org/10.1016/S0960-8524(00)00049-3

http://dx.doi.org/10.1016/S0168-1656(02)00303-6

http://dx.doi.org/10.1016/S0168-1656(02)00303-6

http://dx.doi.org/10.1016/S1010-6030(02)00374-X

http://dx.doi.org/10.1016/S1010-6030(02)00374-X

http://dx.doi.org/10.1016/S1010-6030(02)00374-X

http://dx.doi.org/10.1016/S0043-1354(01)00522-X

http://dx.doi.org/10.1016/S0043-1354(01)00522-X

http://dx.doi.org/10.1016/S0043-1354(01)00522-X

http://dx.doi.org/10.1016/S0043-1354(01)00522-X

http://dx.doi.org/10.1021/es049671d



http://dx.doi.org/10.1016/S0960-8524(01)00172-9

http://dx.doi.org/10.1016/S0960-8524(01)00172-9

http://dx.doi.org/10.1007/s12088-011-0131-4

http://dx.doi.org/10.1007/s12088-011-0131-4

http://dx.doi.org/10.1007/s12088-011-0131-4

http://dx.doi.org/10.1111/j.1574-6968.1993.tb06100.x



http://dx.doi.org/10.1016/j.jcis.2005.08.033

http://dx.doi.org/10.1016/j.jcis.2005.08.033

http://dx.doi.org/10.1186/1735-2746-10-12

http://dx.doi.org/10.1186/1735-2746-10-12

http://dx.doi.org/10.1186/1735-2746-10-12



http://dx.doi.org/10.1007/s00253-007-1302-4

http://dx.doi.org/10.1007/s00253-007-1302-4

http://dx.doi.org/10.1007/s00253-007-1302-4

http://dx.doi.org/10.1007/s11274-004-5957-z

http://dx.doi.org/10.1007/s11274-004-5957-z

http://dx.doi.org/10.1007/s00253-012-3983-6

http://dx.doi.org/10.1007/s00253-012-3983-6

http://dx.doi.org/10.1007/s00253-012-3983-6

http://dx.doi.org/10.1128/AEM.70.2.837-844.2004

http://dx.doi.org/10.1128/AEM.70.2.837-844.2004

http://dx.doi.org/10.1016/S0923-2508(02)01332-3

http://dx.doi.org/10.1016/S0923-2508(02)01332-3

http://dx.doi.org/10.1016/S0923-2508(02)01332-3

http://dx.doi.org/10.1016/j.syapm.2006.02.006



http://dx.doi.org/10.1007/s11274-004-3612-3

http://dx.doi.org/10.1007/s11274-004-3612-3

http://dx.doi.org/10.1007/s11274-004-3612-3

http://dx.doi.org/10.1016/S0003-9861(03)00096-1

http://dx.doi.org/10.1016/S0003-9861(03)00096-1

http://dx.doi.org/10.1002/(SICI)1097-4660(199911)74:11%3C1009::AID-JCTB153%3E3.0.CO;2-N



http://dx.doi.org/10.1007/s002530050016

http://dx.doi.org/10.1007/s002530050016

http://dx.doi.org/10.1016/S1369-703X(02)00074-8

http://dx.doi.org/10.1016/S1369-703X(02)00074-8

http://dx.doi.org/10.1016/S1369-703X(02)00074-8

http://dx.doi.org/10.1016/S0960-8524(00)00080-8



current treatment technologies with a proposed alternative.Bioresour. Technol. 77, 247–255.

Sahasrabudhe, M. M., Saratale, R. G., Saratale, G. D. & Pathade,G. R. Decolorization and detoxification of sulfonatedtoxic diazo dye C.I. Direct red 81 by Enterococcus faecalisYZ 66. J. Environ. Health Sci. Eng. 12, 151.

Saikia, N. & Gopal, M. Biodegradation of β-cyfluthrin byfungi. J. Agr. Food Chem. 52, 1220–1223.

Sani, K. R. & Banerjee, U. C. Decolorization oftriphenylmethane dyes and textile and dyestuff effluent byKurthia sp. Enzyme Microb. Technol. 24, 433–437.

Selvam, K., Swaminathan, K. & Keo-Sang, C. Decolorizationof azo dyes and a dye industry effluent by a white rot fungusThelephora sp. Bioresour. Technol. 88, 115–119.

Seshadri, S., Bishop, P. L. & Agha, A. M. Anaerobic/aerobictreatment of selected azo dyes in wastewater. Waste Manage.15, 127–137.

Sheth, N. T. & Dave, S. R. Optimization for enhanceddecolorization of Reactive Red BS C.I. 111 by Pseudomonasaeruginosa NGKCTS. Biodegradation 20, 827–836.

Singh, S., Chatterji, S., Nandini, P. T., Prasad, A. S. A. & Rao,K. V. B. Biodegradation of azo dye Direct Orange 16 byMicrococcus luteus strain SSN2. Int. J. Environ. Sci. Technol.12, 2161–2168.


Turner, S., Pryer, K. M., Miao, V. P. W. & Palmer, J. D. Investigating deep phylogenetic relationships amongcyanobacteria and plastids by small subunit rRNA sequenceanalysis. J. Eukar. Microbiol. 46, 327–338.

Valderama, L. T., Del Campo, C. M., Rodriguez, C. M., de-Bashan,L. E. & Bashan, Y. Treatment of recalcitrant wastewaterfrom ethanol and citric acid production using the microalgaChlorella vulgaris and the macrophyte Lemna minuscule.Water Res. 36, 4185–4192.

Watanabe, K. & Baker, P. W. Environmentally relevantmicroorganisms. Biosci. Bioeng. 89, 1–11.

Wu, J., Kim, K.-S., Sung, N.-C., Kim, C.-H. & Lee, Y. C. Isolation and characterization of Shewanella oneidensis WL-7 capable of decolorizing azo dye Reactive Black 5. J. Gen.Appl. Microbiol. 55, 51–55.

Xu, M., Guo, J., Xu, Z., Chen, X. & Sun, G. Respiration andgrowth Shewanella oneidensis S12 with an azo compound asthe sole electron. Appl. Environ. Microbiol. 73, 64–72.

Yan, H. & Pan, G. Increase in biodegradation of dimethylphthalate by Closterium lunula using inorganic carbon.Chemosphere, 55, 1281–1285.

Zollinger, H. Colour Chemistry Synthesis Properties andApplication of Organic Dyes and Pigments. VCH, New York,pp. 92–102.

First received 11 November 2017; accepted in revised form 29 January 2018. Available online 20 March 2018

http://dx.doi.org/10.1016/S0960-8524(00)00080-8

http://dx.doi.org/10.1186/s40201-014-0151-1

http://dx.doi.org/10.1186/s40201-014-0151-1

http://dx.doi.org/10.1186/s40201-014-0151-1

http://dx.doi.org/10.1021/jf0349580

http://dx.doi.org/10.1021/jf0349580

http://dx.doi.org/10.1016/S0141-0229(98)00159-8

http://dx.doi.org/10.1016/S0141-0229(98)00159-8

http://dx.doi.org/10.1016/S0141-0229(98)00159-8

http://dx.doi.org/10.1016/S0960-8524(02)00280-8

http://dx.doi.org/10.1016/S0960-8524(02)00280-8

http://dx.doi.org/10.1016/S0960-8524(02)00280-8

http://dx.doi.org/10.1016/0956-053X(94)90005-1

http://dx.doi.org/10.1016/0956-053X(94)90005-1

http://dx.doi.org/10.1007/s10532-009-9270-2

http://dx.doi.org/10.1007/s10532-009-9270-2

http://dx.doi.org/10.1007/s10532-009-9270-2

http://dx.doi.org/10.1007/s13762-014-0588-x

http://dx.doi.org/10.1007/s13762-014-0588-x




http://dx.doi.org/10.1016/S0043-1354(02)00143-4

http://dx.doi.org/10.1016/S0043-1354(02)00143-4

http://dx.doi.org/10.1016/S0043-1354(02)00143-4

http://dx.doi.org/10.1016/S1389-1723(00)88043-3

http://dx.doi.org/10.1016/S1389-1723(00)88043-3

http://dx.doi.org/10.2323/jgam.55.51

http://dx.doi.org/10.2323/jgam.55.51

http://dx.doi.org/10.1128/AEM.01415-06



http://dx.doi.org/10.1016/j.chemosphere.2003.12.019

http://dx.doi.org/10.1016/j.chemosphere.2003.12.019

Ajaz et al. AMB Expr (2019) 9:64 https://doi.org/10.1186/s13568-019-0788-3

ORIGINAL ARTICLE

Degradation of azo dyes by Alcaligenes aquatilis 3c and its potential use in the wastewater treatmentMehvish Ajaz1, Abdul Rehman1*, Zaman Khan2, Muhammad Atif Nisar3 and Syed Hussain4

Abstract

In the present study, Alcaligenes aquatilis was found to decolorize 82% Synazol red 6HBN after incubation of 4 days at 37 °C and pH 7. Maximum decolorization was found under static conditions by using saw dust and yeast extract as carbon and nitrogen source. It also showed promising potential to decolorize mixture of multiple dyes at a rate of more than 86% in 5 days. Decolorization of dye had positive influence on the growth of bacterium as growth rate was increased along with decolorization. The cleavage of azo bond was confirmed through TLC, HPLC and GC–MS analysis. The dye metabolites produced during bacterial treatment are linked to various pathways including ATP synthesis pro-cess. The absence of peaks of wavelength 1612/cm and 1532/cm in bacterially treated FTIR sample demonstrated the cleavage of azo bond. Microbial growth in decolorized dye wastewater shows that bacterially decolorized wastewater is unharmful for the growth of micro-flora. The high decolorization ability of A. aquatilis 3c to convert toxic azo dyes into useful end products may find potential applications in the environmental biotechnology.

Keywords: Azo dyes, Decolorization, A. aquatilis 3c, HPLC, FTIR, GC–MS

© The Author(s) 2019. This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creat iveco mmons .org/licen ses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made.

IntroductionIndustrialization is the backbone for the welfare and development of a country. Apart from its beneficial effects on the economy of a country it also exerts harmful effect on the environment and organisms. The discharge of wastewater from various industries becomes a contin-uous source of environmental pollution. Synthetic dyes, which are widely used in such industries, are a major por-tion of this discharged wastewater. These dyes are even more harmful for the organisms in comparison to other pollutants because of their resistance potential (de Souza et al. 2010).

Azo dyes are the largest class of synthetic dyes which are aromatic in nature structurally having one or more –N=N– bond. These dyes are rapidly used in various industries like textile, cosmetics, paper, food and phar-maceutical industry while the textile industry is its largest

consumer (Alalewi and Jiang 2012). Their extensive com-mercial use is because of the reason that they are easily synthesized and economical for the users (Saratale et al. 2009a).

The loss of dyes from fabrics ranges from 2% for basic dyes to as high as 50% for reactive dyes as all the dye does not bind to the fabric (O’Neil et al. 1999; Pandey et al. 2007). The discharge of dye contaminated wastewater in aquatic ecosystems arises many serious problems. It causes obstruction in light penetration; oxygen transfer is reduced in water bodies, poses acute toxic effects on aquatic flora and fauna and also causes severe environ-mental problems (Solis et al. 2012).

There is an extensive use of physicochemical tech-niques like coagulation and flocculation worldwide to treat industrial effluents (Verma et al. 2012). Certain dis-advantages are associated with the use of such techniques like usage of chemicals, sludge production which serve as source of secondary pollution, production of by products and high cost (Jadhav et al. 2007). In comparison to these physiochemical processes, use of variety of microorgan-isms including yeast, fungi and bacteria is more feasible

Open Access

*Correspondence: [email protected] 1 Department of Microbiology and Molecular Genetics, University of the Punjab, New Campus, Lahore 54590, PakistanFull list of author information is available at the end of the article

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of 12Ajaz et al. AMB Expr (2019) 9:64

as they are environmental friendly and their use result in the cleavage of azo bond which leads to formation of colorless aromatic amines (Kaushik and Malik 2010; Das and Mishra 2016; Meerbergen et al. 2018). The resulting products such as aromatic amines are further degraded to simpler non-toxic forms by multiple-step bioconver-sion occurring aerobically or anaerobically (Shah et al. 2012; Singh et al. 2014).

The current study was aimed at to isolate and char-acterize bacterium from industrial effluents capable to decolorize azo dyes. Optimum growth conditions of the organism were determined and bacterially degraded dye products were also analysed through TLC, HPLC, FTIR and GC–MS.

Materials and methodsSample collection and bacterial isolationIndustrial effluent samples were collected in autoclaved screw-capped bottles from Kot Lakhpat industrial estate, Lahore, Pakistan. Physicochemical parameters such as pH, temperature, as well as color of the samples were also noted at the time of sample collection. The samples were serially diluted and plated on the L-agar plates. L-agar medium was prepared by dissolving tryptone (10 g), yeast extract (5 g), NaCl (5 g) and agar (15 g) in 1000 ml of dis-tilled water. pH of the medium was adjusted to 7.

Evaluation of dye degrading potentialDye degrading potential was determined by inoculating the bacterial isolate in 250 ml Erlenmeyer flask having 100 ml of mineral salt medium (MSM) whose composi-tion is (g/l): (NH4)SO4, 0.28; MgSO4·7H2O, 0.04;NH4Cl, 0.23; KH2PO4, 0.067; FeCl3·6H2O, 0.005; CaCl2·2H2O, 0.022; yeast extract, 0.2; NaCl, 0.15; NaHCO3, 1.0 and 1 ml/l of a trace element solution containing (g/l): MnCl2·4H2O, 0.1;ZnSO4·7H2O, 0.01; CuSO4·5H2O, 0.392; NaB4O7·10 H2O, 0.177; CoCl2·6H2O, 0.248 and NiCl2·6H2O, 0.02 (Parshetti et al. 2006) with glucose and yeast extract (1% each) used as carbon and nitrogen source respectively. The medium was supplemented with dye (Synazol red 6HBN) at a concentration of 50 mg/l for 4 days of incubation at 37 °C. Synazol red 6HBN, Congo red, Methyl red, Phenol red, Brilliant black and Navy blue were purchased from Sigma-Aldrich. All other chemicals were of analytical grade purity.

Bacterial characterizationThe bacterial morphological parameters and biochemi-cal tests were performed according to protocols given in Cappucino and Sherman (2008). For 16S rRNA ribotyp-ing, DNA was isolated (Masneuf-Pomarade et al. 2007)

and 16S rRNA gene was amplified through PCR by using universal bacterial primers (Turner et al. 1999). PCR was performed according to Rehman et al. (2007) and PCR product was purified by Fermentas Gene Jet Gel Extrac-tion kit (#K0691). The purified product was sequenced and submitted to GenBank for obtaining accession numbers. Phylogenetic analysis was done using MEGA7 (Kumar et al. 2016). Phylogenetic tree was constructed using neighbor joining method with 1000 replicates.

Determination of optimal growth conditionsThe physical parameters which were favorable for the bacterial growth i.e., temperature and pH were analyzed by growing bacterium at different temperature and pH. For optimum temperature determination, bacterial iso-late was grown in LB broth and incubated at different temperature i.e., 20, 30, 37 and 45 °C and for optimum pH, the bacterial isolate was grown in LB broth with pH values i.e., 5, 6, 7, 8, 9 and 10. After 24 h of incubation, absorbance was recorded at 600 nm by using spectropho-tometer. In order to determine growth pattern of bacte-rial isolate, LB medium was inoculated with log phase grown bacterial culture (1 ml). Optical density was deter-mined by spectrophotometer at 600 nm at the time of inoculation and then after regular interval of 4 h up to 28 h of growth at optimum temperature and pH.

Decolorization experimentsIn order to optimize the decolorization conditions decol-orization experiments were performed at various condi-tions i.e., temperature (20, 30, 37, 45 and 50 °C), pH (5, 6, 7, 8, 9, 10), incubation condition (static and shaking), carbon sources (saw dust, sugarcane baggase and wheat bran) and nitrogen source (yeast extract, beef extract and peptone) source, inoculum percentage (2%, 4%,6%, 8% and 10%) and dye concentration (3, 5, 7, 10, 20 and 50 mg/l). A volume of 100 ml of MSM was taken in 250 ml of Erlenmeyer flask which was then inoculated with 2% of bacterial suspension and incubated at respec-tive condition. The stock solution of dye was added into the MSM to obtain a final dye concentration of 50 mg/l. Aliquot was taken out at 0 h and after 3 days in order to measure initial and final absorbance, respectively. The aliquot was centrifuged before measuring the opti-cal density at 465 nm. Decolorization percentage of the sample was measured by using following formula. All the treatments and controls were carried out in triplicates.

Decolorization (%)

=

Initial absorbance− Final absorbance

Initial absorbance× 100


Effect of decolorization on growth of bacterial isolateIn order to check the effect of decolorization on growth of bacterial isolate, the bacterium was grown in MSM containing dye concentration of 50 mg/l. The aliquot was obtained after every day up to 5 days and optical den-sity was measured at 600 nm in order to find out growth ratio. Decolorization was also calculated by measuring the optical density of supernatant at 465 nm after centrif-ugation of the sample.

Decolorization of multiple dyesThe bacterial isolate was checked for its ability to decol-orize a mixture of azo dyes i.e., Congo red, Methyl red, Phenol red, Brilliant black and Navy blue. The initial con-centration of each dye was maintained at 50 mg/l. The optical density of centrifuged sample (supernatant) was taken and finally percent decolorization was calculated by using above mentioned formula (Kalyani et al. 2008).

Analysis of dye degraded productsHPLC, TLC and FTIRThe analysis of dye degraded products was done by thin layer chromatography (TLC), high performance liquid chromatography (HPLC) and Fourier transform infra-red spectroscopy (FTIR). The metabolites were extracted from dye degraded sample (100 ml, 5 days) by mixing it with an equal volume of ethyl acetate. The extracts were then dried on anhydrous Na2SO4 and evaporated in rotary evaporator. The dried powder thus obtained was dissolved in HPLC grade methanol and used for analysis.

TLC was performed to analyze the degraded products on silica gel using mobile phase solvent system n-pro-panol, methanol, ethyl acetate, water and glacial ace-tic acid in the ratio 3:2:2:1:0.5 (Kalyani et al. 2008) and the results were visualized under UV illuminator at 254 and 366 nm. HPLC was performed at (Waters model no 2690) C18 column having symmetry 250 × 4.6 mm using methanol as mobile phase with a flow rate of 1.0 ml/min for 15 min and UV detector at 254 nm (Telke et al. 2009).

FTIR (Bruker, alpha-P) was performed to observe the change in structure of dye before and after decoloriza-tion. The FTIR analysis was done in the mid IRF region of 400–4000/cm. Prior to analysis sample was mixed with pure KBr in the ratio of 5:95 and pellet were then fixed in the holder for analysis (Saratale et al. 2009b).

Gas chromatography mass spectrometryThe collected samples were centrifuged for 10 min at 4 °C and the supernatant collected were extracted thrice with an equal volume of ethyl acetate, dried with Na2SO4 and further concentrated in the rotatory evaporator. GC–MS analysis of metabolites was carried out using gas chro-matograph system (GCMS-QP2010 Ultra, Shimadzu)

equipped with capillary column (DB-5 ms). The sam-ples were diluted 1:100 and injected 1.0 µl sample into GC–MS in split mode with injector temperature of 290 °C. Helium was used as a carrier gas with flow rate of 1.02 ml/min under 54.9 kPa inlet pressure. The column temperature was set to 50 °C and hold for 1 min with sub-sequent increase to 280 °C with ramp rate of 30 °C/min without holding and finally to 310 °C with ramp rate of 15 °C/min by holding it for 8 min. The peaks were identi-fied by comparing with NIST27.LIB mass spectra library of GC–MS database.

Effect of decolorized dye wastewater on microbial growthThe extent of bacterially treated wastewater effect was determined on some useful micro-flora including Bacil-lus megatarium (z-28), Bacillus cereus (T358-2) and Bacillus subtilis (z-66) according to Mali et al. (2000). The plates (L-agar) were swabbed by bacteria; a 2 mm well was made and filled with decolorized dye wastewater. The plate’s incubation was done at 37 °C for 24 h and the zone surrounded the well indicating the toxicity index of degraded dye sample.

ResultsPhysicochemical parameters and isolation of dye degrading bacteriumThe temperature of wastewater samples ranged from 28 to 36 °C while the pH ranged between 8 to10. The color of the samples was black, navy blue and maroon. A total of 14 bacteria were isolated from 3 wastewater samples. On the basis of dye degrading potential, one bacterial isolate designated as 3c, showing 82% decolorization after 4 days of incubation at 37 °C, was selected for further research work (Additional file 1: Table S1). The significant visual difference between control and dye containing wastewa-ter is shown in Fig. 1.

Fig. 1 Change in color of Synazol red 6HBN as compared to control in flask containing culture of bacterial isolate 3c from the dye contaminated wastewater


Bacterial identificationThe morphological and biochemical characteristics are given in Table 1. The optimum growth conditions of the bacterium were 37 °C and pH 7 (Additional file 1: Fig. S1). The organism showed maximum growth (O.D) after 16 h of incubation in LB medium (Additional file 1: Fig. S2). The 16S rRNA gene sequence of bacte-rial isolate showed 95% homology with 16S rRNA gene sequence of Alcaligenes aquatilis. Then this 16S rRNA gene sequence was submitted to GenBank under acces-sion number of KY009932. The bacterial strain has also been deposited at First Fungal Culture Bank of Pakistan and has been assigned accession number FCBP-B-728. The phylogenetic tree (Additional file 1: Fig. S3) which was constructed through MEGA7 shows the homol-ogy of A. aquatilis 3c with members of Alcaligenes sp. The frequency of appearance of completely identical sequence among the strains was indicated by number in the parenthesis.

Optimization of dye decolorizationThe result showed that the A. aquatilis 3c showed maxi-mum decolorization at 37 °C (Fig. 2a), pH 7 (Fig. 2b) and

at static condition (Fig. 2c). The bacterium efficiently uti-lized saw dust and yeast extract as carbon and nitrogen sources, respectively (Fig. 2d, e) to show maximum per-centage of decolorization. Moreover, efficient decoloriza-tion was determined when inoculum percentage was 6 and dye concentration was 10 mg/l (Fig. 2f, g).

Effect of decolorization on growth of bacterial isolateIt was determined that with the passage of time growth rate of A. aquatilis 3c was also increased along with the increase in decolorization percentage (Fig. 3a).

Multiple dyes decolorizationWhen A. aquatilis 3c was grown in the medium contain-ing multiple dyes, it showed 27, 38, 62, 80 and 86% decol-orization after incubation of 1, 2, 3, 4 and 5 days (Fig. 3b).

Dye degraded products analysisTLCTLC analysis showed two bands with Rf value of 0.96 and 0.94 as compared to the control (original dye) band with Rf value of 0.83 when visualized under UV range of 254 and 366 nm (Additional file 1: Fig. S4).

HPLCThe chromatogram of the untreated dye sample showed three detectable peaks at retention time of 1.80, 2.88 and 5.00 min (Fig. 4a) while A. aquatilis 3c treated dye sam-ple showed peaks at retention time of 1.99, 2.30, 2.95 and 3.76 min after 5 days of incubation (Fig. 4b).

FTIRA comparison between the FTIR spectrum of control dye and extracted metabolites was given in Fig. 5. Two specific peaks of wavelength 1612/cm and 1532/cm are shown in FTIR spectrum of control dye which are due to the presence of –N=N– stretching. The peak with wave-length 1395/cm indicates C–O–H bending. The peak with a wavelength 1037/cm is due to C–O stretching. There is a variation in the peaks in the FTIR spectrum of metabolites extracted from bacterial decolorized sample of dye when compared to the control dye spectrum. The absence of peaks with wavelength 1612/cm and 1532/cm indicates the reductive cleavage of azo bond. The peak with wavelength 2850/cm and 2923/cm is due to stretch-ing of alkanes. The peaks with 1755/cm and 1717/cm wavelength indicate the presence of C=O group. A peak of wavelength 1289/cm depicts the presence of carboxyl group. Two peaks with wavelength 1104/cm and 1037/cm show the presence of amines (C–N). The presence of peak with wavelength 769/cm indicates the presence of conjugated C=C.

Table 1 Morphological and biochemical characteristics of A. aquatilis 3c

+ve: positive, –ve: negative

Morphological characteristics A. aquatilis 3c

Shape Spherical

Size 1 mm

Color Pale yellow

Elevation Non elevated

Edges Smooth

Texture Sticky

Gram’s staining Gram negative

Biochemical characteristics

Fermentation

Lactose –ve

Sucrose –ve

Dextrose –ve

H2S production –ve

Nitrate reduction –ve

Indole production –ve

Methl red reaction –ve

Voges Proskauer reaction –ve

Citrate use ± Urease activity –ve

Catalase activity +ve

Oxidase activity +ve


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GC–MS analysisAccording to GC–MS analysis, various end prod-ucts of azo dye were found including pentadecanal, 2-acetyl-3-methylhexahydropyrrolo[1,2-a]pyrazine-1,4-dione, [Z]-hexadec-9-enoic acid, palmitic acid, 3-isobutylhexahydropyrrolo[1,2-a]pyrazine-1,4-dione, 3-benzylhexahydropyrrolo[1,2-a]pyrazine-1,4-dione, bis(6 methylheptyl) phthalate, chlorobenzene, and N′-(3,6-dichloro-2,7-bis(2-(ethyl(methyl)amino) ethoxy-9H-flu-oren-9 ylidene) pivalohydrazide (Additional file 1: Fig. S5).

In the present study, it is confirmed through GC–MS analysis that azo dye enzymatically converted into various end products. These metabolites are used in different pathways for example pyrrolo[1,2-a]pyrazine-1,4-dione derivative can be used as a substrate in amino acid metabolism. The amino acid catabolism can synthe-size 3C compound (pyruvate) which can be converted into acetyl-CoA. The acetyl-CoA further undergoes Krebs cycle to generate reduced molecules (NADH2 and FADH2). Moreover, another end product, phthalate

derivative can be changed into different fatty acids and aldehydes and these molecules can directly/indirectly enter into fatty acid β-oxidation reactions to produce NADH2 and FADH2. These reduced molecules may be used in the process of ATP synthesis (Fig. 6).

Impact of decolorized dye wastewater on microbial growthThe plates were overlaid with microbial growth showing no inhibition zones and indicating that the decolorized dye wastewater is unharmful for the microbial growth (Fig. 7). Likewise, A. aquatilis 3c treated dye wastewater was tested for microbial toxicity assessment and was also found unharmful for microbial growth.

DiscussionNumerous microbes, counting fungi, yeasts, bacteria as well as algae, may decolourize and even cause whole mineralization of several azo colorants underneath sure ecological circumstances. Several assessments are acces-sible on the physiochemical as well as microbiological approaches for azo dyes decolourization (Kaushik and Malik 2010, 2011; Singh et al. 2014; Das and Mishra 2016; Liu et al., 2018; Meerbergen et al. 2018). Aftab et al. (2011) reported a Corynebacterium sp. which decolorizes Reactive black 5 up to 60% and Reactive yellow 15 up to 76% within a period of 4 days when the initial concentra-tion was 100 µg/ml.

Shah (2014) reported three bacteria namely Pseu-domonas putida, P. aeruginosa and B. subtilis capable of degrading multiple dyes i.e., Blue RR, Black B, Red RR, Yellow RR and Navy blue. P. putida showed maximum decolorization of Blue RR (95%), P. aeruginosa of Black B (93%) and Navy blue (70.58%) and B. subtilis of Yellow RR (65%) and Red RR (91%). Joe et al. (2008) reported a bacterium, Clostridium biofermentans, which was able to decolorize dyes Reactive red 3B-A, Reactive black 5, and Reactive yellow 3B-A, by over 90% within a period of 36 h. Likewise, Kaushik and Malik (2011) reported that the high dye removal efficiency (99.97%) and high uptake capacity (97.54 mg/g) of Aspergillus lentulus FJ172995 in 24 h using optimum process variables.

In the current study, A. aquatilis was found to decolor-ize 82% Synazol red 6HBN after incubation of 4 days at 37 °C and pH 7. Saha et al. (2017) reported two A. faeca-lis species namely A. faecalis E5.Cd and A. faecalis Fal.3 which decolorize up to 93% of Blue H/C and Red 3B dye at pH 7 and 94% of Yellow 3R at pH 8 within a period of 96 h. Both the strains decolorize up to 91% of dye at 35 °C, 92% with 50 ppm initial dye concentration and 93% with 20% inoculum size and supplementation of 1% co-substrate respectively. Shah et al. (2013) reported that Bacillus species i.e., B. cereus and B. megatarium hav-ing 95% and 98% dye decolorizing potential respectively

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under optimum conditions. Optimal condition for B. cereus was found to be 37 °C, pH 7, 1% sucrose, 0.25% peptone and 8% inoculum and that for B. megaterium was found to be 37 °C, pH 6, glucose 1%, 0.25% yeast extract and 10% inoculum.

Different bands with Rf value were visualized in Direct orange 16 treated with Micrococcus luteus strain SSN2 and untreated samples when visualized under UV range of 254 nm (Singh et al. 2015). Likewise, a band with Rf value of 0.71 was visualized in the TLC chromatogram of decolorized sample of Direct red by Enterococcus faeca-lis YZ66 as compared to the untreated original dye which had a band with Rf value of 0.97 (Sahasrabudhe et al.

2014). Similar results were obtained against Amarnath mono azo dye decolorized by Acinetobacter calcoaceticus NCIM 2890 (Ghodake et al. 2011). In the present investi-gation, TLC analysis showed two bands with Rf value of 0.96 and 0.94 as compared to the control dye band with Rf value of 0.83 when visualized under UV range of 254 and 366 nm.

Lade et al. (2015) observed three major peaks with retention time 2.521, 3.241 and 3.564 and two minor peaks of retention time 3.123 and 3.910 when extracted metabolites from Provedencia rettgeri decolorized C.I reactive blue 172 were analyzed through HPLC. The HPLC analysis of the control dye results in the presence

Fig. 4 HPLC analysis of reactive Synazol red 6HBN with a mobile phase of profile of methanol at 1.0 mL/min. The column was C-18 (250 × 4.6 mm) a represents dye chromatogram while b represents A. aquatilis 3c degraded dye products extracted after 5 days of incubation at 37 °C


of one major peak with retention time of 2.702 min three minor peaks at retention time of 2.125, 2.801 and 3.394 min. HPLC analysis demonstrated the presence of two major peaks at retention time 2.45 and 2.68 and two minor peaks at retention time 3.50 and 6.50 min when B. halodurans decolorized Acid black-24. The chromato-gram of original dye had only one peak at retention time of 2.81 min (Prasad and Rao 2014). In this study, A. aqua-tilis 3c treated dye sample showed peaks at retention time of 1.99, 2.30, 2.95 and 3.76 min after 16 days of incu-bation (Fig. 4b) while dye sample showed three detect-able peaks at retention time of 1.80, 2.88 and 5.00 min.

In the present study, two specific peaks of wavelength 1612 and 1532/cm are shown in FTIR spectrum of con-trol dye while such peaks are absent in bacterium treated dye sample indicates the reductive cleavage of azo bond. The absence of peak of wavelength 1631/cm in FTIR spectrum of Marinobacter sp. strain HBRA treated

Direct blue-1 sample as compared to the spectrum of control dye clearly demonstrated the degradation of azo bond (Prasad et al. 2013). Shyamala et al. (2014) reported the breakdown of Methyl orange by halotolerant Bacillus sp. as peaks of wavelength 1567/cm and 1424/cm which were present in FTIR spectrum of control dye were absent in spectrum of bacterially treated sample.

Hungerer et al. (1996) reported that the azo-dye degraded compound 1,2-benzene dicarboxylic acid, butyl 8-methylnonyl ester was cleaved to form phthalic acid and palmitic acid. Our findings are similar with other studies that reported that the predominant compounds found in the final dye treated sample were 3-Aminobu-tanoic acid, pyrrolo pyrazine-1, 4-dione and palmitic acid (Harwood and Parales 1996; Walker and Van der Donk 2016; Shanmugam et al. 2017). In the current research, it is confirmed through GC–MS analysis that azo dye

Fig. 5 FTIR spectrum of control dye (Synazol red 6HBN) and A. aquatilis 3c treated sample

Fig. 6 Synazol red 6HBN dye enters into the cell (A. aquatilis 3c) by unknown mechanism. Upon entrance into the cell the dye is enzymatically processed into various end products. Most probably, azo group (–N=N–) is reduced, followed by cleavage reaction and different end products are produced. Secondly, desulfonation and oxidative deamination results in synthesis of pyrrolo[1,2-a]pyrazine-1,4-dione derivative which can be used as substrates in amino acid metabolism. The amino acid catabolism can synthesize pyruvate (3C compound) which can be converted into acetyl-CoA. The acetyl-CoA undergoes Krebs cycle to produce NADH2 and FADH2 (substrates of electron transport chain). Moreover, dye desulfonation, oxidative deamination and carboxylation lead to produce phthalate derivatives, which can be transformed into different fatty acids and aldehydes. The phthalate, fatty acids and aldehydes can directly/indirectly enter into fatty acid oxidation reactions (β-oxidation) to produce acetyl-CoA, NADH2 and FADH2

(See figure on next page.)


enzymatically converted into various end products which may be involved in the synthesis of energy yielding processes.

In the current work, A. aquatilis 3c treated dye waste-water was tested for microbial toxicity assessment and was also found unharmful for microbial growth. Aspergil-lus niger and Nigrospora sp. degraded dye end products containing wastewater was found safe for the growth of soil micro-flora (Ilyas and Rehman 2013). The degrada-tion products of Sudan 1 by B. circulans BWL1061 and S. putrefaciens CN32 showed a decreased toxicity to E. coli BL21 and B. subtilis 168 (Liu et al. 2018). Similarly, Shah et al. (2012) reported the degradation of Reactive Orange 13 through enzyme assay and GC–MS analysis. The final products, naphthalene and 6-[(4-chloro-1,3,5-triazin-2-yl) amino]-2-iminonaphthalen-1(2H)-one were non-toxic revealed by phytotoxicity study.

In conclusion, A. aquatilis 3c has a promising potential to decolorize Synazol red 6HBN (82%) after incubation of 4 days at its optimum growth conditions. The maximum dye decolorization was found under static conditions by using saw dust and yeast extract as carbon and nitrogen source. The bacterium also showed promising poten-tial to decolorize multiple dyes at a rate of 86% in 5 days and dye degradation had positive effect on the growth of organism. This degradation was confirmed through TLC, HPLC, FTIR and GC–MS analysis and the end products are used in various metabolic pathways including ATP synthesis process. It was tested that bacterially decolor-ized dye wastewater is safe and sound for useful micro-bial flora. By using this promising decolorization ability of A. aquatilis 3c wastewater can be ameliorated and used at least for crops irrigation.

Fig. 7 Bacterial decolorized broth used for incubation of a B. subtilis (z-66), b B. megatarium (z-28) and c B. cereus (T358-2) at 37 °C for 48 h. No zone of inhibition was observed in any case


Additional file

Additional file 1: Table S1. Decolorization (%) of bacteria isolated from 3 industrial samples. Figure S1. Growth of A. aquatilis 3c at various (a) temperature and (b) pH after incubation of 16 h. Figure S2. Growth of A. aquatilis 3c in LB medium. Optical density was taken at 600 nm after regular time interval. Figure S3. Phylogenetic tree constructed through MEGA7 to show homology of A. aqualitis 3c with Alcaligenes sp. Figure S4. TLC chromatogram of extracted metabolites of A. aquatilis 3c decolor-ized dye samples visualize UV range of (a) 254 and (b) 366 nm. Figure S5. Chromatograms of extracted metabolites of A. aquatilis 3c decolorized dye sample through GC-MS analysis.

AcknowledgementsNot applicable.

Authors’ contributionsMA performed experiments and analyzed the results. AR designed and supervised the study and helped in manuscript preparation. SZH helped in conducting experiments. ZK and MAN helped in analyzing the data and manuscript preparation. All authors read and approved the final manuscript.

FundingThis is to acknowledge the support of University of the Punjab (Grant no. Env-67), Lahore, Pakistan to accomplish this research work.

Availability of data and materialsPlease contact author for data requests.

Ethics approval and consent to participateNot applicable.

Consent for publicationNot applicable.

Competing interestsThe authors declare that they have no competing interests.

Author details1 Department of Microbiology and Molecular Genetics, University of the Pun-jab, New Campus, Lahore 54590, Pakistan. 2 University, Institute of Medical Laboratory Technology (UIMLT), Faculty of Allied Health Sciences (FAHS), The University of Lahore, Lahore, Pakistan. 3 Department of Microbiology, Govern-ment College University Faisalabad (GCUF), Faisalabad, Pakistan. 4 Department of Chemistry, SBA School of Sciences and Engineering (SBASSE) Lahore, Uni-versity of Management Sciences (LUMS), DHA, Cantt, Lahore 54792, Pakistan.

Received: 10 December 2018 Accepted: 30 April 2019

ReferencesAftab U, Khan MR, Mahfooz M, Ali M, Hassan SH, Rehman A (2011) Decoloriza-

tion and degradation of textile azo dyes by Corynebacterium sp. isolated from industrial effluent. Pakistan J Zool 43:1–8

Alalewi A, Jiang C (2012) Bacterial influence on textile wastewater decoloriza-tion. J Environ Protect 3:889–901

Cappuccino JG, Sherman N (2008) Microbiology: a laboratory manual, vol 9. Pearson/Benjamin Cummings, Boston

Das A, Mishra S (2016) Decolorization of different textile azo dyes using an isolated bacterium Enterococcus durans GM13. Int J Curr Microbiol Appl Sci 5:675–686

de Souza SM, Bonilla KA, de Souza AA (2010) Removal of COD and color from hydrolyzed textile azo dye by combined ozonation and biological treat-ment. J Hazard Mater 179:35–42

Ghodake G, Jadhav U, Tamboli D, Kagalkar A, Govindwar S (2011) Decoloriza-tion of textile dyes and degradation of mono-azo dye Amaranth by Acinetobacter calcoaceticus NCIM 2890. Indian J Microbiol 51:501–508

Harwood CS, Parales RE (1996) The beta-ketoadipate pathway and the biology of self-identity. Annu Rev Microbiol 50:553–590

Hungerer C, Troup B, Römling U, Jahn UD (1996) Regulation of the hemA gene during 5-aminolevulinic acid formation in Pseudomonas aeruginosa. J Bacteriol 177:1435–1443

Ilyas S, Rehman A (2013) Decolorization and detoxification of Synozol red HF-6BN azo dye, by Aspergillus niger and Nigrospora sp. Iranian J Environ Health Sci Eng 10:12

Jadhav JP, Parshetti GK, Kalme SD, Govindwar SP (2007) Decolorization of azo dye methyl red by Saccharomyces cerevisiae MTCC 463. Chemosphere 68:394–400

Joe MH, Lim SY, Kim DH, Lee S (2008) Decolourization of reactive dyes by Clostridium bifermentans SL186 isolated from contaminated soil. World J Microbiol Biotechnol 24:117–121

Kalyani DC, Telke AA, Dhanve RS, Jadhav JP (2008) Ecofriendly biodegrada-tion and detoxification of reactive red 2 textile dye by newly isolated Pseudomonas sp. SUK1. J Hazard Mater 163:735–742

Kaushik P, Malik A (2010) Effect of nutritional conditions on dye removal from textile effluent by Aspergillus lentulus. World J Microbiol Biotechnol 26:1957–1964

Kaushik P, Malik A (2011) Process optimization for efficient dye removal by Aspergillus lentulus FJ172995. J Hazard Mater 185(2–3):837–843

Kumar S, Stecher G, Tamura K (2016) MEGA7: molecular evolutionary genetics analysis version 7.0. Mol Biol Evol 33:1870–1874

Lade H, Govindwar S, Paul D (2015) Low-cost biodegradation and detoxifica-tion of textile azo dye C.I. Reactive blue 172 by Providencia rettgeri strain HSL1. J Chem 2015:1–10. https ://doi.org/10.1155/2015/89410 9

Liu W, You Y, Sun D, Wang S, Zhu J, Liu C (2018) Decolorization and detoxifi-cation of water-insoluble Sudan dye by Shewanella putrefaciens CN32 co-cultured with Bacillus circulans BWL1061. Ecotoxicol Environ Saf 166:11–17

Mali PL, Mahajan MM, Patil DP, Kulkarni MV (2000) Biodecolorization of members of triphenyl methane and azo group of dyes. J Sci Ind Res 59:221–224

Masneuf-Pomarade I, Jeuneb CL, Durrensc P, Lollierb M, Aigled M, Dubourdieu D (2007) Molecular typing of wine yeast strains Saccharomyces bayanus var. uvarum using microsatellite markers. Syst Appl Microbiol 30:75–82

Meerbergen K, Willems KA, Dewil R, Van Impe J, Appels L, Lievens B (2018) Isolation and screening of bacterial isolates from wastewater treatment plants to decolorize azo dyes. J Biosci Bioeng 125(4):448–456

O’Neil C, Hawkes FR, Hawkes DL, Lourenco ND, Pinheiro HM, Delee W (1999) Colour in textile effluents-sources, measurement, discharge consents and simulation: a review. J Chem Technol Biotechnol 74:1009–1018

Pandey A, Singh P, Iyengar L (2007) Bacterial decolorization and degradation of azo dyes. Int Biodeterior Biodegradation 59:73–84

Parshetti G, Kalme S, Saratale G, Govindwar S (2006) Biodegradation of mala-chite green by Kocuria rosea MTCC 1532. Acta Chim Slov 53:492–498

Prasad ASA, Rao KVB (2014) Aerobic biodegradation of azo dye Acid Black-24 by Bacillus halodurans. J Environ Biol 35:549–554

Prasad ASA, Satyanarayana VSV, Rao KVB (2013) Biotransformation of direct blue 1 by a moderately halophilic bacterium Marinobacter sp. strain HBRA and toxicity assessment of degraded metabolites. J Hazard Mater 262:674–684

Rehman A, Ali A, Muneer B, Shakoori AR (2007) Resistance and biosorption of mercury by bacteria isolated from industrial effluents. Pakistan J Zool 39:137–146

Saha AK, Sultana N, Mohanta MK, Mandald A, Haquee M (2017) Identification and characterization of azo dye decolourizing bacterial strains, Alcaligenes faecalis E5.Cd and A. faecalis Fal. 3 isolated from textile effluents. Am Sci Res J Eng Technol Sci 3:163–175

Sahasrabudhe MM, Saratale RG, Saratale GD, Pathade GR (2014) Decoloriza-tion and detoxification of sulfonated toxic diazo dye C.I. Direct red 81 by Enterococcus faecalis YZ 66. J Environ Health Sci Eng 12:151

Saratale RG, Saratale GD, Kalyani DC, Chang JS, Govindwar SP (2009a) Enhanced decolorization and biodegradation of textile azo dye Scarlet R by using developed microbial consortium-GR. Bioresour Technol 100:2493–2500

Saratale RG, Saratale GD, Chang JS, Govindwar SP (2009b) Decolorization and biodegradation of textile dye navy blue HER by Trichosporon beigelii NCIM-3326. J Hazard Mater 166:1421–1428

https://doi.org/10.1186/s13568-019-0788-3

https://doi.org/10.1155/2015/894109


Shah MP (2014) Biodegradation of azo dyes by three isolated bacterial strains: an environmental bioremedial approach. J Microbial Biochem Technol S 3:007

Shah PD, Dave SR, Rao MS (2012) Enzymatic degradation of textile dye reactive orange 13 by newly isolated bacterial strain Alcaligenes faecalis PMS-1. Int Biodeterior Biodegradation 69:41–50

Shah MP, Patel KA, Nair SS, Darji AM (2013) Potential effect of two Bacillus sp. on decolorization of azo dye. J Bioremed Biodegrad 4:199

Shanmugam BK, Easwaran SN, Lakra R, Deepa PR, Mahadevan S (2017) Meta-bolic pathway and role of individual species in the bacterial consortium for biodegradation of azo dye: a biocalorimetric investigation. Chemos-phere 188:81–89

Shyamala A, Hemapriya J, Vadakkan K, Vijayanand S (2014) Bioremediation of Methyl orange, a synthetic textile azo dye by a halotolerant bacterial strain. Int J Curr Res Acad Rev 2:373–381

Singh RP, Singh PK, Singh RL (2014) Bacterial decolorization of textile azo dye acid orange by Staphylococcus hominis RMLRT03. Toxicol Int 21:160–166

Singh S, Chatterji S, Nandini PT, Prasad ASA, Rao KVB (2015) Biodegradation of azo dye Direct Orange 16 by Micrococcus luteus strain SSN2. Int J Environ Sci Technol 12:2161–2168

Solis M, Solis A, Perez HI, Manjarrez N, Flores M (2012) Microbial decolouration of azo dyes: a review. Process Biochem 47:1723–1748

Telke AA, Kalyani D, Dawkar V, Govindwar S (2009) Influence of organic and inorganic compounds on oxidoreductive decolourization of sulfonated azo dye C.I. Reactive Orange 16. J Hazard Mater 172:298–309

Turner S, Pryer KM, Miao VPW, Palmer JD (1999) Investigating deep phyloge-netic relationships among cyanobacteria and plastids by small subunit rRNA sequence analysis. J Eukar Microbiol 46:327–338

Verma AK, Dash RR, Bhunia P (2012) A review on chemical coagulation floc-culation technologies for removal of colour from textile wastewaters. J Environ Manage 93:154–168

Walker MC, Van der Donk WA (2016) The many roles of glutamate in metabo-lism. J Ind Microbiol Biotechnol 43:419–430

Publisher’s NoteSpringer Nature remains neutral with regard to jurisdictional claims in pub-lished maps and institutional affiliations.

REVIEW

Microbial use for azo dye degradation—a strategyfor dye bioremediation

Mehvish Ajaz1 & Sana Shakeel1 & Abdul Rehman1

Received: 27 April 2019 /Revised: 8 October 2019 /Accepted: 17 October 2019# Springer Nature Switzerland AG 2019

AbstractAzo dyes are aromatic compounds with one to many –N=N– groups as well as the leading class of synthetic dyes utilised incommercial solicitations. Azo dyes, released in the environment through textile effluents, have hazardous effects on the aquatic aswell as human life. Their persistence and discharge into the environment are becoming a global concern; thus, the remediation ofthese contaminants has acquired great attention. The current review comprehensively discusses some of the main aspects ofbiodegradation of azo dyes. A variety of physicochemical approaches has already been utilised for treatment of textile effluentscounting filtration, coagulation and chemical flocculation. Though these conventional techniques are effective, yet they are lavishand also comprise formation of concentrated sludge that makes a secondary disposal problem. In this regard, microbial usage isan effective, economical, bio-friendly and ecologically benign approach.

Keywords Azo dyes .Microorganisms . Decolourisation . Degradation . Bioremediation

Introduction

For centuries, mankind had been using dyes and byNeanderthal man the very first proof of dye use came about180,000 years back (Christie 2007). Without exception, thedyes utilised, until the late nineteenth century, were roughlyobtained from the raw materials such as leaves, roots, vegetalextracts, branches, berries, blossoms of numerous plants, mol-luscs and insects, and were usually have limited range of col-ours (Gupta 2009).

Yet, pale purple dye in 1856 as a world’s first commercialsynthetic colourant was exposed via W.H. Perkin unexpected-ly (Hunger 2007). These dyes could be delineated as colouredconstituents giving an everlasting colour to fibres whensmeared plus retain itself even on exposure to light, water,perspiration and several chemicals, including oxidising prox-ies in addition to by the bout of microorganisms (Rai et al.2005). Synthetic colourants have largely substituted the natu-ral dyes, due to the huge range of unique colours as well astheir cost effectiveness (Gabriel and Hong 2008).

In 1994, it was estimated that from the entire worldwidefabrication of colourants, that was approximately 1,000,000tons, more than 50% were azo dyes (Olligaard et al. 1999;Stolz 2001). Azo dyes, named on the basis of their chemicalconfiguration, characteristically contain single to countless (–N=N–) azo groups (Chang and Kuo 2000). Azo complexeshave been established as one of the major and most versatileassortments of man-made colourants that are widely used innumerous trades such as foodstuff, paper printing, textile andcosmetic manufacturing (Pandey et al. 2007). Latest interpre-tations conducted via Rajaguru et al. (2002) and de AragaoUmbuzeiro et al. (2005) report the azo colourants causing anincrease in mutagenic bustle of surface and ground waterwhen filthy by textile effluvium whose discharge into surfacewater poses serious harm to the soil prolificacy, natural chat-tels and aquatic creatures, and also upsets legitimacy of eco-system (Mester and Tien 2000; Puvaneswari et al. 2006; Zhaoet al. 2011) by altering pH, increasing the BOD and COD andnegatively affect quality of water. Approximately 280,000tons of textile colourants are given out in the atmosphereworldwide per year (Jin et al. 2007), most of it get mixed intothe aquatic environment. Accordingly, elimination of dyesfrom the textile effluvium is becoming a serious problem.

Extensive work had been done on utilising physical andchemical techniques to eradicate colourants from sewage(Swaminathan et al. 2003; Behnajady et al. 2004; Golab

* Abdul [email protected]

1 Department of Microbiology and Molecular Genetics, University ofthe Punjab, New Campus, Lahore 54590, Pakistan

International Microbiologyhttps://doi.org/10.1007/s10123-019-00103-2

http://crossmark.crossref.org/dialog/?doi=10.1007/s10123-019-00103-2&domain=pdf

mailto:[email protected]

et al. 2005; Chen et al. 2010; Malik et al. 2017). However,these approaches are impractical and ineffective as they arenot only complex processes but also demand large quantitiesof chemicals and energy. These are incapable of fully elimi-nate either the recalcitrant azo dyes or both dyes with theirorganic metabolites, and the use of chemicals has led to pro-duction of huge quantities of sludge which cause secondarycontamination problems (Forgacs et al. 2004; Zhang et al.2004). In contrast, biodegradation is a cheap, eco-friendlyand innovative approach that could clean the water systemby eliminating dyes through the processes such as desalina-tion, adsorption, reduction precipitation, reverse osmosis,photocatalysis and membrane filtration (Verma andMadamwar 2003; Rai et al. 2005; Rafatullah et al.2010; Mishra and Maiti 2018). In the present paper, we woulddiscuss some of the biodegradation approaches to treat thetextile effluents comprising azo colourants.

Bioremediation strategies

Bioremediation approach is based on the use of native micro-organisms’ degradation ability. As xenobiotic azo dyes arecomplex and their degradation is not easy (Mohana et al.2008), azo dyes are degraded by both aerobic and anaerobicpathway; aerobic degradation is greatly favoured; however,anaerobic degradation yields by-products like aromaticamines that are not only hazardous but also have cancinogenicpotential (Senan and Abraham 2004).

In physiochemical methods, the extensively used floccula-tion/coagulation, adsorption and membrane filtrationmethods, all yield huge amounts of sludge, which further re-quire safe dumping (Maier et al. 2004; Ramya et al. 2007).However, the biological approaches are generally regarded asenvironmental friendly, as they can mineralise organic con-taminants without producing secondary toxic sludge(Jirasripongpun et al. 2007). This is usually observed thatcolourants (dyes, pigments) repel biodegradation inorthodox-activated sludge treatment process (Stolz 2001).Numerous microbes, such as fungi, yeasts, bacteria and algae,could not only decolourise several azo dyes but also complete-ly degrade them under certain ecological circumstances.Several investigations have been reported on the physiochem-ical as well as microbiological approaches for azo dyedecolourisation (McMullan et al. 2001; Stolz 2001; Rai et al.2005; Van der Zee and Villaverde 2005; Mani et al. 2019).

Fungal degradation of azo dyes

Being an economical, eco-friendly, less sludge producing pro-cess, fungal treatment is very advantageous process used byresearchers to find a new way for remediation. Contrary to bac-teria, fungi robustly degrade multifaceted organic complexes by

utilising methods such as production of extracellular ligninolyticenzymes comprising manganese peroxidase, laccase and ligninperoxidase; thus, scientists are more interested in using fungi inthe current scenario (He et al. 2018; Guo et al. 2019; Mani et al.2019; Zahran et al. 2019). For the dye decolourisation, the po-tential of ligninolytic fungi is extensively explored (Bumpus2004). Numerous investigations validate the white rot fungi po-tential for degrading and decolourising contaminants via extra-cellular ligninolytic enzymes (Champagne and Ramsay 2010;Kunjadia et al. 2012). In relation to azo dye degradation anddecolourisation, white rot fungus Phanerochaete chrysosporiumis extensively studied; however, some others too have gainedsignificant attention, such as Aspergillus ochraceus, Trametesversicolor, species of Pleurotus and Phlebia (Pointing andVrijmoed 2000; Saratale et al. 2006; Humnabadkar et al. 2008;Sen et al. 2016; Kunjadia et al. 2016). Gahlout et al. (2013)testified that Ganoderma cupreum AG-1 has the potential fordecolourising numerous azo colourants. Though, solicitation ofwhite rot fungus to eliminate dye from textile wastewater has afew disadvantages, for example, the lengthy growth cycle andthe requisition for N2 restraining situations. However, white rotfungus is naturally absent in wastewaters; therefore, enzyme fab-rication for the degradation of dyes is an independent phenome-non (Robinson et al. 2001). Furthermore, the lengthy hydraulicretention period mandatory to completely decolourise likewiseconfines the function of the fungal decolourisation scheme(Banat et al. 1996; Chang et al. 2004), as well as themaintenanceof fungus, in bioreactors, is a problem of consideration as well(Stolz 2001).

Azo dye degradation via yeasts

To ascertain the decolourising potential of yeasts, minor workhas been done, especially focussing on the bio-sorption in-stead. Few ascomycetes yeast species, for instance, Candidatropicalis, C. zeylanoides Debaryomyces polymorphus (Yanget al. 2003) and Issatchenkia occidentalis (Ramalho et al.2004), are utilised for assessing the enzymatic biodegradationaccompanied with the decolourisation of various azocolourants. Lately, S. cerevisiae MTCC-463 had testified fordecolourising Methyl Red as well as Malachite Green (Jadhavand Govindwar 2006; Jadhav et al. 2007). GalactomycesgeotrichumMTCC 1360 may degrade azo, triphenyl methaneand the reactive highly used textile colourants (Jadhav et al.2008a). Via Trichosporon beigelii NCIM-3326, Navy BlueHER decolourisation could also be carried out with the enzy-matic mechanism (Saratale et al. 2009c). Lucas et al. (2007)reported that presence of Candida oleophila decolourised theazo dye Reactive black 5 dye in an assay containing Fenton’sreagent. More recently, a thorough study revealed thatCyberlindnera samutprakarnensis could decolourise AcidRed B (ARB) within 18 h with the efficiency of 97% underoptimal conditions (Song et al. 2018).

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Algal remediation of azo dyes

Photosynthetic organisms, like algae and cyanobacteria, areubiquitous in nature and also have been studied for theirdecolourisation potential of wastewater and it has been pro-posed that algae are proficient in degradation of azo dyes byutilising its azoreductases (Omar 2008; Vijayaraghavan andYun 2008). Azo dyes can be removed by algae through threediverse mechanisms of assimilative chromophores usage: (1)for the creation of algal biomass, (2) H2O as well as CO2

alteration from coloured to non-coloured molecules, (3) chro-mophores adsorption on algal biomass (Acuner and Dilek2004; Mahalakshmi et al. 2015). The decolourisation to bio-sorption trailed through bio-coagulation as well as bioconver-sion by utilising algae is characterised byMohan et al. (2002).More than 30 azo complexes and basic aromatic amines havebeen reported to get degraded completely via Chlorellavulgaris, C. pyrenoidosa and Oscillateria sp. (Yan and Pan2004). Therefore, algae might play a vital part for eliminationof azo colourants as well as aromatic amines from steadypools. Furthermore, present bio-sorption procedure is accept-ed as a profitable, active method for possible utilisation indecolourising the effluents, substituting the much costlier sup-plies (Banat et al. 1996; Daeshwar et al. 2007).

Azo dye degradation utilising plants

Patil et al. (2009) stated that azo dye decolourisation via plantsis an emerging practice ensuring the effective and cheap wayto remediate the soils and groundwater which were pollutedwith organic contaminants and heavy metals. The key advan-tage associated with the remediation by plants is that beingautotrophic, plants require only petite input of nutrients andare generally accepted by the community due to its ecologicalpracticability (Ghodake et al. 2009a; Kagalkar et al. 2009).Significant decline (72–77%) in colour intensity was observedwhen the wetlands were surrounded by coco yam vegetation(Mbuligwe 2005). Besides, the decolourisation potential ofvarious plant species such as Sorghum vulgare, Brassicajuncea and Phaseolus mungo was assessed against the reac-tive group of azo dyes from the industrial effluents. S. vulgare,B. juncea and P. mungo could effectively decolourise indus-trial effluent with the efficiency of 57%, 79% and 53%, cor-respondingly (Ghodake et al. 2009a) (Table 1). Likewise, anherb Blumea malcommi is used for the degradation ofReactive Red 5B textile dye (Kagalkar et al. 2009). ReactiveRed 198 was found to be degrade via the hirsute root culture ofTagetes patula (Marigold) (Patil et al. 2009). Though, plantsplay a major role in the azo dye bioremediation; however,there are problems which limit their use in this scenario suchas the level of contaminants tolerated by plant, the bio-available portion of pollutants, evapotranspiration of unstableorganic contaminants and necessitating great zones for

grafting the approach (Ghodake et al. 2009b). The two speciesof plants, Sesbania cannabina Pers and Medicago sativa,have been proved to tolerate the poisonousness and salinityof the azo dye wastewater as well as involve in bio-decolourisation of the wastewater (Zhou and Xiang 2013).

Enzymes in azo dye degradation

Azo reductase facilitates the azo cleavage in the presence ofreducing agents such as FADH and NADH. Azo reductaseshave been recognised in numerous anaerobic bacteria and inthe microflora of human intestine. The earliest azo reductasesrecognised from the anaerobic microbes were oxygen sensi-tive; however, currently designated as E. gallinarum strain,obtained from the textile industry wastewater treatment plant,is able to decolourise DB38 azo dye via azoreductase enzymewhich acts optimally underneath aerobic circumstances(Bafana et al. (2008); Fu et al. 2019). Similarly, azoreductasegene from Bacillus latrosporus RRK1 produces azoreductasecapable of decolourising numerous azo colourants underneathaerobic circumstances, when replicated and articulated inE. coli, could decolourise Remazol Red only when a specificamount of dissolved oxygen (0.8 mg/L) had been provided(Sandhya et al. 2008). Duran et al. (2002) reported that fungifrequently destroy colourants via exoenzymes similar to theperoxidases. Kalsoom et al. (2013) reported that SoybeanPeroxidase may proficiently degrade a diazo dye, TrypanBlue, and it is also an eco-friendlier approach.

The Lacasses, catalysing the reduction of 4e− of O2 to H2O,are mostly obtained via fungus causing lignin degradation forinstance T. versicolor, T.villoa, Cladospora cladosporioidesand Fusarium soloni (Abedin 2008; Pandi et al. 2019).

Bacterial degradation and decolourisation of azo dyes

Usually, azo dye decolourisation happens under tradition-al aerobic, anaerobic also facultative anaerobic circum-stances via diverse bacteria. In eliminating colour, anaer-obic microbial approach could be considered as success-ful, chiefly via azo reductases slicing −N=N− giving re-sultant amines, which are not only lethal nonetheless like-wise oncogenic as well as mutagenic, and above repelfurther decolourisation under anaerobic environment(O’neill et al. 2000; Van der Zee et al. 2001; Gottliebet al. 2003; Pinheiro et al. 2004; Van der Zee andVillaverde 2005; Sarkar et al. 2017; Fu et al. 2019; deSouza et al. 2019). Under anaerobic respiratory oxidationof carbon sources, the azo dyes act as a terminal e− ac-ceptor (Carliell et al. 1995; Ryan et al. 2010; de Souzaet al. 2019). Contrastly, activated sludge processes beingaerobic in nature, such bio-approaches, proved to be inef-fective for the treatment of textile wastewaters as theycause minute or no colour elimination from azo dye

Int Microbiol

(Chern and Chien 2003; Brik et al. 2006; Singh et al.2007; Roy et al. 2018). Bacteria having capacity of aero-bic decolourisation of colourants, especially sulfonatedazo dye, are hard to sequester (McMullan et al. 2001;Pearce et al. 2003‚ Lin et al. 2010). Though, pure inaddition to mixed bacterial culture have been exposed todyes capable of aerobic degradation and cause detoxifica-t ion of aromat ic amines formed via anaerobicdecolourisation of azo colourants (Khalid et al. 2009;Mishra and Maiti 2018).

In one of the studies from this laboratory (Ajaz et al. 2019),via GC-MS examination, it has been long established that en-zymatically azo colurant is transformed into numerous endproducts by using Staphylococcus lentus 1 M. These metabo-lites are used in different pathways; for example, during aminoacid metabolism, 4-guanidinobutyric acid is utilised as a sub-strate. The amino acid catabolism may produce pyruvate that istransformed into acetyl coenzyme Awhich further go throughTCA cycle to generate reduced molecules. Moreover, anotherend product, phthalate derivative, can be changed into differentfatty acids as well as aldehydes and these molecules may movein a direct or indirect way into fatty acid β-oxidation reactionsyielding NADH2 also FADH2. These reduced molecules maybe used in the process of ATP synthesis (Fig. 1).

Because of recalcitrant nature of azo dyes to highly aerobiccircumstances as well as the production and accumulation ofecologically harmful amines under anaerobic circumstancesfor the whole mineralisation of azo dyes, successiveanaerobic–aerobic biological approach has been establishedas the utmost effective and economical methods.

Anaerobic decolourisation via reductive cleavage of azodye’s linkages consequently produces colourless, hazardousaromatic amines which could be degraded in the aerobic stage(Popli and Patel 2015). Furthermore, the high salinity and pHbeing the attributes of textile dye wastewaters were exposed toconstraining azo colourant decolourisation via wastewater mi-crobial populations representing that tolerance to such hostilecircumstances is likely to be a vital limit for the effective bio-transformation of azo colourant left-over watercourses (Manuand Chauhari 2003; Şen and Demirer 2003; Asad et al. 2007).Thus, it is vital to isolate appropriate microbial strains and also

determine the factors indispensable for the efficient bio-treatment of azo colourants (Franciscon et al. 2012).

Utilising co-culture as well as mixed bacterial cultures

Azo dye decolourisation with pure cultures of bacterial strainsis effective; however, distinct bacterial isolates frequentlycould not be able to decolourise azo dyes entirely and alsoare responsible for the production of carcinogenic aromaticamines as the transitional products, which needs further deg-radation (Joshi et al. 2008). Therefore, the treatment schemesusually consist of mixed microbial communities which attainan advanced level of mineralisation and biodegradation ma-jorly because of the synergistic metabolic actions of the mi-crobial communities, also these are significantly beneficial ascompared with the use of pure cultures (Khehra et al. 2005;Chen and Chang 2007; Saratale et al. 2010).

To aid the decomposition process, different isolates mightsecrete the colourant molecule, in a microbial consortium, atdiverse locations or might utilise metabolites made through theco-existing isolates (Chang et al. 2004; Forgacs et al. 2004;Jadhav et al. 2008b; Saratale et al. 2009b; Vikrant et al. 2018).Under aerobic conditions, at ambient temperature (28 ± 2 °C)and basic pH (9–10.5), when the salinity ranges between 0.9 and3.68 g/L, bacterial consortium containing two isolates (BF1,BF2) and a Pseudomonas putida (MTCC1194) strain effectivelydegrade a combination of textile azo dyes (Senan and Abraham2004). Das and Mishra (2017) reported that the bacterial consor-tium of Bacillus pumilus HKG212 and Zobellella taiwanensisAT1–3 could eliminate the reactive green-19 dye underoptimised static conditions; however, after the addition of yeastextract, the removal efficacy of consortium was reached up to97%within 24 h of incubation when the initial conc. of colourantwas 100 mg/L. Moreover, Waghmode et al. (2011) determinedthat the bacterial consortium consisting of Brevibacilluslaterosporus (NCIM2298) and GG-BL Galactomycesgeotrichum (MTCC 1360) could decolourise the golden yellowHER under aerobic-microaerophilic conditions. The authors’findings revealed that the rigorous action of isolatesmay substan-tially diminish the COD (84%), TOC (63%) and also colour(100%) within 24 h from aq. solution loaded with dye. Inside

Table 1 Dye decolourisation potential (%) in bacterial and fungal strains

Sr. no. Organism Dye used Dye decolourisationpotential (%)

Reference

1 P. aeruginosa Reactive Red BS C.I. 111 91 Sheth and Dave (2009)

2 Aspergillus niger Synazol red 6HBN 88 Ilyas and Rehman (2013)

3 Nigrospora sp. Synazol red 6HBN 96 Ilyas and Rehman (2013)

4 A. faecalis E5.Cd Red 3B 93 Saha et al. (2017)

5 Alishewanella sp. CBL-2 Sumifex Tourqi blue 83 Ajaz et al. (2018)

6 Alcaligenes aquatilis 3C Synazol red 6HBN 82 Ajaz et al. (2019)

Int Microbiol

the microbial consortium, the initial breakdown of azo dyes re-sulted in the aromatic amines, which further get broken via cor-responding entities present; thus, the practice becomes extra ef-ficient as well as effective (Moosvi et al. 2005, 2007; Sarataleet al. 2009a, b; Tony et al. 2009a, b; Verma et al. 2019).

Using pure bacterial culture

The mixed cultures merely offer an ordinary macroscopic ap-proach to what is going on within the system, as well as the

outcomes are not simply represented. In the 1970s, laboursisolated pure bacterial cultures of Aeromonas hydrophila,Bacillus cereus and Bacillus subtilis able to degrade azo dyes(Wuhrmann et al. 1980). Lately, extensive studies on the dyedegradation have been investigated by utilising single bacte-rial cultures for instance Proteus mirabilis and Pseudomonasluteola which showed very promising outcomes underneathanoxic situations (Chang et al. 2001; Yu et al. 2001; Kalyaniet al. 2008). Navitan Fast Blue S5R, a marketable colourant,has been decolourised by Pseudomonas aeruginosa when

Fig. 1 Synazol Red HF 6BN dye enters into the cell (Staphylococcuslentus 1 M) by unknown mechanism. Upon entrance into the cell, thedye is enzymatically processed into various end products. Most probably,azo group (–N=N–) is reduced, followed by cleavage reaction anddifferent end products are produced. Secondly, desulfonation andoxidative deamination eventually result in synthesis of 4-guanidinobutanoic acid derivative which can be used as substrates inamino acid metabolism. The amino acid catabolism can synthesise

pyruvate (3C compound) which can be converted into acetyl-CoA. Theacetyl-CoA undergoes Krebs cycle to produce NADH2 and FADH2 (sub-strates of electron transport chain). Moreover, dye desulfonation, oxida-tive deamination and carboxylation lead to produce phthalate derivatives,which can be transformed into different fatty acids and aldehydes. Thephthalate, fatty acids and aldehydes can directly/indirectly enter into fattyacid oxidation reactions (β-oxidation) to produce acetyl-CoA, NADH2

and FADH2

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supplementedwith glucose under aerobic circumstances and itmay also degrade many different azo colourants as well(Nachiyar and Rajkumar 2005).

In 2008, from dye-contaminated soil, Kalyani et al. seclud-ed a facultative anaerobic bacterium Pseudomonas sp. SUK1for biodegradation studies of reactive red-BLI dye under staticanaerobic circumstance. Investigators found the bacterial iso-late displaying decolourisation efficacy up to 99.28% within1-h incubation when the initial concentration was l50 mg/Ldye, where the bacterial strain degrades poisonous dye intoharmless products. The decolourisation of reactive black-5dye utilising Bacillus sp. YZU1, isolated by dye-polluted soil,has been studied byWang et al. (2013) in static condition. Theinvestigators found that azoreductase enzyme formed by thestrain catalysed the degradation (95% degradation, within 120h, of 100 mg colourant/L) of dye. Enterococcus faecalis YZ66 strain showed the capability of degrading manyindustrialised dyes amongst which, it displayed whole degra-dation of Direct Red 81 diazo dye within incubation period of1.5 h under static anoxic circumstances (Sahasrabudhe et al.2014). For the decolourisation of reactive sulfonated azocolourants, Brevibacterium sp. may be utilised in a sequentialstatic/aerobic procedure which was sequestered from one tex-tile dye wastewater treatment plant (Franciscon et al. 2012).Also, in a study, decolourisation of the azo dye Acid Orange51 from Trametes trogii using rudimentary laccase has beenproved (Daâssi et al. 2013).

In the recent past for the decolourisation of the combinationof reactive dyes, several bacterial species were examined (Dasand Mishra 2017; Perumal et al. 2017; Meerbergen et al.2018). Lucious et al. (2014) have treated the liquid wastesupplemented with the mixture of dyes acid blue and acidred with the Pseudomonas aeruginosa under aerobic condi-tions. Scientists stated that bacteria exhibiting decolourisationability maximally remove 90% of dye, which seemed im-proved as compared with the reference strain, i.e. Bacillussubtilis. Under anaerobic–aerobic system, the reductivedecolourisation of azo colourants (acid red-88, direct red-88as well as reactive black-5) via Shewanella sp. IFN4 fromsynthetic wastewater with efficiency of 90% has been reported(Imran et al. 2014; de Souza et al. 2019).

Using whole bacterial cells

Specifically, the capability of whole bacterial cells has beenbroadly examined for metabolising azo dyes. Underneath aer-obic situations, azo dye could not freely metabolise (Robinsonet al. 2001). Though, in anaerobic situations, several bacteriacould shrink the extremely e− loving azo bond in the dyemolecule, seemingly via the action of short specificity cyto-plasmic azo reductases, to yield colourless aromatic amines.Such amines are impervious for more degradation; thus, an-aerobic mineralisation may be mutagenic otherwise lethal for

the faunas. Luckily, after the removal of xenobiotic azo ele-ment of the colourant molecule, the resulting amino com-plexes can be easily aerobically biodegrade. Amines may un-dergo mineralisation under aerobic circumstances, hiring asuccessive anaerobic–aerobic scheme to treat the wastewater,via a hydroxylation trail comprising a ring opening system(Lourenco et al. 2000). The utilisation of whole cells incoloured wastewater (containing dyes) is beneficial relativelyto the isolated enzymes, since expenses allied with purifica-tion of enzymes are annulled as well as the cells may too offerdefence to the enzymes from the severe environment process(Pearce et al. 2003; Sandesh et al. 2019).

In situ and ex situ bioremediation

Both strategies are being used for the treatment of industrialwastewater containing toxic pollutants including azo dyes. Exsitu bioremediation involves the removal of contaminated soilfrom ground and land farming is one of the techniques in whichcontaminated soil is excavated. It then spread over a preparedbed and periodically tilled until pollutants are degraded.Usually, this practice is limited to the treatment of superficial10–35 cm in soil (Shah 2014). Wang et al. (2018) reported thatthe in situ generated bio-Pd by Klebsiella oxytoca strain isefficient for azo dye reduction without complex preparationprocesses. This is of great importance for the removal and sub-sequent safe disposal of toxic metal ions and azo dyes.

Factors influencing bacterial decolourisation

During biotic treatment procedures, various physicochemicalworking limits, for instance the agitation level, temperature,oxygen, pH, concentration and structure of dye, electron do-nor and redox mediator, provision of various nitrogen andcarbon sources, sprightly affect the bacterial decolourisationof azo colourants (Krishnan et al. 2017). Therefore, for mak-ing the procedure super effective, earlier determination of theoutcome of individual feature on the bacterial decolourisationof azo colourants is crucial.

Usually to degrade dyes biologically the inert state isdesirable, where the low oxygen level is retained in thereaction medium (Bouraie and Din 2016). In 2003,Pearce et al . c la imed that for achieving worthydecolourisation, the optimal pH of 6.0–10.0 is requisitewhile 25–35 °C temperature range could stay appropriateaimed at bacterial cell growth within the decolourisationmedium. Though, azoreductase enzyme formed via bacte-ria has been seen to be thermostable at 60 °C, being ener-getic for short interval (Misal et al. 2011). For bio-treatment of the dye-polluted wastewater, accessibility ofa particular electron donor is essential (Al-Amrani et al.

Int Microbiol

2014). Dyes having low molecular weight and unpreten-tious construction are seen to degrade simply, althoughdecolourisation rate declines with dyes having e− with-drawing substitution groups at the p-position in the phenylring (Pandey et al. 2007).

Mechanisms of microbial dye degradation

Azo complexes stand vulnerable to aerobic as well as anaerobicbiodegradation (Khehra et al. 2005). Generally, remediation ofazo colourants by microbes encompasses breakdown of the azobonds by azoreductase enzyme (EC 1.7.1.6) in anaerobic condi-tions, also involving transference of four electrons. Then at theazo linkage there are two stages involved, each stage involvingtransfer of two electrons to the azo dyes (acting as an ultimateelectron acceptor), causing dye degradation and formingcolourless mixtures (Chang and Kuo 2000). The degradation ofresultant transitional metabolites, including aromatic amines, iscarried out anaerobically/aerobically (Bromley-Challenor et al.2000; Chang et al. 2004; Pandey et al. 2007) (Fig. 2).

Azoreductase is a reducing enzyme that degrades azo dyethrough reductive cleavage process into colourless amines. Itneeds low molecular weight reducing equivalent like FADHorNADH as the electron donor in the form of a redox reaction.Azoreductase is categorised into three types on the basis ofcoenzyme use: using NADH only, using NADPH only orusing both. This enzyme can be either cytoplasmic or mem-brane bound. It has gained importance due to its use in waste-water treatment containing azo dyes (Sarkar et al. 2017).

Few researchers proposed that the occurrence of O2 custom-arily impedes the action of azo bond reduction, as aerobic res-piration might tower over the NADH use, consequently ham-pering the transference of electron towards azo bond from

NADH (Chang et al. 2001). Otherwise, decolourisation maybe attributed to unspecified extracellular responses happeningamid reduced composites produced via the anoxygenic fuel(Van der Zee et al. 2001). Currently, only in anoxygenic set-tings, the mechanism of azo dye decolourisation is well ac-knowledged (Singh et al. 2015). Considerable investigationscovering the anaerobic decolourisation of colours (mainly azodyes) were piloted utilising monocultures. The infusion of azodyes in anaerobic conditions in the microbial cells from biolog-ical membrane is stated to be the major rate-limiting feature ofdecolourisation (Kodam et al. 2005). In aerobic environments,amalgamation of O2 in the aromatic ring of carbon-based com-plexes from O2 is catalysed by mono- as well as di-oxygenaseenzymes earlier to ring fission (Sarayu and Sandhya 2010). In2010, Lin et al. stated the few oxygenic bacteria reducing azocomplexes with oxygen catalysed azoreductases yielding aro-matic amines. The aerobic azo reductases were reported to uti-lise equally NADH as well as NAD(P)H as cofactors causingreductive cleavage of the bacterial carboxylated growth sub-strates and the sulfonated structural equivalents too (Kecket al. 1997; Nachiyar and Rajkumar 2005). Furthermore, fewbacteria are capable of growing on azo complexes by way ofsingle carbon source. Such bacteria after reductively cleavingazo bonds use amines as carbon source as well as energy togrow. Pigmentiphaga kullae K24 and Xenophilus azovoransKF 46 are instances of bacterial strains having this attribute,which may aerobically breed upon carboxy-orange II as wellas I, correspondingly (McMullan et al. 2001). Merely, a smallnumber of bacteria having specific enzymes capable of reduc-ing azo dye are seen for breaking azo dyes underneath entirelyaerobic circumstances (Zimmermann et al. 1984; Nachiyar andRajkumar 2003). In 2003, Blümel and Stolz (2003) also repli-cated as well as described the triplet code, from Pagmentiphagakullae K24, of the aerobic azo reductase.

Fig. 2 Suggested mechanism ofutilising whole bacterial cells forreduction of azo dyes with somemodification (Pearce et al. 2003)

Int Microbiol

Conclusions

Dyes discharged into the environment have negative effects tothe biosphere and biota. The chemical treatment of dyes is notrecommended by many researchers due to the generation ofsecondary pollutants. Microbial degradation of azo dyes hasnoteworthy prospective to discourse this problem because oftheir cheap and eco-friendlier nature. Amongst microbes, bac-teria are extensively used to degrade dyes because of theirrapidly growing ability and extraordinary hydraulic retentionperiod; hence, they may be effective in treatment high-forteorganic wastewater. Microbial consortium, mixture of definedmicrobial population, releases a variety of enzymes that attackon the chemical structure in different ways to rapidly degradethe complex chemical molecules/compounds as wastewatercontains complex mixture of various chemicals. The use ofmicrobes is, therefore, recommended for the treatment ofwastewater containing azo dyes.

Compliance with ethical standards

Conflict of interest The authors declare that they have no conflict ofinterest.

Research involving human participants and/or animals This articledoes not contain any studies with human participants or animals per-formed by any of the author.

Informed consent N/A

References

Acuner E, Dilek FB (2004) Treatment of tectilon yellow 2G by Chlorellavulgaris. Process Biochem 39:623–631

Abedin MAR (2008) Decolorization and biodegradation of crystal Violetand Malachite Green by Fusarium solani (Martius) Saccardo, acomparative study on biosorption of dyes by the dead fungal bio-mass. Am-Euras J Bot 12:17–31

AjazM, Elahi A, RehmanA (2018) Degradation of azo dye by bacterium,Alishewanella sp. CBL-2 isolated from industrial effluent and itspotential use in decontamination of wastewater. J Water ReuseDesalination 8:507–515

AjazM, RehmanA, Khan Z, NisarMA,Hussain S (2019)Degradation ofazo dyes by Alcaligenes aquatilis 3c and its potential use in thewastewater treatment. AMB Express 9:64

Al-Amrani WA, Lim PE, Seng CE, Ngah WSW (2014) Factors affectingbio-decolorization of azo dyes and COD removal in anoxic–aerobicREACToperated sequencing batch reactor. J Taiwan Inst Chem Eng45:609–616

Asad S, Amoozegar MA, Pourbabaee A, Sarbolouki MN, DastgheibSMM (2007) Decolorization of textile azo dyes by newly isolatedhalophilic and halotolerant bacteria. Bioresour Technol 98:2082–2088

Bafana A, Chakrabarti T, Muthal P, Kanade G (2008) Detoxification ofbenzidine-based azo dye by E. gallinarum: time-course study.Ecotoxicol Environ Saf 72(3):960–964

Banat IM, Nigam P, Singh D, Marchant R (1996) Microbial decoloriza-tion of textile-dye-containing effluents: a review. Bioresour Technol58:217

Behnajady MA, Modirshahla N, Shokri M (2004) Photodestruction ofacid orange 7 (AO7) in aqueous solutions by UV/H2O2: influenceof operational parameters. Chemosphere 55:129–134

Blümel S, Stolz A (2003) Cloning and characterization of the gene codingfor the aerobic azoreductase from Pigmentiphaga kullae K24. ApplMicrobiol Biotechnol 62:186

Bouraie ME, Din WSE (2016) Biodegradation of reactive black 5 byAeromonas hydrophila strain isolated from dye-contaminated textilewastewater. Sustain Environ Res 26:209–216

Brik M, Schoeberl P, Chamam B, Braun R, Fuchs W (2006) Advancedtreatment of textile wastewater towards reuse using amembrane bio-reactor. Process Biochem 41:1751–1757

Bromley-Challenor CA, Knapp JS, Zhang Z, Gray NC, Hetheridge MJ,Evans MR (2000) Decolorization of an azo dye by unacclimatedactivated sludge under anaerobic conditions. Water Res 34(18):4410–4418

Bumpus JA (2004) Biodegradation of azo dyes by fungi. In: Arora DK(ed) Fungal biotechnology in agricultural, food and environmentalapplications. Marcel Dekker, New York, pp 457–480

Carliell CM, Barclay SJ, NaidooN, Buckley CA,Mulholland DA, SeniorE (1995) Microbial decolourization of a reactive azo dye underanaerobic conditions. Water SA 21:61–61

Champagne PP, Ramsay JA (2010) Dye decolorization and detoxificationby laccase immobilized on porous glass beads. Bioresour Technol101:2230–2235

Chang JS, Kuo TS (2000) Kinetics of bacterial decolorization of azo dyewith Escherichia coli NO3. Bioresour Technol 75:107–111

Chang JS, Chou C, Lin YC, Lin PJ, Ho JY, Hu TL (2001) Kinetic char-acteristics of bacterial azo-dye decolorization by Pseudomonasluteola. Water Res 35:2841–2850

Chang JS, Chen BY, Lin YS (2004) Stimulation of bacterial decoloriza-tion of an azo dye by extracellular metabolites from Escherichia colistrain NO3. Bioresour Technol 91:243–248

Chen BY, Chang JS (2007) Assessment upon species evolution of mixedconsortia for azo dye decolorization. J Chin Inst Chem Eng 38:259–266

Chen T, Gao B, Yue Q (2010) Effect of dosing method and pH on colorremoval performance and floc aggregation of polyferric chloride–polyamine dual-coagulant in synthetic dyeing wastewater treatment.Colloids Surf A Physicochem Eng Asp 355:121–129

Chern JM, Chien YW (2003) Competitive adsorption of benzoic acid andp-nitrophenol onto activated carbon: isotherm and breakthroughcurves. Water Res 37:2347–2356

Christie RM (2007) Environmental aspects of textile dyeing. ElsevierDaâssi D, Zouari-Mechichi H, Frikha F, Martinez MJ, Nasri M, Mechichi

T (2013) Decolorization of the azo dye acid orange 51 by laccaseproduced in solid culture of a newly isolated Trametes trogii strain. 3Biotech 3:115–125

Daeshwar N, Ayazloo M, Khataee AR, Pourhassan M (2007) Biologicaldecolorization of dye solution containing malachite green bymicroalgae Cosmarium sp. Bioresour Technol 98:1176

Das A, Mishra S (2017) Removal of textile dye reactive green-19 usingbacterial consortium: process optimization using response surfacemethodology and kinetics study. J Environ Chem Eng 5:612–627

de Aragao Umbuzeiro G, Freeman HS, Warren SH, De Oliveira DP,Terao Y, Watanabe T, Claxton LD (2005) The contribution of azodyes to the mutagenic activity of the Cristais River. Chemosphere60:55–64

de Souza NA, Ramaiah N, Damare S, Furtado B, Mohandass C, Patil A,De Lima M (2019) Differential protein expression in Shewanellaseohaensis decolorizing azo dyes. Curr Proteomics 16:156–164

Int Microbiol

Duran N, Rosa MA, D’Annibale A, Gianfreda L (2002) Applications oflaccases and tyrosinases (phenoloxidases) immobilized on differentsupports: a review. Enzym Microb Technol 31:907

Forgacs E, Cserhati T, Oros G (2004) Removal of synthetic dyes fromwastewaters: a review. Environ Int 30:953–971

Franciscon E, Grossman MJ, Paschoal JAR, Reyes FGR, Durrant LR(2012) Decolorization and biodegradation of reactive sulfonatedazo dyes by a newly isolated Brevibacterium sp. strain VN-15.SpringerPlus 1:37

Fu L, Bai YN, Lu YZ, Ding J, Zhou D, Zeng RJ (2019) Degradation oforganic pollutants by anaerobic methane-oxidizing microorganismsusing methyl orange as example. J Hazard Mater 364:264–271

Gabriel H, Hong J (2008)Decolouration of reactive blue dye KN-R. Res JAppl Sci 3:216–224

Gahlout M, Gupte S, Gupte A (2013) Optimization of culture conditionfor enhanced decolorization and degradation of azo dye reactiveviolet 1 with concomitant production of ligninolytic enzymes byGanoderma cupreum AG-1. 3 Biotech 3:143–152

Ghodake GS, Talke AA, Jadhav JP, Govindwar SP (2009a) Potential ofBrassica juncea in order to treat textile-effluent-contaminated sites.Int J Phytoremed 11(4):297–312

Ghodake G, Jadhav S, Dawkar VS, Govindwar S (2009b)Biodegradation of Diazo dye Direct Brown MR by Acinetobactercalcoaceticus NCIM 2890. Int Biodeterior Biodegradation 63:433

Golab V, Vinder A, Simonic M (2005) Efficiency of the coagulation/flocculation method for the treatment of dye bath effluent. DyesPigments 67:93–97

Gottlieb A, Shaw C, Smith A, Wheatley A, Forsythe S (2003) The tox-icity of textile reactive azo dyes after hydrolysis and decolourisation.J Biotechnol 101:49–56

Guo G, Tian F, Zhang C, Liu T, Yang F, Hu Z, Liu C, Wang S, Ding K(2019) Performance of a newly enriched bacterial consortium fordegrading and detoxifying azo dyes. Water Sci Technol 79:2036–2045

Gupta VK (2009) Application of low-cost adsorbents for dye removal-areview. J Environ Manag 90:2313–2342

He XL, Song C, Li YY, Wang N, Xu L, Han X, Wei DS (2018) Efficientdegradation of azo dyes by a newly isolated fungus Trichodermatomentosum under non-sterile conditions. Ecotoxicol Environ Saf150:232–239

Humnabadkar RP, Saratale GD, Govindwar SP (2008) Decolorization ofpurple 2R by Aspergillus ochraceus (NCIM-1146). Asian JMicrobiol Biotechnol Environ 10:693–697

Hunger K (2007) Industrial dyes: chemistry, properties, applications.Wiley

Ilyas S, Rehman A (2013) Decolorization and detoxification of Synozolred HF-6BN azo dye, by Aspergillus niger and Nigrospora sp.Iranian J Environ Health Sci Eng 10:12

Imran M, Arshad M, Asghar HN, Asghar M, Crowley DE (2014)Potential of Shewanella sp. strain IFN4 to decolourize azo dyesunder optimal conditions. Int J Agric Biol 16:578–584

Jadhav JP, Govindwar SP (2006) Biotransformation of malachite greenby Saccharomyces cerevisiaeMTCC 463. Yeast 23:315–323

Jadhav JP, Parshetti GK, Kalme SD, Govindwar SP (2007)Decolourization of azo dye methyl red by Saccharomyces cerevisiaeMTCC 463. Chemosphere 68:394–400

Jadhav SU, Jadhav MU, Kagalkar AN, Govindwar SP (2008a)Decolorization of Brilliant Blue G dye mediated by degradation ofthe microbial consortium of Galactomyces geotrichum and Bacillussp. J Chin Inst Chem Eng 39:563

Jadhav SU, Jadhav UU, Dawkar VV, Govindwar SP (2008b)Biodegradation of Disperse dye brown 3REL by microbial consor-tium of Galactomyces geotrichum MTCC 1360 and Bacillus sp.VUS. Biotechnol Bioprocess Eng 13:232

Jin XC, Liu GQ, XuZH, TaoWY (2007) Decolorization of a dye industryeffluent by Aspergillus fumigatus XC6. Appl Microbiol Biotechnol74:239–243

Jirasripongpun K, Nasanit R, Niruntasook J, Chotikasatian B (2007)Decolorization and degradation of CI Reactive Red 195 byEnterobacter sp. Thammasat Int J Sci Technol 12:6–11

Joshi T, Iyengar L, Singh K, Garg S (2008) Isolation, identification andapplication of novel bacterial consortium TJ-1 for thedecolourization of structurally different azo dyes. BioresourTechnol 99:7115–7121

Kagalkar AN, Jagtap UB, Jadhav JP, Bapat VA, Govindwar SP (2009)Biotechnological strategies for phytoremediation of the sulfonatedazo dye Direct Red 5B using Blumea malcolmii Hook. BioresourTechnol 100:4104–4110

Kalsoom U, Ashraf SS, Meetani MA, Rauf MA, Bhatti HN (2013)Mechanistic study of a diazo dye degradation by SoybeanPeroxidase. Chem Cent J 7:93

Kalyani DC, Telke AA, Dhanve RS, Jadhav JP (2008) Ecofriendly bio-degradation and detoxification of Reactive Red 2 textile dye bynewly isolated Pseudomonas sp. SUK1. J Hazard Mater 163:735–742

Keck A, Klein J, Kudlich M, Stolz A, Knackmuss H-J, Mattes R (1997)Reduction of azo dyes by redox mediators originating in the naph-thalene sulfonic acid degradation pathway of Sphingomonas sp.strain BN6. Appl Environ Microbiol 63(9):3684–3690

Khalid A, Arshad M, Crowley DE (2009) Biodegradation potential ofpure and mixed bacterial cultures for removal of 4-nitroaniline fromtextile dye wastewater. Water Res 43:1110–1116

Khehra MS, Saini HS, Sharma DK, Chadha BS, Chimni SS (2005)Comparative studies on potential of consortium and constituent purebacterial isolates to decolorize azo dyes. Water Res 39:5135–5141

Kodam KM, Soojhawon I, Lokhande PD, Gawai KR (2005) Microbialdecolorization of reactive azo dyes under aerobic conditions. WorldJ Microbiol Biotechnol 21:367–370

Krishnan J, Kishore AA, Suresh A, Madhumeetha B, Prakash DG (2017)Effect of pH, inoculum dose and initial dye concentration on theremoval of azo dye mixture under aerobic conditions. IntBiodeterior Biodegrad 119:16–27

Kunjadia PD, Patel FD, Nagee A, Mukhopadhyaya PN, Dave GS (2012)Crystal violet (triphenylmethane dye) decolorization potential ofPleurotus ostreatus (MTCC 142). BioResources 7:1189–1199

Kunjadia PD, Sanghvi GV, Kunjadia AP, Mukhopadhyay PN, Dave GS(2016) Role of ligninolytic enzymes of white rot fungi (Pleurotusspp.) grown with azo dyes. SpringerPlus 5:1487

Lin J, Zhang X, Li Z, Lei L (2010) Biodegradation of Reactive Blue 13 ina two-stage anaerobic/aerobic fluidized beds system with aPseudomonas sp. isolate. Bioresour Technol 101:34

Lourenco ND, Novais JM, Pinheiro HM (2000) Reactive textile dyecolour removal in a sequencing batch reactor. Water Sci Technol42:321–328

LucasMS, Dias AA, SampaioA, Amaral C, Peres JA (2007) Degradationof a textile reactive azo dye by a combined chemical–biologicalprocess: Fenton’s reagent-yeast. Water Res 41:1103–1109

Lucious S, Reddy ES, Anuradha V, Yogananth N, Ali MYS, Vijaya P,Rajan R, Parveen PMK (2014) Decolorization of acid dyes byB. cereus and P. aeruginosa isolated from effluent of dyeing indus-try. Int J Pure Appl Biosci 2(3):23–29

Mahalakshmi S, Lakshmi D, Menaga U (2015) Biodegradation of differ-ent concentration of dye (Congo red dye) by using green and bluegreen algae. Int J Environ Res 9:735–744

Maier J, Kandelbauer A, Erlacher A, Cavaco-Paulo A, Gübitz GM (2004)A new alkali-thermostable azoreductase from Bacillus sp. strain SF.Appl Environ Microbiol 70:837–844

Malik SN, Ghosh PC, Vaidya AN, Waindeskar V, Das S, Mudliar SN(2017) Comparison of coagulation, ozone and ferrate treatment

Int Microbiol

processes for color, COD and toxicity removal from complex textilewastewater. Water Sci Technol 76:1001–1010

Mani S, Chowdhary P, Bharagava RN (2019) Textile wastewater dyes:toxicity profile and treatment approaches. In: Emerging and eco-friendly approaches for waste management. Springer, Singapore,pp 219–244

Manu B, Chauhari S (2003) Decolorization of indigo and azo dyes insemicontinuous reactors with long hydraulic retention time. ProcessBiochem 38:1213–1221

Mbuligwe SE (2005) Comparative treatment of dye-rich wastewater inengineered wetland systems (EWSs) vegetated with different plants.Water Res 39:271–280

McMullan G, Meehan C, Conneely A, Kirby N, Robinson T, Nigam P,Smyth WF (2001) Microbial decolourisation and degradation oftextile dyes. Appl Microbiol Biotechnol 56:81–87

Meerbergen K, Willems KA, Dewil R, Van Impe J, Appels L, Lievens B(2018) Isolation and screening of bacterial isolates from wastewatertreatment plants to decolorize azo dyes. J Biosci Bioeng 125:448–456

Mester T, Tien M (2000) Oxidation mechanism of ligninolytic enzymesinvolved in the degradation of environmental pollutants. IntBiodeterior Biodegrad 46:51–59

Misal SA, Lingojwar DP, Shinde RM, Gawai KR (2011) Purification andcharacterization of azoreductase from alkaliphilic strain Bacillusbadius. Process Biochem 46(6):1264–1269

Mishra S, Maiti A (2018) The efficacy of bacterial species to decolourisereactive azo, anthroquinone and triphenylmethane dyes from waste-water: a review. Environ Sci Pollut Res 5:8286–8314

Mohan SV, Rao NC, Prasad KK, Karthikeyan J (2002) Treatment ofsimulated reactive yellow 22 (Azo) dye effluents using Spirogyraspecies. Waste Manag 22:575–582

Mohana S, Shrivastava S, Divecha J, Madamwar D (2008) Responsesurface methodology for optimization of medium for decolorizationof textile dye Direct Black 22 by a novel bacterial consortium.Bioresour Technol 99:562–569

Moosvi S, Keharia H, Madamwar D (2005) Decolourization of textiledye reactive violet 5 by a newly isolated bacterial consortium RVM11.1. World J Microbiol Biotechnol 21:667–672

Moosvi S, Kher X, Madamwar D (2007) Isolation, characterization anddecolorization of textile dyes by a mixed bacterial consortium JW-2.Dyes Pigments 74:723–729

Nachiyar CV, Rajkumar GS (2003) Degradation of tannery and textiledye, Navian Fast Blue S5R by Pseudomonas aerugenosa. World JMicrobiol Biotechnol 19:609

Nachiyar CV, Rajkumar GS (2005) Purification and characterization of anoxygen insensitive azoreductase from Pseudomonas aeruginosa.Enzym Microb Technol 36:503–509

O’neill C, Lopez A, Esteves S, Hawkes FR, Hawkes DL, Wilcox S(2000) Azo-dye degradation in an anaerobic-aerobic treatment sys-tem operating on simulated textile effluent. Appl MicrobiolBiotechnol 53:249–254

Olligaard H, Frost L, Gaister J, Hansen OC (1999) Survey of azocolourant in Denmark. Miljoproject. 509 Danish EPA

Omar HH (2008) Algal decolorization and degradation of monoazo anddiazo dyes. Pak J Biol Sci 11:1310–1316

Pandey A, Singh P, Iyengar L (2007) Bacterial decolorization and degra-dation of azo dyes. Int Biodeterior Biodegrad 59:73–84

Pandi A, Kuppuswami GM, Ramudu KN, Palanivel S (2019) A sustain-able approach for degradation of leather dyes by a new fungallaccase. J Clean Prod 211:590–597

Patil P, Desai N, Govindwar S, Jadhav JP, Bapat V (2009) Degradationanalysis of reactive red 198 by hairy roots of Tagetes patulaL.(Marigold). Planta 230:725–735

Pearce CI, Lloyd JR, Guthrie JT (2003) The removal of colour fromtextile wastewater using whole bacterial cells: a review. DyesPigments 58:179–196

Perumal K, Malleswari RB, Catherin A, Moorthy TS (2017)Decolourization of Congo red dye by bacterial consortium isolatedfrom dye contaminated soil, Paramakudi, Tamil Nadu. J MicrobiolBiotechnol Res 2:475–480

Pinheiro HM, Touraud E, Thomas O (2004) Aromatic amines from azodye reduction: status review with emphasis on direct UV spectro-photometric detection in textile industry wastewaters. DyesPigments 61:121–139

Pointing SB, Vrijmoed LLP (2000) Decolorization of azo and triphenyl-methane dyes by Pycnoporus sanguineus producing laccase as thesole phenoloxidase. World J Microbiol Biotechnol 16:317–318

Popli S, Patel UD (2015) Destruction of azo dyes by anaerobic–aerobicsequential biological treatment: a review. Int J Environ Sci Technol12:405–420

Puvaneswari N,Muthukrishnan J, Gunasekaran P (2006) Toxicity assess-ment and microbial degradation of azo dyes. Indian J Exp Biol 44:618–626

Rafatullah M, Sulaiman O, Hashim R, Ahmad A (2010) Adsorption ofmethylene blue on low-cost adsorbents: a review. J Hazard Mater177:70–80

Rai HS, Bhattacharyya MS, Singh J, Bansal TK, Vats P, Banerjee UC(2005) Removal of dyes from the effluent of textile and dyestuffmanufacturing industry: a review of emerging techniques with ref-erence to biological treatment. Crit Rev Environ Sci Technol 35:219–238

Rajaguru P, Vidya L, Baskarasethupathi B, Kumar PA, Palanivel M,Kalaiselvi K (2002) Genotoxicity evaluation of polluted groundwater in human peripheral blood lymphocytes using the comet as-say. Mutat Res/Genet Toxicol Environm Mutagen 517:29–37

Ramalho PA, Cardoso MH, Cavaco-Paulo A, Ramalho MT (2004)Characterization of azo reduction activity in a novel ascomyceteyeast strain. Appl Environ Microbiol 70:2279–2288

Ramya M, Anusha B, Kalavathy S, Devilaksmi S (2007)Biodecolorization and biodegradation of reactive blue byAspergillus sp. Afr J Biotechnol 6:1441–1445

Robinson T, McMullan G, Marchant R, Nigam P (2001) Remediation ofdyes in textile effluent: a critical review on current treatment tech-nologies with a proposed alternative. Bioresour Technol 77:247–255

Roy U,Manna S, Sengupta S, Das P, Datta S, Mukhopadhyay A, BhowalA (2018) Dye removal using microbial biosorbents. In: Green ad-sorbents for pollutant removal. Springer, pp 253–280

Ryan A, Wang CJ, Laurieri N, Westwood I, Sim E (2010) Reactionmechanism of azoreductases suggests convergent evolution withquinone oxidoreductases. Protein Cell 1:780–790

Saha AK, Sultana N, Mohanta MK, Mandald A, Haquee M (2017)Identification and characterization of azo dye decolourizing bacterialstrains, Alcaligenes faecalis E5.Cd and A. faecalis Fal.3 isolatedfrom textile effluents. Am Sci Res J Eng Technol Sci 3:163–175

Sahasrabudhe MM, Saratale RG, Saratale GD, Pathade GR (2014)Decolorization and detoxification of sulfonated toxic diazo dye CIdirect red 81 by Enterococcus faecalis YZ 66. J Environ Health SciEng 12:151

Sandesh K, Kumar G, Chidananda B, Ujwal P (2019) Optimization ofdirect blue-14 dye degradation by Bacillus fermus (Kx898362) analkaliphilic plant endophyte and assessment of degraded metabolitetoxicity. J Hazard Mater 364:742–751

Sandhya S, Sarayu K, Swaminathan K (2008) Determination of kineticconstants of hybrid textile wastewater treatment system. BioresourTechnol 99:5793–5797

Saratale GD, Kalme SD, Govindwar SP (2006) Decolorisation of textiledyes byAspergillus ochraceus (NCIM-1146). Indian J Biotechnol 5:407–410

Saratale RG, Saratale GD, Chang JS, Govindwar SP (2009a) Ecofriendlydegradation of sulphonated diazo dye reactive green 19A using

Int Microbiol

Micrococcus glutamicus NCIM 2168. Bioresour Technol 100:3897–3905

Saratale RG, Saratale GD, Kalyani DC, Chang JS, Govindwar SP(2009b) Enhanced decolorization and biodegradation of textile azodye scarlet R by using developed microbial consortium-GR.Bioresour Technol 100:2493

Saratale RG, Saratale GD, Chang JS, Govindwar SP (2009c)Decolorization and biodegradation of textile dye navy blue HERby Trichosporon beigelii NCIM-3326. J Hazard Mater 166:1421

Saratale RG, Saratale GD, Chang JS, Govindwar SP (2010)Decolorization and biodegradation of reactive dyes and dye waste-water by a developed bacterial consortium. Biodegradation 21:999–1015

Sarayu K, Sandhya S (2010) Aerobic biodegradation pathway forremazol orange by Pseudomonas aeruginosa. Appl BiochemBiotechnol 160:1241–1253

Sarkar S, Banerjee A, Halder U, Biswas R, Bandopadhyay R (2017)Degradation of synthetic azo dyes of textile industry: a sustainableapproach using microbial enzymes. Water Conserv Sci Eng 2:121–131

Şen S, Demirer GN (2003) Anaerobic treatment of real textile wastewaterwith a fluidized bed reactor. Water Res 37:1868–1878

Sen SK, Raut S, Bandyopadhyay P, Raut S (2016) Fungal decolourationand degradation of azo dyes: a review. Fungal Biol Rev 30:112–133

Senan RC, Abraham TE (2004) Bioremediation of textile azo dyes byaerobic bacterial consortium aerobic degradation of selected azodyes by bacterial consortium. Biodegradation 15:275–280

Shah MP (2014) Environmental bioremediation: a low cost nature’s nat-ural biotechnology for environmental clean-up. J Pet EnvironBiotechnol 5:1–12

Sheth NT, Dave SR (2009) Optimization for enhanced decolorization ofReactive Red BS C.I. 111 by Pseudomonas aeruginosa NGKCTS.Biodegradation 20:827–836

Singh P, Sanghi R, Pandey A, Iyengar L (2007) Decolourization andpartial degradation of monoazo dyes in sequential fixed film anaer-obic batch reactor (SFABR). Bioresour Technol 98:2053–2056

Singh S, Chatterji S, Nandini PT, Prasad ASA, Rao KVB (2015)Biodegradation of azo dye Direct Orange 16 byMicrococcus luteusstrain SSN2. Int J Environ Sci Technol 12:2161–2168

Song Z, Song L, Shao Y, Tan L (2018) Degradation and detoxification ofazo dyes by a salt-tolerant yeast Cyberlindnera samutprakarnensisS4 under high-salt conditions. World J Microbiol Biotechnol 34:131

Stolz A (2001) Basic and applied aspects in the microbial degradation ofazo dyes. Appl Microbiol Biotechnol 56:69–80

Swaminathan K, Sandhya S, Sophia AC, Pachhade K, SubrahmanyamYV (2003) Decolorization and degradation of H-acid and other dyesusing ferrous-hydrogen peroxide system. Chemosphere 50:619–625

Tony BD, Goyal D, Khanna S (2009a) Decolorization of textile azo dyesby aerobic bacterial consortium. Int Biodeterior Biodegradation 63:462

Tony BD, Goyal D, Khanna S (2009b) Decolorization of direct red 28 bymixed bacterial culture in an Up-Flow immobilized bioreactor. J IndMicrobiol Biotechnol 36:955

Van der Zee FP, Villaverde S (2005) Combined anaerobic-aerobic treat-ment of azo dyes-a short review of bioreactor studies. Water Res 39:1425–1440

Van der Zee FP, Lettinga G, Field JA (2001) Azo dye decolourisation byanaerobic granular sludge. Chemosphere 44:1169–1176

Verma P, Madamwar D (2003) Decolorization of synthetic dyes by anewly isolated strain of Serratia maerascens. World J MicrobiolBiotechnol 19:615–618

Verma K, Saha G, Kundu LM, Dubey VK (2019) Biochemical charac-terization of a stable azoreductase enzyme from Chromobacteriumviolaceum: application in industrial effluent dye degradation. Int JBiol Macromol 121:1011–1018

Vijayaraghavan K, Yun YS (2008) Biosorption of CI reactive black 5from aqueous solution using acid-treated biomass of brown seaweedLaminaria sp. Dyes Pigments 76:726–732

Vikrant K, Giri BS, Raza N, Roy K, Kim KH, Rai BN, Singh RS (2018)Recent advancements in bioremediation of dye: current status andchallenges. Bioresour Technol 253:355–367

Waghmode TR, Kurade MB, Khandare RV, Govindwar SP (2011) Asequential aerobic/microaerophilic decolorization of sulfonatedmono azo dye Golden Yellow HER by microbial consortium GG–BL. Int Biodeter Biodegrad 65:1024–1034

Wang ZW, Liang JS, Liang Y (2013) Decolorization of reactive black 5by a newly isolated bacterium Bacillus sp. YZU1. Int BiodeteriorBiodegrad 76:41–48

Wang P-T, Song Y-H, Fan H-C, Yu L (2018) Bioreduction of azo dyeswas enhanced by in-situ biogenic palladium nanoparticles.Bioresour Technol 266:176–180

Wuhrmann K, Mechsner K, Kappeler TH (1980) Investigation on ratedetermining factors in the microbial reduction of azo dyes. ApplMicrobiol Biotechnol 9:325–338

Yan H, Pan G (2004) Increase in biodegradation of dimethyl phthalate byClosterium lunula using inorganic carbon. Chemosphere 55:1281–1285

Yang Q, Yang M, Pritsch K, Yediler A, Hagn A, Schloter M, Kettrup A(2003) Decolorization of synthetic dyes and production ofmanganese-dependent peroxidase by new fungal isolates.Biotechnol Lett 25:709–713

Yu J, Wang X, Yue PL (2001) Optimal decolorization and kinetic model-ing of synthetic dyes by Pseudomonas strains. Water Res 35:3579–3586

Zahran SA, Ali-Tammam M, Hashem AM, Aziz RK, Ali AE (2019)Azoreductase activity of dye-decolorizing bacteria isolated fromthe human gut microbiota. Sci Rep 9:5508

Zhang F, Yediler A, Liang X, Kettrup A (2004) Effects of dye additiveson the ozonation process and oxidation by products: a comparativestudy using hydrolysed CI Reactive red 120. Dyes Pigments 60:1–7

Zhao G, Jiang L, He Y, Li J, Dong H, Wang X, Hu W (2011) Sulfonatedgraphene for persistent aromatic pollutant management. Adv Mater23:3959–3963

Zhou X, Xiang X (2013) Effect of different plants on azo-dye wastewaterbio-decolorization. Procedia Environ Sci 18:540–546

Zimmermann T, Gasser F, Kulla H, Leisinger T (1984) Comparison oftwo bacterial azoreductases acquired during adaptation to growth onazo dyes. Arch Microbiol 138:37

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Int Microbiol

Clarity - Chromatography SWDataApex 2006

www.dataapex.com

Chromatogram Info:

File Name : C:\CLARITY\WORK1\DATA\C1.PRM File Created : 6/5/2017 3:14:57 PM

Origin : Acquired Acquired Date : 6/5/2017 3:14:57 PM

Project : c:\Clarity\Projects\Work1.PRJ By : Administrator

Printed Version Info:

Printed Version : 6/5/2017 3:14:57 PM Printed Date : 6/5/2017 3:15:57 PM

Report Sty le : c:\Clarity\Common\Chromatogram.sty By : Administrator

Calibration File : None

Sample Info:

Sample ID : c1 Amount : 0

Sample : secondary metabolite ISTD Amount : 0

Inj. Volume [ml] : 20 Dilution : 1

Method : Bioactive secondary metabolite By : Administrator

Description : Bioactive secondary metabolites

Created : 1/17/2013 10:08 PM Modified : 6/5/2017 3:14 PM

Column : RPC18 Detection : UV254

Mobile Phase : Methanol Temperature : Ambiant

Flow Rate : 1 ml/min Pressure :

Note :

Autostop : 15.00 min External Start : Start - Stop, Down

Detector 1 : Signal 1 Range 1 : Bipolar, 1250 mV, 12.5 Samp. per Sec.

Subtraction Chromatogram : (None) Matching : No Change

Base : Not Used Calibration File : None Calculation : Uncal

Scale Factor : Not Used Units After Scaling : Not Used Uncal. Response : 0

Unretained Time : 0.00 min Column Length : 50.00 mm Column Calc. : From Width at 50% of Height

Result Table Reports : All Peaks Hide ISTD Peak : Enabled

[min.]Time

0 2 4 6 8 10 12 14

[mV]

Volta

ge

0

50

100

150

[%]

Components

0

20

40

60

80

100

0.72

1.22

1.

39

1.80

2.60

2.

88

5.00

6.70

7.17

7.

52

9.65

10.3

2

13.0

8

13.6

2

14.1

1 14

.38

14.4

6

6/5/2017 3:15 PM Page 1 of 2Chromatogram C:\CLARITY\WORK1\DATA\C1.PRM

Result Table (Uncal - C1)

Reten. Time[min]

Area[mV.s]

Height[mV]

Area[%]

Height[%]

W05[min]

1 0.720 41.554 1.313 0.8 0.5 0.72

2 1.224 12.926 1.366 0.2 0.6 0.17

3 1.388 24.998 1.720 0.5 0.7 0.28

4 1.804 519.123 22.770 9.8 9.4 0.31

5 2.596 73.253 6.446 1.4 2.7 0.20

6 2.876 2990.816 154.332 56.2 63.6 0.22

7 4.996 901.448 38.401 16.9 15.8 0.21

8 6.696 36.854 2.595 0.7 1.1 0.27

9 7.168 44.532 2.561 0.8 1.1 0.35

10 7.524 183.586 3.115 3.4 1.3 1.25

11 9.648 146.977 2.329 2.8 1.0 1.27

12 10.324 307.997 2.609 5.8 1.1 2.73

13 13.080 7.618 0.542 0.1 0.2 0.13

14 13.624 2.008 0.152 0.0 0.1 0.02

15 14.112 14.342 0.882 0.3 0.4 0.25

16 14.380 6.662 0.702 0.1 0.3 0.18

17 14.460 7.273 0.727 0.1 0.3 0.21

Total 5321.967 242.562 100.0 100.0

6/5/2017 3:15 PM Page 2 of 2Chromatogram C:\CLARITY\WORK1\DATA\C1.PRM


www.dataapex.com

Chromatogram Info:

File Name : C:\CLARITY\WORK1\DATA\1B(AGAIN).PRM File Created : 6/5/2017 1:37:08 PM







Sample Info:

Sample ID : 1b(again) Amount : 0









Note :








[min.]Time

0 2 4 6 8 10 12 14

[mV]

Volta

ge

0

200

400

600

800

[%]

Components

0

20

40

60

80

100

0.76

1.97

2.92

4.99

5.39

7.62

7.

93

8.23

8.89

9.77

11.3

5

11.7

4 12

.05

12.6

7

13.3

8

14.9

6

6/5/2017 1:38 PM Page 1 of 2Chromatogram C:\CLARITY\WORK1\DATA\1B(AGAIN).PRM

Result Table (Uncal - 1B(AGAIN))

Reten. Time[min]

Area[mV.s]

Height[mV]

Area[%]

Height[%]

W05[min]

1 0.760 29.637 1.278 0.2 0.1 0.43

2 1.972 1034.413 47.013 5.9 5.1 0.37

3 2.916 14192.904 774.317 80.4 84.7 0.24

4 4.988 137.866 6.807 0.8 0.7 0.43

5 5.392 195.871 3.930 1.1 0.4 1.00

6 7.620 230.357 4.166 1.3 0.5 1.32

7 7.928 105.543 4.666 0.6 0.5 0.41

8 8.228 214.134 16.786 1.2 1.8 0.21

9 8.888 600.227 23.048 3.4 2.5 0.48

10 9.772 599.731 23.076 3.4 2.5 0.45

11 11.348 144.130 2.287 0.8 0.3 1.18

12 11.736 55.870 2.390 0.3 0.3 0.47

13 12.052 39.800 2.134 0.2 0.2 0.36

14 12.672 32.671 1.570 0.2 0.2 0.42

15 13.384 33.119 0.934 0.2 0.1 0.51

16 14.964 0.317 0.100 0.0 0.0 0.02

Total 17646.591 914.503 100.0 100.0

6/5/2017 1:38 PM Page 2 of 2Chromatogram C:\CLARITY\WORK1\DATA\1B(AGAIN).PRM


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Chromatogram Info:

File Name : c:\Clarity\Work1\DATA\00000003.PRM File Created : 3/15/2018 3:45:01 PM







Sample Info:

Sample ID : c1 Amount : 0

Sample : Secondary metbolites ISTD Amount : 0








Note :








[min.]Time

0 2 4 6 8 10 12 14

[V]

Volta

ge

-1.2

-1.1

-1.0

-0.9

-0.8[%]

Components

0

20

40

60

80

100

0.19

0.

42

0.57

0.

72

0.85

0.

98

1.17

1.

34

1.70

2.11

3.01

4.32

4.

61

4.75

4.

98

5.32

5.

68

5.94

6.

27

6.39

6.78

7.

03

7.18

7.64

7.

93

8.21

8.

45

8.58

8.

82

9.00

9.

09

9.41

9.

57

9.79

10

.11

10.2

5 10

.36

10.5

6 10

.68

10.9

3 11

.09

11.2

6 11

.40

11.7

0 11

.97

12.0

6 12

.29

12.4

2 12

.65

12.8

1 13

.00

13.2

5 13

.40

13.6

6 13

.91

14.2

0 14

.34

14.5

1 14

.64

14.8

6

3/15/2018 3:54 PM Page 1 of 2Chromatogram c:\Clarity\Work1\DATA\00000003.PRM

Result Table (Uncal - 00000003)

Reten. Time[min]

Area[mV.s]

Height[mV]

Area[%]

Height[%]

W05[min]

1 0.092 4.156 0.913 0.0 0.1 0.08

2 0.188 3.362 0.777 0.0 0.1 0.08

3 0.416 21.076 2.393 0.2 0.4 0.10

4 0.568 9.349 1.724 0.1 0.3 0.11

5 0.724 3.944 0.792 0.0 0.1 0.10

6 0.848 4.724 1.012 0.0 0.2 0.09

7 0.984 13.676 1.757 0.1 0.3 0.14

8 1.168 10.305 1.890 0.1 0.3 0.09

9 1.336 15.821 1.699 0.1 0.3 0.17

10 1.700 47.680 4.475 0.4 0.7 0.20

11 2.108 1095.653 44.010 8.4 7.1 0.32

12 3.008 10337.866 420.250 79.7 68.0 0.31

13 4.324 158.295 10.878 1.2 1.8 0.28

14 4.608 75.943 8.285 0.6 1.3 0.18

15 4.748 69.444 6.924 0.5 1.1 0.20

16 4.976 45.506 5.066 0.4 0.8 0.16

17 5.316 137.392 6.902 1.1 1.1 0.39

18 5.684 26.921 4.054 0.2 0.7 0.13

19 5.936 76.910 4.839 0.6 0.8 0.30

20 6.268 10.467 1.851 0.1 0.3 0.12

21 6.388 47.338 2.979 0.4 0.5 0.34

22 6.776 33.193 2.809 0.3 0.5 0.23

23 7.028 15.066 2.817 0.1 0.5 0.10

24 7.184 43.185 3.336 0.3 0.5 0.27

25 7.636 55.355 2.885 0.4 0.5 0.42

26 7.928 30.346 2.726 0.2 0.4 0.22

27 8.212 36.556 3.382 0.3 0.5 0.21

28 8.448 13.708 2.200 0.1 0.4 0.12

29 8.584 15.529 2.335 0.1 0.4 0.13

30 8.824 25.314 2.911 0.2 0.5 0.17

31 9.004 13.891 2.387 0.1 0.4 0.11

32 9.092 26.553 2.917 0.2 0.5 0.18

33 9.412 27.437 2.252 0.2 0.4 0.24

34 9.568 14.008 2.134 0.1 0.3 0.13

35 9.792 20.859 2.317 0.2 0.4 0.18

36 10.108 19.996 1.398 0.2 0.2 0.29

37 10.252 7.522 1.380 0.1 0.2 0.10

38 10.360 12.742 1.697 0.1 0.3 0.14

39 10.564 7.284 1.474 0.1 0.2 0.09

40 10.684 26.651 2.100 0.2 0.3 0.23

41 10.932 5.740 1.074 0.0 0.2 0.10

42 11.092 16.340 2.554 0.1 0.4 0.11

43 11.260 6.892 1.314 0.1 0.2 0.09

44 11.400 25.747 2.508 0.2 0.4 0.19

45 11.704 17.646 1.684 0.1 0.3 0.17

46 11.968 7.621 1.287 0.1 0.2 0.11

47 12.060 13.091 1.966 0.1 0.3 0.12

48 12.292 9.039 1.226 0.1 0.2 0.12

49 12.420 13.665 1.631 0.1 0.3 0.16

50 12.652 15.619 2.301 0.1 0.4 0.12

51 12.812 14.150 1.840 0.1 0.3 0.15

52 13.004 13.133 1.990 0.1 0.3 0.11

53 13.248 15.762 2.297 0.1 0.4 0.12

54 13.404 13.476 1.752 0.1 0.3 0.15

55 13.660 28.255 2.659 0.2 0.4 0.20

56 13.908 36.171 2.542 0.3 0.4 0.28

57 14.204 11.600 1.647 0.1 0.3 0.14

58 14.336 9.066 1.318 0.1 0.2 0.14

59 14.508 12.811 2.007 0.1 0.3 0.12

60 14.644 18.821 1.785 0.1 0.3 0.20

61 14.856 12.511 1.954 0.1 0.3 0.10

Total 12978.180 618.262 100.0 100.0

3/15/2018 3:54 PM Page 2 of 2Chromatogram c:\Clarity\Work1\DATA\00000003.PRM


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Chromatogram Info:

File Name : C:\CLARITY\WORK1\DATA\2P.PRM File Created : 6/5/2017 2:05:22 PM







Sample Info:

Sample ID : 2p Amount : 0









Note :








[min.]Time

0 2 4 6 8 10 12 14

[mV]

Volta

ge

0

50

100

150

200

250

300

[%]

Components

0

20

40

60

80

100

0.36

0.82

1.75

1.92

2.61

2.

89

4.62

4.86

6.72

7.70

8.48

10.2

6

10.8

4 10

.95

12.7

2

13.2

2

14.0

7 14

.38

14.9

6

6/5/2017 3:16 PM Page 1 of 2Chromatogram C:\CLARITY\WORK1\DATA\2P.PRM

Result Table (Uncal - 2P)

Reten. Time[min]

Area[mV.s]

Height[mV]

Area[%]

Height[%]

W05[min]

1 0.364 6.867 0.515 0.1 0.1 0.07

2 0.820 4.026 0.640 0.1 0.2 0.08

3 1.748 42.660 1.135 0.8 0.3 0.59

4 1.916 318.113 19.911 6.0 5.5 0.17

5 2.612 46.247 4.104 0.9 1.1 0.21

6 2.892 4224.109 294.905 79.1 81.7 0.17

7 4.624 13.761 2.068 0.3 0.6 0.13

8 4.864 492.202 29.676 9.2 8.2 0.19

9 6.724 8.834 0.586 0.2 0.2 0.15

10 7.696 33.966 0.968 0.6 0.3 0.48

11 8.476 10.858 0.581 0.2 0.2 0.14

12 10.260 45.171 0.993 0.8 0.3 0.95

13 10.844 15.934 1.170 0.3 0.3 0.23

14 10.952 28.129 0.943 0.5 0.3 0.16

15 12.716 10.088 0.744 0.2 0.2 0.22

16 13.224 2.844 0.579 0.1 0.2 0.08

17 14.072 25.697 0.572 0.5 0.2 0.07

18 14.384 9.862 0.529 0.2 0.1 0.23

19 14.960 3.128 0.190 0.1 0.1 0.04

Total 5342.497 360.810 100.0 100.0

6/5/2017 3:16 PM Page 2 of 2Chromatogram C:\CLARITY\WORK1\DATA\2P.PRM


www.dataapex.com

Chromatogram Info:

File Name : C:\CLARITY\WORK1\DATA\4V.PRM File Created : 6/5/2017 2:48:21 PM







Sample Info:

Sample ID : 4 Amount : 0









Note :








[min.]Time

0 2 4 6 8 10 12 14

[mV]

Volta

ge

0

100

200

300

400

500

[%]

Components

0

20

40

60

80

100

0.24

0.90

1.50

1.

56

1.94

2.91

4.92

6.37

6.95

7.

30

8.27

9.29

10.3

1 10

.51

10.6

3 10

.80

11.3

2

12.0

3

12.5

3

13.7

3

6/5/2017 3:16 PM Page 1 of 2Chromatogram C:\CLARITY\WORK1\DATA\4V.PRM

Result Table (Uncal - 4V)

Reten. Time[min]

Area[mV.s]

Height[mV]

Area[%]

Height[%]

W05[min]

1 0.244 3.514 0.863 0.0 0.1 0.07

2 0.896 48.174 1.822 0.4 0.3 0.24

3 1.496 33.227 1.778 0.3 0.3 0.38

4 1.556 17.976 2.042 0.2 0.3 0.16

5 1.940 560.499 27.988 4.8 4.0 0.26

6 2.912 8506.937 521.400 72.2 75.1 0.20

7 4.916 1337.122 78.881 11.3 11.4 0.19

8 6.368 266.880 18.990 2.3 2.7 0.23

9 6.952 34.779 1.713 0.3 0.2 0.35

10 7.304 18.994 1.226 0.2 0.2 0.12

11 8.268 45.738 1.212 0.4 0.2 0.61

12 9.292 17.250 1.196 0.1 0.2 0.12

13 10.312 35.784 1.112 0.3 0.2 0.64

14 10.508 15.214 1.504 0.1 0.2 0.19

15 10.628 17.161 1.735 0.1 0.2 0.20

16 10.800 46.550 2.939 0.4 0.4 0.13

17 11.320 23.207 1.937 0.2 0.3 0.14

18 12.028 70.256 1.733 0.6 0.2 0.81

19 12.532 63.801 2.204 0.5 0.3 0.75

20 13.732 619.199 21.935 5.3 3.2 0.50

Total 11782.264 694.211 100.0 100.0

6/5/2017 3:16 PM Page 2 of 2Chromatogram C:\CLARITY\WORK1\DATA\4V.PRM


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Chromatogram Info:

File Name : C:\CLARITY\WORK1\DATA\3C.PRM File Created : 2/23/2017 1:24:21 PM







Sample Info:

Sample ID : 3c Amount : 0

Sample : Bioactive Secondary Metabolites ISTD Amount : 0








Note :








[min.]Time

0 2 4 6 8 10 12 14

[V]

Volta

ge

0.0

0.2

0.4

0.6

0.8

1.0

[%]

Components

0

20

40

60

80

100

0.44

0.

72

1.90

1.

99

2.30

2.95

3.76

4.22

5.03

5.

18

5.53

5.

84

6.72

7.32

7.

40

7.65

7.

98

8.24

8.

50

10.5

2 10

.62

12.8

0

13.6

9 13

.82

14.7

2

2/24/2017 3:17 PM Page 1 of 2Chromatogram C:\CLARITY\WORK1\DATA\3C.PRM

Result Table (Uncal - 3C)

Reten. Time[min]

Area[mV.s]

Height[mV]

Area[%]

Height[%]

W05[min]

1 0.128 4.506 0.382 0.0 0.0 0.13

2 0.436 2.318 0.362 0.0 0.0 0.13

3 0.716 24.661 1.246 0.2 0.1 0.30

4 1.896 35.679 4.975 0.3 0.5 0.12

5 1.992 243.322 27.524 2.3 2.6 0.14

6 2.300 292.056 17.784 2.7 1.7 0.27

7 2.948 9384.845 976.853 88.2 91.9 0.14

8 3.756 191.200 12.755 1.8 1.2 0.15

9 4.224 68.468 3.468 0.6 0.3 0.31

10 5.032 24.811 1.623 0.2 0.2 0.34

11 5.184 25.318 1.786 0.2 0.2 0.27

12 5.532 27.882 1.443 0.3 0.1 0.36

13 5.836 53.809 1.818 0.5 0.2 0.47

14 6.720 40.995 1.674 0.4 0.2 0.35

15 7.320 2.265 0.365 0.0 0.0 0.05

16 7.396 3.440 0.591 0.0 0.1 0.09

17 7.648 3.034 0.384 0.0 0.0 0.08

18 7.976 7.823 0.769 0.1 0.1 0.20

19 8.240 13.618 0.905 0.1 0.1 0.30

20 8.504 96.406 1.689 0.9 0.2 0.44

21 10.524 3.199 0.357 0.0 0.0 0.06

22 10.624 15.876 1.118 0.1 0.1 0.11

23 12.804 13.699 0.350 0.1 0.0 0.42

24 13.688 4.042 0.397 0.0 0.0 0.23

25 13.824 9.248 1.428 0.1 0.1 0.09

26 14.720 41.933 1.313 0.4 0.1 0.56

Total 10634.453 1063.358 100.0 100.0

2/24/2017 3:17 PM Page 2 of 2Chromatogram C:\CLARITY\WORK1\DATA\3C.PRM


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Chromatogram Info:

File Name : C:\CLARITY\WORK1\DATA\3D.PRM File Created : 2/23/2017 1:03:59 PM







Sample Info:

Sample ID : 3d Amount : 0

Sample : Bioactive Secondary Metabolites ISTD Amount : 0








Note :








[min.]Time

0 2 4 6 8 10 12 14

[mV]

Volta

ge

0

100

200

300

400

500

600

700[%]

Components

0

20

40

60

80

100

0.99

1.93

2.

09

2.95

3.75

4.70

5.18

7.28

7.91

8.65

9.32

12.1

7

13.7

4

14.6

8

2/24/2017 3:17 PM Page 1 of 2Chromatogram C:\CLARITY\WORK1\DATA\3D.PRM

Result Table (Uncal - 3D)

Reten. Time[min]

Area[mV.s]

Height[mV]

Area[%]

Height[%]

W05[min]

1 0.992 5.022 0.950 0.0 0.1 0.09

2 1.928 142.454 13.261 1.3 1.6 0.18

3 2.092 1378.844 122.456 13.0 15.0 0.10

4 2.952 8328.501 654.345 78.3 80.2 0.18

5 3.748 279.502 10.351 2.6 1.3 0.25

6 4.700 51.554 2.939 0.5 0.4 0.40

7 5.176 67.574 3.389 0.6 0.4 0.26

8 7.276 50.320 1.418 0.5 0.2 0.88

9 7.912 58.751 1.479 0.6 0.2 0.77

10 8.648 52.843 1.714 0.5 0.2 0.62

11 9.324 96.503 1.713 0.9 0.2 0.66

12 12.172 73.807 0.933 0.7 0.1 0.70

13 13.736 31.109 0.920 0.3 0.1 0.43

14 14.684 17.389 0.496 0.2 0.1 0.50

Total 10634.175 816.364 100.0 100.0

2/24/2017 3:17 PM Page 2 of 2Chromatogram C:\CLARITY\WORK1\DATA\3D.PRM

azo dyes removal by bacteria isolated from industrial wastewater

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Transcript of azo dyes removal by bacteria isolated from industrial wastewater