PB

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Journal of Hazardous Materials 192 (2011) 1746– 1755

Contents lists available at ScienceDirect

Journal of Hazardous Materials

j our na l ho me p age: www.elsev ier .com/ locate / jhazmat

referential biodegradation of structurally dissimilar dyes from a mixture byrevibacillus laterosporus

ayur B. Kuradea, Tatoba R. Waghmodeb, Sanjay P. Govindwarb,∗

Department of Biotechnology, Shivaji University, Kolhapur 416004, IndiaDepartment of Biochemistry, Shivaji University, Kolhapur 416004, India

r t i c l e i n f o

rticle history:eceived 26 March 2011eceived in revised form 23 June 2011ccepted 3 July 2011vailable online 8 July 2011

a b s t r a c t

Biodegradation of a mixture containing seven commercial textile dyes with different structures andcolor properties has been investigated by an ecofriendly strain – Brevibacillus laterosporus MTCC 2298.It showed 87% decolorization in terms of ADMI removal (American Dye Manufacturing Institute) within24 h. The effective decolorization of dye mixture was attained in the presence of metal salt – CaCl2 andnitrogen sources. The induction of oxido-reductive enzymes such as veratryl alcohol oxidase, tyrosinase,

eywords:PTLCecolorizationDMIiotransformation

NADH-DCIP reductase and azo reductase was found to be responsible for biotransformation of dyes. Highperformance thin layer chromatography exposed the mechanism of preferential biodegradation of dyesat different time periods. Significant change in the high pressure liquid chromatography and Fouriertransform infrared spectroscopy of sample before and after treatment confirmed the biodegradation ofdye mixture. Phytotoxicity study revealed the much less toxic nature of the metabolites produced after

ixtu

ye mixture the degradation of dyes m

. Introduction

Synthetic dyes and pigments released to the environment in theorm of effluents by textile, leather and printing industries causeevere ecological damages. Ichalkaranji area, India is heavily indus-rialized, that generate millions of liters of untreated effluents peray which are directly discharged into the Panchganga River. Thislters pH, increases biochemical oxygen demand (BOD) and chem-cal oxygen demand (COD) and gives intense coloration. Even a

inimal amount of dye as low as 0.005 ppm is highly visible andan be toxic to aquatic organisms as it interferes with the penetra-ion of sunlight [1]. In addition, several dyes have been found to bearcinogenic and mutagenic [2,3].

Conventional waste water treatments (chemical oxidation,everse osmosis, coagulation–flocculation, filtration, adsorption,nd photodegradation) are not efficient to remove recalcitrantyestuffs from effluents. These technologies are not suitable due toigh cost, low efficiency and inapplicability to a wide variety of dyes4,5]. Instead, bioremediation is definitely an attractive tool whichs currently of interest to overcome the problems arising from the

ndustrial waste water as this is an ecofriendly, cost effective tech-ology. Many biological agents are capable of degrading textile

∗ Corresponding author. Tel.: +91 231 2609152; fax: +91 231 2691533.E-mail addresses: spg biochem@unishivaji.ac.in, spgovindwar@rediffmail.com

S.P. Govindwar).

304-3894/$ – see front matter © 2011 Elsevier B.V. All rights reserved.oi:10.1016/j.jhazmat.2011.07.004

re.© 2011 Elsevier B.V. All rights reserved.

dyes, including bacteria [6], fungi [7], yeast [8,9], actinomycetes[10], algae [11] and plants [12,13].

Textile effluents often include more than one component andstudy of the possible interaction between different chromophoreswill be useful for the treatment. Various approaches were imple-mented to degrade the mixture of dyes includes ozonation[14], flocculation [15], photodegradation [16] and biodegradation[17,18]. The significant role of oxido-reductive enzymes has beenreported in the biodegradation of mixture of dyes [19–21].

The present study is a novel attempt to find out the preferentialdegradation of individual dye in the mixture by Brevibacilluslaterosporus MTCC 2298. The capability of this strain to detoxifythe textile dyes has been explored earlier [22,23]. The physico-chemical parameters to decolorize dye mixture have beenoptimized along with degradation pattern of structurally differentdyes in the mixture using HPTLC.

2. Experimental methods

2.1. Microorganisms and culture conditions

B. laterosporus MTCC 2298 was obtained from Microbial Type

Culture Collection, Chandigarh, India. The pure culture was main-tained on the Nutrient agar slant containing (g/l): NaCl, 5.0;bacteriological peptone, 5.0; yeast extract, 2.0; beef extract, 1.0 andagar powder 15.0 at 4 ◦C.

rdous

2

GfAHLp

2

Edyc–lstDDd1scipiweDc

A

wt

A

2

as0d

2

m((Tgawb1fic(d

M.B. Kurade et al. / Journal of Haza

.2. Dyes and chemicals

Textile dyes Remazol red, Rubine GFL, Brown 3 REL, Scarlet RR,olden yellow HER, methyl red and Brilliant blue GL were obtained

rom Manpasand Textile Processing Industry, Ichalkaranji, India.ll required chemicals were obtained from Sigma–Aldrich, USA,iMedia Laboratories Pvt. Ltd., Mumbai, India and Sisco Researchaboratory (SRLs), India. All chemicals used were of the highesturity available and of the analytical grade.

.3. Decolorization experiment

Decolorization experiments were performed in the 250 mlrlenmeyer flasks containing 100 ml nutrient broth. Mixture ofyes (Remazol red, Rubine GFL, Brown 3 REL, Scarlet RR, Goldenellow HER, methyl red and Brilliant blue) was added in pre-grownulture (24 h) at a concentration of 70 mg/l (each dye concentration

10 mg/l) and 5 ml of culture medium was withdrawn at regu-ar time intervals. Aliquot was centrifuged (4000 × g for 20 min) toeparate cell mass and clear supernatant was used to determinehe decolorization. Decolorization was analyzed using Americanye Manufacturing Institute (ADMI) 3WL tristimulus method [24].ecolorization was carried out under static as well as shaking con-ition (120 rpm) at 30 ◦C in 250 ml Erlenmeyer flasks containing00 ml nutrient broth. Faster decolorization rate was observed intatic culture, hence further studies were carried out under staticondition. Effect of pH on the decolorization performance was stud-ed by adjusting different pH (3.0, 5.0, 7.0, 9.0 and 11.0) of the 24 hre-grown culture before dye addition. In another set of exper-

ment, effect of temperature on the decolorization performanceas studied by incubating pre-grown (at 30 ◦C) culture at differ-

nt temperatures (10, 20, 30, 40 and 50 ◦C) after the dye addition.ecolorization was expressed in terms of ADMI removal ratio andalculated as follows.

DMI removal (%) = ADMI (to) − ADMI (t)ADMI (to)

× 100

here ADMI (to) and ADMI (t) are the initial ADMI value (at 0 h) andhe ADMI value after a particular reaction time (t), respectively.

All the decolorization experiments were carried out in triplicate.biotic controls (without microorganisms) were always included.

.4. Effect of metal salt on the decolorization of mixture of dyes

The effect of various metal salts, viz., CaCl2, ZnSO4, FeCl3, MgCl2,nd CuSO4 on the decolorization of dye mixture was studied byupplementing them in nutrient medium at the concentration of.5 mM and the influence of metal salt was quantified using percentecolorization in terms of ADMI removal after 24 h incubation.

.5. Effect of carbon and nitrogen sources on decolorization

Different carbon and nitrogen sources (0.5% w/v) were supple-ented in Bushnell Haas medium (BHM) [g/l: MgSO4 (0.2), K2HPO4

1.0), CaCl2 (0.02), FeCl3 (0.05), NH4NO3 (1.0)] along with 0.05%w/v) peptone to determine their effect on the dye decolorization.he tested carbon sources were glucose and starch while nitro-en sources used were yeast extract, peptone, ammonium chloride,nd urea. In addition to this, rice husk, bagasse, wheat bran andood shavings were tested as agricultural waste products. 1 g of

agasse powder, wheat bran and wood shavings were mixed with00 ml distilled water individually, boiled for 15 min, cooled, and

ltered after boiling. The 5% (v/v) ml boiled extract of each agri-ultural waste was used as an inducer with Bushnell Haas mediumBHM). The effect of these substrates was calculated using percentecolorization in terms of ADMI removal after 24 h incubation.

Materials 192 (2011) 1746– 1755 1747

2.6. Preparation of cell free extract

The cells of B. laterosporus were grown for 24 h in nutrient broth.The grown cells were harvested by centrifugation (10,000 × g,20 min at 4 ◦C) and suspended in 50 mM potassium phosphatebuffer (pH 7.4), gently homogenized and sonicated (30 s, 40 ampli-tude, 7 strokes) at 4 ◦C. The sonicated cells were centrifuged(10,000 × g, 15 min at 4 ◦C) and the supernatant was used as sourceof intracellular enzyme. The resulting extract was then used as anenzyme source. The culture supernatant obtained after centrifuga-tion during the harvesting of cell biomass was directly used as asource of extracellular enzymes.

2.7. Determination of enzyme activities

Veratryl alcohol oxidase activity was determined by using theprocedure reported earlier by Jadhav et al. [25]. Tyrosinase enzymeactivity was determined by the formation of o-benzoquinone anddehydro-ascorbic acid in 3 ml reaction mixture containing 50 mMof catechol and 2.1 mM of ascorbic acid in 50 mM potassium phos-phate buffer (pH 6.8) by measuring decrease in optical densityat 265 nm [26]. Riboflavin reductase and NADH-dichlorophenolindophenol (NADH-DCIP) reductase activity was determined usinga procedure reported by Tamboli et al. [21]. Azo reductase assaywas performed as the procedure reported earlier by Telke et al.[27]. Composition of the assay mixture (2.0 ml) was 4.45 �M ofmethyl red (MR), 100 �M NADH, 1.7 ml of potassium phosphatebuffer (20 mM, pH 7.5). The reaction mixture was pre-incubatedfor 4 min followed by the addition of NADH and monitored for thedecrease in color absorbance (430 nm) at room temperature. Thereaction was initiated by addition of 0.1 ml of the enzyme solution.Methyl red reduction was calculated by using its molar extinctioncoefficient of 0.023 �M/cm and activity was expressed in units. Oneunit of enzyme activity was defined as amount of enzyme requiredto reduce 1 �M of substrate/min.

2.8. Extraction and analysis of metabolites obtained after dyedecolorization

Biomass was removed after decolorization by centrifugation(10,000 × g at 4 ◦C for 20 min) and supernatant was processed forextraction of degradation metabolites with equal volume of ethylacetate. The extracted residue were dried over anhydrous Na2SO4and evaporated to dryness in a rotary evaporator. The crystalsobtained were dissolved in small volume of HPLC grade methanoland used for HPLC, FTIR and HPTLC analysis. High performanceliquid chromatography (HPLC) analysis was carried out (Watersmodel no. 2690) using C8 column (symmetry, 4.6 mm × 250 mm)by isocratic elution with 10 min run time. The mobile phase wasmethanol with flow rate of 1 ml/min and UV detector at 530 and260 nm. Decolorization of mixture of dyes was monitored usingUV–vis spectrophotometer (Hitachi U 2800). The Fourier trans-form infrared spectroscopy (FTIR) analysis of extracted metaboliteswas done on Perkin-Elmer, spectrum one instrument and com-pared with control dye in the mid IR region of 400–4000/cm with8 scan speed. The samples were mixed with spectroscopicallypure KBr in the ratio 5:95, pellets were fixed in sample holder,and the analysis was carried out. To confirm the biodegradationof dyes in the mixture, the same metabolites were analyzed byhigh performance thin layer chromatography (HPTLC) using silicagel plates (HPTLC Lichrospher silica gel 60 F254S). 15 �l of sam-ple was applied on the HPTLC plate by micro syringe using sample

applicator (Linomat V, Camag, Switzerland). The solvent system asmobile phase used was toluene:ethyl acetate:methanol (7:2:1). Thetwin trough chamber was saturated with mobile phase for 20 minprior to plate development. After development scanning was done

1 rdous

udoWv

2

amtI

2

Te

3

3

hotoodptAdoAt

748 M.B. Kurade et al. / Journal of Haza

sing TLC scanner (Camag, Switzerland) at 254 and 530 nm usingeuterium and tungsten lamp respectively with slit dimensionf 5 × 0.45 mm. The chromatogram was analyzed by using HPTLCin-CATS 1.4.4.6337 software. The plate was observed in the ultra-

iolet (254 and 366 nm) as well as visible light.

.9. Phytotoxicity study

This test was performed to assess the toxicity of the untreatednd treated samples at a final concentration of 2000 ppm ofetabolites and dye mixture. The test was carried out according

o Parshetti et al. [28], on two kinds of seeds commonly used inndian agriculture; Sorghum vulgare and Phaseolus mungo.

.10. Statistical analysis

Data were analyzed by one-way analysis of variance usingukey–Kramer multiple comparison test. Values are mean of threexperiments. Values were considered significant when P was <0.05.

. Results and discussion

.1. Decolorization experiment

Textile industry effluent generally possesses the variety ofighly complex dyes. Therefore it is necessary to study the capacityf pollutant cleaner microorganism towards decolorization of mix-ure of dyes instead of single dye. The characteristics of mixturef dyes are highly erratic in both, the hues and the concentrationf color and thus it is more complicated to quantify than singleye. Being a complex mixture of dyes, it did not show well definedeaks in the visible region hence the decolorization of the mix-ure of dyes was determined in terms of the ADMI value [24]. TheDMI value provides a true measurement of water color, indepen-

ent of hue and thus opens the way to a more accurate definitionf wastewater and colored mixture [29]. In the present study, thisDMI value was found to be reduced significantly by 87% within

ime span of 24 h in the optimum environment of pH 7, 40 ◦C and at

Fig. 1. Effect of pH and temperature on decoloriz

Materials 192 (2011) 1746– 1755

static condition, with 71 and 49% reduction of COD and TOC respec-tively suggesting biodegradation of dyes to intermediates and notcompletely mineralized into simple forms as CO2, H2O, CH4 andNH3. The negligible decolorization in shaking condition might bedue to the competition of oxygen and the azo compounds for thereduced electron carriers under aerobic condition [30]. Fig. 1 repre-sents the effect of pH and temperature on decolorization of mixtureof dyes. Decolorization of most of the dyes by microbial culturesis generally carried out in neutral to slightly alkaline pH range.Meiying et al. [31] showed the optimum pH range of 6.0–8.0 forfaster decolorization of anthraquinone dye using Shewanella decol-orationis S12. The optimum temperature for rapid decolorizationof dye mixture was found to be 40 ◦C from the studied tempera-ture range (10–50 ◦C). Natural microbial consortium required thesame temperature range for maximum decolorization of mixture ofdyes [19]. The loss of cell viability or the negligible oxido-reductiveenzyme activities might be the cause of reduced decolorization inculture broth which was incubated beyond the optimum temper-ature range. The broad specificity of B. laterosporus to decolorizestructurally different dyes is presented in Table 1.

3.2. Effect of metal salts on decolorization of mixture of dyes

One of the major problem associated with textile processingeffluents are heavy metal ions, which may arise from materials usedin the dyeing process or from metal containing dyes. For this rea-son, this study was carried out to examine the effect of selectedmetals on mixture of dyes decolorizing ability of B. laterosporus. Inthis study, although similar growth of organism was seen in all testflasks, supplementation of Cu2+, Zn2+ and Fe3+ strongly diminishedthe decolorization efficiency; however Ca2+ showed no significantchange when compared to control (Table 2). Biodegradation ofdyes is mostly performed using enzymes in microbial cells [32,33]and the performance of these enzymes may dependent upon the

presence of heavy metal. Metal ions may alter the active site con-firmation which directly leads to the inhibition of enzymes or it maybe utilized as co-factor for that enzyme which fastens the reactivityof the enzyme. During the decolorization of mixture of dyes, induc-

ation of mixture of dyes by B. laterosporus.

M.B.

Kurade

et al.

/ Journal

of H

azardous M

aterials 192 (2011) 1746– 1755

1749

Table 1Decolorization of structurally different textile dyes by B. laterosporus.

Sr. no. Dye name and class Structure �max (nm) Time (h) Decolorization (%) Reference

1 Remazol red (reactive azo)

N

N

N

N

S OO

O-

OH

S

O

O

O-

NH

Cl NHS

O

O

O-

S CH2 CH2 S

O

O-

O

O

ON

530 30 100 This study

2 GY-HER (reactive azo)N N

CH3

OH

NH

Cl

Cl

SO2Na

NaO2S

420 48 87 [23]

3 Scarlet RR (disperse)

N

O

NNH

Cl

CH3

O

N+

O O

470 36 92 This study

4 Rubine GFL (azo) N N

C N

N+

O

O-

N

CH2 CH2 C N

CH2 CH 3530 48 96 This study

5 Methyl red (azo)HO

ONN

NCH3

CH3

440 12 93 [22]

1750M

.B. K

urade et

al. /

Journal of

Hazardous

Materials

192 (2011) 1746– 1755

6 Brilliant blue (disperse)

S-

O

OO

N

CH3

CH3

HN

O

CH3

N+

CH3

S-

O O

O

620 120 55 This study

7 Brown 3 REL (disperse)

N

N

Cl

Cl

O CH2 C

O

NH C

O

NH

440 42 94 This study

M.B. Kurade et al. / Journal of Hazardous

Table 2Effect of different metals and carbon, nitrogen sources on decolorization of mixtureof dyes by B. laterosporus.

Sr. no. Metal % ADMI removal

1 Control 872 CuSO4 103 MgCl2 564 FeCl3 385 CaCl2 906 ZnCl2 32

Carbon/nitrogen source1 Control 262 Glucose 43 Starch 34 Peptone 505 Yeast extract 446 Wood shavings 47 Bagasse 298 Rice husk 239 Wheat bran 3510 Soya waste 30

ttvaZcdat

3m

tasoswdcocsTcwrMmiSbce[ytddt

alkanes. It also showed formation of oximes at 1660/cm. Peak at1512/cm corresponds to N O stretch as in aromatic nitro com-

ion of veratryl alcohol oxidase was observed. It has been reportedhat the presence of Ca2+ in reaction mixture induces the activity oferatryl alcohol oxidase; on the other hand, the existence of Cu2+

nd Zn2+ caused inhibition [25]. Although these metal ions (Cu2+,n2+, Fe3+ and Mg2+) were responsible for reduced decolorization,omplete inhibition was not seen. Telke et al. [34] reported higherecolorization using Ca2+ and inhibitory effects of Mg2+, Cu2+, Fe3+

nd Zn2+ during the decolorization of sulfonated azo dye C.I. Reac-ive orange 16.

.3. Effect of carbon and nitrogen sources on decolorization ofixture of dyes

The rate of dye decolorization/degradation process depends onhe availability and type of a co-substrate used, because it acts asn electron donor for the dye reduction. Some carbon and nitrogenources were chosen to determine their effects on decolorizationf mixture of dyes by B. laterosporus (Table 2). Although there wasimilar growth of this strain in all tested sources, large differenceas observed in the decolorization pattern. There was reducedecolorization in the presence of carbon source as compared to theontrol medium indicating the inhibitory effect of carbon sourcen the enzyme system responsible for decolorization of dyes. Inontrast, amplified decolorization was obtained in the mediumupplemented with nitrogen source like peptone and yeast extract.here was 4 and 3% decolorization in pure carbon sources, glu-ose and starch respectively, whereas 50 and 44% decolorizationas obtained in medium containing peptone and yeast extract

espectively as compared to 26% decolorization in the control flask.eiying et al. [31] reported the lower decolorization rate in theedium containing carbon source such as glucose and sucrose dur-

ng the decolorization of anthraquinine dye using S. decolorationis12. Similarly glucose inhibited the decolorization ability of mixedacterial consortium [35]. Faster decolorization in the mediumontaining yeast extract might be due to the metabolism of yeastxtract, which is considered essential for the regeneration of NADH36]. Similar observations of higher decolorization in presence ofeast extract have been reported by Telke et al. [6]. Some agricul-ural waste as supplementary nutrient also used to decolorize theye. Wheat bran was found to be helpful waste source with 35%

ecolorization of mixture of dyes as compared to 26% decoloriza-ion in control medium.

Materials 192 (2011) 1746– 1755 1751

3.4. Enzymatic analysis

A major mechanism of biodegradation in the microbial cell isbecause of the biotransformation enzymes present in that cell.Data shown in Table 3 represents the enzymatic behavior dur-ing the degradation process of dyes. Enzymatic analysis detectedsignificant induction of veratryl alcohol oxidase, intracellular andextracellular tyrosinase and NADH-DCIP reductase with 1230,1567, 394 and 315% induction respectively at 6 h of decolorizationprogress. At 12 h of decolorization, the induction of veratryl alco-hol oxidase and tyrosinase was found to be reduced, after which itregained the initial induction. But considering overall process, allthe activities of the oxidative enzymes in the test was observed tobe greater than control (0 h). There might be feed-back inhibitionin which the metabolites produced after 6 h might have interferedwith the oxidative enzyme system due to which oxidative enzymescould not maintain the previous level after 6 h, causing cessationin their induction. At the end of 24 h; veratryl alcohol oxidase,intracellular and extracellular tyrosinase managed to retain theirinductive activities at 895, 1534 and 358%. In case of reductiveenzymes; riboflavin reductase activity induced significantly whileazo reductase activity was found to be completely inhibited at first6 h. But at 12 h, 19% increase in the azo reductase activity wasdetected. These enzymes have been reported in the positive roleof biodegradation of textile dyes [20,25,27,6,37].

3.5. Analysis of degradation products

In HPLC analysis, the control showed three distinct peaks at2.494, 2.754 and 3.060 min while this chromatogram was com-pletely altered in case of metabolites of mixture of dyes extractedafter its decolorization (Fig. 2). There were eight peaks in HPLC chro-matogram of metabolites as 1.306, 1.463, 1.849, 2.328, 2.808, 3.005,5.193 and 5.406 min. These variations suggest the biotransforma-tion of mixture of dyes. Significant reduction in the absorbanceof culture supernatant was noticed as compared to the control(data not shown). The general observation can be made that decol-orization is a consequence of the transformation of the molecularstructure of the dye chromophore [38].

The significant difference in the FTIR spectrum of control ofmixture of dyes and metabolites obtained after its decolorizationsuggests the biodegradation of mixture of dyes (Fig. 3). Peaks incontrol spectrum at 1311.16/cm corresponds to C–N vibration,3452.70/cm for O–H stretch, presence of alkanes (cyclopropane) at2863.90, 2970.48 and 3069.33/cm. Peak at 638.46 and 687.16/cmshowed presence of halides (C–Cl) (this peak corresponds to cyclo-propane ring structure containing alkane and halides in Goldenyellow HER in dye mixture). The peak at 2250.52/cm correspondsto saturated nitriles in Rubine GFL, whereas the peak at 1599.04/cmcorresponds to N N stretch in azo dyes present in dye mixture. Thepeak at 1713.33/cm showed C O stretch (present in the Brown3 REL and Scarlet RR dye in dye mixture), the peaks at 765.77 to993.86/cm showed C–H deformation. The presence of NO2 stretchat 1486.20 and 1552.27/cm corresponds to the aromatic nitro com-pound. N–H deformation was observed at the peak of 1524.78/cm,S O stretch was observed at the peak 1042.56, 1147.68, 1199.28and 1228.21/cm (supports the dye mixture contains the sulfonateddye compound), the peak at 1116.34/cm showed the presence ofsecondary alcohol. The FTIR spectrum of metabolites extracted afterdecolorization of mixture of dyes showed peaks at 3429/cm corre-sponding to the O–H stretch as in alcohols, C O stretch as in ketonesand N–H stretch in primary amines, 2960/cm for C–H stretch as in

pounds, N–H deformation in acyclic amides, 1455/cm for C–Hdeformation in alkanes, 1337/cm for O–H deformation as in phe-

1752 M.B. Kurade et al. / Journal of Hazardous Materials 192 (2011) 1746– 1755

Table 3Enzyme activities of control cells and cells obtained after decolorization of mixture of dyes.

Enzyme 0 h 6 h 12 h 18 h 24 h

Veratryl alcohol oxidasea 0.237 ± 0.012 3.153 ± 0.058** 0.835 ± 0.046** 1.406 ± 0.089** 2.359 ± 0.208**

Tyrosinaseb

Intracellular 358.0 ± 25.0 5969.0 ± 484.0** 3426.0 ± 92.0** 2843.0 ± 68.0** 5850.0 ± 75.0**

Extracellular 579.0 ± 62.0 2862.0 ± 214.0** 568.0 ± 10.0 435.0 ± 8.3** 2652.0 ± 112.0**

NADH DCIP reductasec 62.67 ± 4.64 259.9 ± 11.08** 265.4 ± 5.83** 437.1 ± 9.95** 290.7 ± 8.67**

Azo reductased 2.869 ± 0.398 NA 3.416 ± 0.564* 2.315 ± 0.281* NARiboflavin reductasee NA 2.759 ± 0.059** 2.055 ± 0.094** 2.510 ± 0.093** 3.254 ± 0.189**

Values are mean of three experiments ±SEM.* Significantly different from control cells at P < 0.01 by one-way ANOVA with Tukey–Kramer comparison test.

** Significantly different from control cells at P < 0.001 by one-way ANOVA with Tukey–Kramer comparison test.a Enzyme unit’s min−1 mg protein−1.b Activity in IU.

n1mamcTtsts

[ot

c �g of DCIP reduced min−1 mg protein−1.d �M of methyl red reduced min−1 mg protein−1.e �g of riboflavin reduced min−1 mg protein−1.

ols and tertiary alcohols and CN vibrations in aromatic amines,306/cm for C–N as in aromatic primary amines and C–H defor-ation in alkenes, 1112/cm for C–OH stretch in secondary alcohols

nd C–O stretch as in phenols. 915/cm for C–H deformation as inonosubstituted alkenes and 702/cm for C–Cl stretch as in poly-

hlorinated compounds and C–S stretch as in sulphur compounds.he disappearance of peak at 1599/cm confirmed the reduction ofoxic azo groups in the dyes. The significant change in the FTIRpectrum of metabolites compared to control spectrum suggesthe biotransformation of complex dyes present in the mixture intoimple form.

There are few reports on biodegradation of mixture of dyes18,20], but it did not propose the mechanistic way of the through-ut process by which it was degraded. To reveal this mechanism,he analysis of degradation process by HPTLC was crucial, which

Fig. 2. HPLC profile of control dyes mixture (A) and m

disclosed the progressive pattern of biodegradation of mixture ofdyes (Fig. 4). In first 6 h, seven dyes were degraded into 15 metabo-lites as 15 bands were obtained in that lane. Further, mineralizationof these intermediates led into the formation of total 17, 18 and 19metabolites in 12, 18 and 24 h respectively. Within first 6 h, thesubstrate priority of oxido-reductive enzyme system present in B.laterosporus was Golden yellow HER, Rubine GFL, Scarlet RR, methylred and Brown 3 REL as the color capability of all these dyes waslost within first 6 h except Remazol red and Brilliant blue GL. It doesnot mean that Remazol red and Brilliant blue were inaccessibleto the enzymatic machinery; because at 6 h, the color intensity of

Remazol red was reduced considerably. After 6 h Remazol red wascompletely mineralized. But in case of Brilliant blue; being recalci-trant and complex nature, it might be very tough for enzyme systemto break this molecule into simple form. But it succeeded to remove

etabolites obtained after its decolorization (B).

M.B. Kurade et al. / Journal of Hazardous Materials 192 (2011) 1746– 1755 1753

) and m

omfafBg2soabH(fihswrdT

Fig. 3. FTIR spectrum of control dyes mixture (A

ne or more strongly polar groups (sulphonic) other than chro-ophoric group from parent structure, as its Rf value was changed

rom 0.0 to 0.37 after 6 h (blue colored band in 12, 18 and 24 h). Inddition to this, the intensity of this band was gradually increasedurther at 18 and 24 h, suggesting the continuous conversion of therilliant blue GL dye into that unique metabolite by losing its polarroup. Further breakdown of this metabolite was not possible up to4 h. Desulphonation by veratryl alcohol oxidase from Comamonasp. UVS has been reported earlier [25]. The structural complexityf Brilliant blue and Remazol red might be acted like a barrier forccessibility of the enzyme system. Except metabolite of Brilliantlue, entirely noncolored and fluorescent nature of the bands onPTLC plate visualized at 254 and 366 nm and change in Rf values

data not shown) confirmed the removal of chromophoric groupsrom the parent dyes and breakdown of these complex speciesnto simple form. As represented in Table 1, much longer timeave been needed to decolorize each dye varied according to theirtructures; but in the case of mixture of dyes, 87% decolorization

as achieved within only 24 h. Methyl red and Golden yellow HER

equired 12 and 48 h respectively for complete breakdown whenecolorized individually in the culture of B. laterosporus [22,23].he activities of oxidative enzymes monitored in the cells during

etabolites obtained after its decolorization (B).

decolorization of mixture of dyes were found to be higher thantheir levels during the single dye decolorization (data not shown).This could be the most possible reason for the faster decoloriza-tion of the dyes within the mixture. From the HPTLC data, it can beconcluded that the same enzyme may act on more than one sub-strate (dye) at the same time, as within first 6 h, significant changesoccurred in five dyes. In this mixture, one of the dyes might havebeen acted like redox mediator for the other dye which ultimatelycaused quicker substrate utilization by the oxido-reductiveenzymes.

3.6. Phytotoxicity

This study is of particular relevance since the Panchganga Riverand Ichalkaranji area near Kolhapur, India are heavily industrial-ized, with significant wastewater discharge from textile industriesto the environment which causes the harmful impacts on thenearby flora and fauna. Table 4 represents the toxicity analysis of

the mixture of dyes and its metabolites obtained after decoloriza-tion. The toxic nature of the dyes in the mixture before treatmentwas confirmed when phytotoxicity study that showed the 60 and40% germination inhibition of S. vulgare and P. mungo respectively.

1754 M.B. Kurade et al. / Journal of Hazardous Materials 192 (2011) 1746– 1755

Fig. 4. HPTLC plate at different wavelength (A) and HPTLC chromatogram at 254 nm (B).

Table 4Phytotoxicity study of mixture of dyes and the metabolites obtained after its decolorization.

Observations Sorghum vulgare Phaseolus mungo

I II III I II III

Germination (%) 100 40 90 100 60 100Shoot length (cm) 3.25 ± 0.73 1.10 ± 0.16* 2.90 ± 0.34** 15.22 ± 0.91 7.16 ± 0.29* 13.43 ± 0.35**

Root length (cm) 7.96 ± 0.55 1.52 ± 0.23* 3.86 ± 0.32** 3.45 ± 0.21 1.79 ± 0.23* 3.64 ± 0.21**

I: seeds germinated in distilled water; II: seeds germinated in mixture of dyes; III: seeds germinated in metabolites obtained after decolorization of mixture of dyes.Data was analyzed by one way analysis of variance (ANOVA) with Turkey–Kramer multiple comparison test using mean values of germinated seeds of three experiments.

ermineeds g

Bacbot

* Seeds germinated in mixture of dyes are significantly different from the seeds g** Seeds germinated in degradation products are significantly different from the s

ut after treatment with B. laterosporus, germination of S. vulgarend P. mungo was enhanced by 125 and 66% respectively whenompared to untreated dyes. Reduction in shoot and root length of

oth the plants was observed due to toxicity of untreated mixturef dyes. A significant improvement in shoot and root length of bothhe plants was observed in the treated dye sample.

ated in plain distilled water at P < 0.001.erminated in mixture of dyes at P < 0.01.

4. Conclusion

Enzymatic studies indicate the involvement of veratryl alcoholoxidase, tyrosinase and NADH-DCIP reductase in biotransformationof dyes present in the mixture. The activity of oxidative enzymesin cellular organization amplifies during the stress of structurally

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M.B. Kurade et al. / Journal of Haza

ifferent dyes. B. laterosporus can progressively degrade the dyeithin short span when combined with other dyes in culture;

therwise it needed long time to degrade the same dye when itoncerned about single dye decolorization. B. laterosporus haveroad specificity as it can degrade structurally diverse range ofyes with significant reduction in its toxicity and have preference

n metabolism. B. laterosporus decolorized the mixture of dyes with7% ADMI removal, 71% COD and 49% TOC reduction, hence, it coulde applied in the effluent treatment plant containing different dyes.

cknowledgement

M.B. Kurade and T.R. Waghmode wish to thank Departmentf Biotechnology, New Delhi, India for providing Junior Researchellowship.

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