Licensee OA Publishing London 2013. Creative Commons Attribution License (CC-BY)

Shah MP, Patel KA, Nair SS, Darji AM. Molecular characterization and optimization of Azo dye degrading Bacillus subtillis ETL-2013. OA Molecular & Cell Biology 2013 Apr 01;1(1):2.

Com

petin

g in

tere

sts:

non

e de

clar

ed. C

onfli

ct o

f int

eres

ts: n

one

decl

ared

. Al

l aut

hors

con

trib

uted

to c

once

ptio

n an

d de

sign,

man

uscr

ipt p

repa

ratio

n, re

ad a

nd a

ppro

ved

the

final

man

uscr

ipt.

Al

l aut

hors

abi

de b

y th

e As

soci

atio

n fo

r Med

ical

Eth

ics (

AME)

eth

ical

rule

s of d

isclo

sure

.

Molecular characterization and optimization of

Azo dye degrading Bacillus subtillis ETL-2013

Shah MP*, Patel KA, Nair SS, Darji AM

Industrial Waste Water Research Laboratory,

Division of Applied & Environmental Microbiology Lab

Enviro Technology Limited, Plot No: 2413/14,

GIDC, Ankleshwar-393002

Gujarat, India

Corresponding author. Maulin P Shah, Industrial Waste Water Research Laboratory, Applied &

Environmental Microbiology Lab, Enviro Technology Limited (CETP), Ankleshwar- 393 002, Gujarat, India, Tel:

+91-90 999 65504, Fax No: +91-2646-250707, E-mail: shahmp@uniphos.com,

1) Abstract:

Triphenylmethane dyes belong to the most important group of synthetic colorants and are used

extensively in the textile industries for dying cotton, wool, silk, nylon, etc. They are generally

considered as the xenobiotic compounds, which are very recalcitrant to biodegradation. The

organism isolated from contaminated sites of textile industry soil sample that degrade bromo

phenol blue was subjected to 16s rRNA and the organism was characterized as Bacillus subtilis.

It was able to decolorize 88% of bromo phenol blue dye (5 mg/l) within 12hr at optimized

conditions pH 6, temperature 40ºC, and carbon source (glucose).

Keywords: Bromo phenol blue, decolorization, Bacillus subtilis ETL-1982

2) Introduction

Rapid industrialization has necessitated the manufacture and use of different chemicals in day to

day life (Shah et al., 2013). Reactive dyes, including many structurally different dyes, are

extensively used in the textile industry because of their wide variety of color shades, high wet

fastness profiles, ease of application, brilliant colors, and minimal energy consumption. The

three most common groups are azo, anthraquinone and phthalocyanine (Shah et al., 2013). A

major class of synthetic dyes includes the azo, anthroquinone dyes. Approximately 10,000

different dyes and pigments are used industrially and over 0.7 million tons of synthetic dyes are

produced annually worldwide. Azo dyes belong to the most important group of synthetic

colorants and are used extensively in the textile industries for dying cotton, wool, silk, nylon, etc.

Azo dyes, which are aromatic compounds with one or more(–N=N–) groups, accounts for the

majority of all textile dyestuffs produced and have been the most commonly used synthetic dyes

in the textile, food, paper making, printing, leather and cosmetic industries. They are generally

considered as the xenobiotic compounds, which are very recalcitrant to biodegradation (Zollinger,

1991; Stolz, 2001). During the dyeing processes about 10–90% of the dye stuff do not bind to the

fibers and therefore, released into the sewage treatment system or the environment (Zollinger

1991; Abdullah, et al., 2000). The inefficiency in dying process has resulted in 10-15% of

unused dye stuff entering the wastewater directly. Improper textile dye effluent disposal in

aqueous ecosystems leads to the reduction in sunlight penetration which in turn decreases

photosynthetic activity, dissolved oxygen concentration, water quality and depicts acute toxic

effects on aquatic flora and fauna, causing severe environmental problems worldwide. In

addition to their visual effect, triphenylmethane dyes and their metabolites are toxic,

carcinogenic, mutagenic, leading to potential health hazard to humankind Removal of hazardous

industrial effluents is one of the growing needs of the present time (Mittal et al. 2005). Dyes are

synthetic aromatic water-soluble dispersible organic Colorants, having potential application in

various industries (Mohan et al. 2002). The dye degradation of physical/chemical methods has

inherent drawbacks: economically unfeasible (more energy and chemicals), unable to remove the

recalcitrant triphenylmethane dyes and their organic metabolites completely from such effluents,

generate a significant amount of sludge may cause secondary pollution problems, substantially

increases the cost of these treatment methods and complicated procedures. Biological

degradation, being inexpensive and ecofriendly, is considered a valuable removal method for

many toxic pollutants. Several microorganisms, including a number of bacteria, yeast, and fungi,

have been investigated for their ability to biodecolorize triphenylmethane dyes (Wamik et al.

1988; Sani and Banerjee 1999; Cha et al. 2001; An et al. 2002; Liu et al. 2004; Jadhav and

Govindwar 2006; Ren et al. 2006b; Utkarsha et al. 2008). Biochemical studies of the

decolorization process indicate that laccase, peroxidase, and lignin peroxidase and

triphenylmethane reductase (TMR) are involved in the enzymatic decolorization of dyes (Tekere

et al. 2001; Shin and Kim 1998; Jang et al. 2005). This study aims to investigate the potential of

Bacillus subtillis isolated from textile soil for decolorizing a solution containing a bromo phenol

blue dye at various optimized conditions like temperature, pH, and carbon source and dye

concentration.

3) Experiment

3.1) Sample collection: Soil sample was collected from contaminated sites of Ankleshwar

textile industry, Ankleshwar, Gujarat, India. The collected soil sample was placed in a sterile

polythene bags and stored at 4ºC.

3.2) Isolation of bacteria: The soil sample collected from textile industry was subjected to

serial dilution (Madigan et al. 2000). From that 10-5 and 10-6 dilutions are plated on Nutrient agar

(peptone 5g/l, NaCl 5g/l, yeast extract 1.5g/l, beef extract 1.5g/l, agar20g/l temperature-25ºC,

pH7.4±0.2) by spread plate method. The single isolated colonies were sub cultured on LB (casein

enzymatic hydrolysate-10g/l, yeast extract-5g/l, sodium chloride-10g/l, agar-15g/l and ph(at

25)7.5±0.2) agar plates. And pure culture was maintained on LB (Luria Bertani) agar plates.

3.3) Dye Degradation assay: The purified two colonies from 10-5 dilution were streaked on the

screening media (Prepare 30ml LB agar medium and 10mg/lit bromophenolblue dye was added

to medium) and were kept for incubation for 2days for preliminary identification of dye

degrading bacteria. A control plate was also maintained for comparison.

3.4) Genomic DNA isolation: Method described by Krsek and Wellington (1999), with some

modifications. The enzymatic, chemical and mechanical lysis was performed. For the enzymatic

lysis, the overnight grown culture was suspended in 2 ml of a lysozyme solution (150mM NaCl,

100 mM EDTA, 5 mg/ml lysozyme). The tube was inverted several times to mix the contents

and placed in a 37°C water bath for 2hr. Subsequently, 500µl proteinase K (2.5mg/ml) was

added, and the contents of the tube gently mixed and placed in a 55°C water bath for 15min. For

the chemical lysis, 2ml SDS solution (500 mM NaCl, 500 mM Tris-HCl, 10% w/v SDS) was

added to the tube and the contents mixed by inversion several times for 5 min. Finally, for the

mechanical lysis process, the samples were subjected to three cycles of freezing in liquid

nitrogen and thawing at 100ºC. An equivalent volume (4.5ml) of phenol- chloroform-isoamyl

alcohol (25:24:1) was added and the mixture gently vortexed for 1 min. To precipitate DNA, 0.7

vol. cooled isopropanol and 1/10 vol. 3M sodium acetate were added to the supernatant. The

mixture was gently mixed (5-10 times) and kept at -20°C overnight. The sample was centrifuged

at 4100rpm for 10min, and the pellet was washed three times with cold 70% ethanol and then

resuspended in 100µlTE buffer 10/0.1 (10 mM Tris-HCl, 0.1 mM EDTA, pH 8.0). For

visualizing the DNA extract, 10µl of each extract was electrophoresed on 1% agarose gel in 1X

TAE buffer, which were then stained with ethidium bromide and examined under UV light.

3.5) Strain characterization: The Genomic DNA was isolated from the overnight grown

bacterial culture was amplified with universal bacterial primers. The 25μl of reaction mixture

contains, 15µl of master mix (10X assay buffer, DNTP’s, Taq), 1µl of forward primer

(GGCGAACGGGTGAGTAA), 1µl of reverse primer (ACTGCTGCCTCCCGTAG), 2µl of

template DNA and 6µl of distilled water. PCR was carried out by the thermal cycler PCR under

the following conditions-- 30 cycles of 94ºC for 4min, 94ºC for 1min, 52ºC for 1min, 72ºC for

1.15min followed by final extension at 72ºC for 1min and holding temperature at 10ºC for 1min.

The amplified DNA fragments were observed by agarose gel electrophoresis in 2% agarose gel

and sequenced. The unknown bacterium was identified using GenBank database. Partial 16S

rRNA sequences obtained from isolates were assembled in a contig using the

phred/Phrap/CONSED program (Godon et al., 1997; Ewing et al., 1998). Identification was

achieved by comparing the contiguous 16S rRNA sequences obtained with the 16S rRNA

sequence data and type strains available in the public databases GenBank using the BLASTn

sequence match routines. The sequences were aligned using the CLUSTAL X program and

analyzed with the MEGA software 2001 (Thompson et al., 1994; Kumar et al., 2004).

Evolutionary distances were derived from sequence-pair dissimilarities, calculated as

implemented in MEGA, using Kimura’s DNA substitution model (Kimura, 1980). The

phylogenetic reconstruction was done using the neighbourjoining (NJ) algorithm, with bootstrap

values calculated from 1000 replicate runs, using the routines included in the MEGA software

(Saitou and Nei, 1987).

3.6) Decolorization experiments: Decolorization of dyes was determined as relative in decrease

in absorbance for each dye at their absorbance maximum at particular time interval. A loop full

of microbial culture (10µl) was inoculated in 250ml of Erlenmeyer flask containing 100ml LB

broth and incubated at 30ºC for 24hr. After 24hr of incubation, bromo phenol blue dye was

added at concentration of 5 mg/l and 3ml of the culture media was withdrawn at different time

intervals. Aliquot was centrifuged at 8000rpm for 10 minutes to separate the bacterial cell mass,

clear supernatant was used to measure the decolorization at the absorbance maxima of the

respective dyes. Abiotic controls (without microorganism) were always included. (Saratale, G.D,

Kalme, S.D and Govindwar, S.P, 2006). Controls were carried out in the same conditions but

without dyes or inoculum. Concentration of dyes were selected which obeys lamberts- beer law

The percentage decolorization was calculated as follows

% Degradation = Initial absorbance - Observed absorbance

_________________________________×100

Initial absorbance

3.7) Effect of initial dye concentration on decolorization: The decolorization of bromo phenol

blue blue was studied at different concentrations of the dye (5, 10, 20, 30, 40, 50 and 60 mg/l) as

described by (El-Naggar et al. 2004). The bromo phenol blue blue dye was added separately to

150 ml Erlenmeyer flasks containing 50 ml LB medium. The flask was inoculated with 1ml

Bcillus subtilis ETL-2013 (isolated from Industrial Textile soil). Dye absorbance was measured

by spectrophotometer at 620nm.

3.8) Effect of pH on decolorization: The effect of pH on bromophenolblue dye decolorization

was studied by performing the experiment at different pH values from pH 6-pH 10 using 1M

NAOH and 1M HCL and at 620nm absorbance was measured by spectrophotometer (Oranusi

and Ogugbue 2005).

3.9) Effect of Temperature on decolorization: For studying the effects of temperature on

decolorization of bromophenolblue dye, the bacterial culture was incubated at different

temperatures from 10ºC-40ºC. After 24hrs incubation the bromophenolblue dye was added

separately to cultures. Dye absorbance was measured by spectrophotometer at 620nm.

3.10) Effect of carbon source on decolorization: For enhancing the decolorization of

bromophenolblue was observed by addition of different carbon sources to culture media. The

rate of decolorization was decreased with increasing concentration of dye. Absorbance was

measured by spectrophotometer at 620nm (Oranusi, N.A. and Mbah, A.N. 2005).

4) Results and Discussion:

4.1) Dye Degradation assay:

In LB agar medium, after two days incubation the plates were observed for decolorization of the

dye at and around the culture position on comparison with control.

4.2) Genomic DNA isolation:

Result of isolated genomic DNA from dye industry. Agarose gel electrophoresis of genomic

DNA. Lanes 1 and 2 were isolated Genomic DNA from dye industry sample.

4.3) Effect of initial dye concentration on decolorization:

The decolorization of bromophenolblue was studied at various increasing concentration of dye

i.e. from 5, 10, 20, 30, 40, 50, 60 mg/L. The rate of decolorization was decreased with increasing

concentration of dye. 61% decolorization of bromophenolblue was observed at 5mg/lt,

concentration within 24 h respectively. Only 51%, 45%,36%,28%,23% and 22% decolorization

was observed at 10, 20, 30, 40, 50, 60 mg/L dye concentration respectively.

4.4) Effect of PH on decolorization: The effect of pH on dye decolorization was studied by

performing the experiment at pH6- pH10. 68% decolorization of BPB was observed at Ph8,

within 12h respectively. Only 64%. 63%, 60% and 39% decolorization was observed at ph

8,10,9 and 7 dye concentration respectively.

4.5) Effect of Temperature on decolorization: For studying the effects of temperature on

decolorization of Bromophenolblue the culture was incubated at 10-40ºC. 88% decolorization of

bromophenolblue was observed at 40ºC, concentration within 12h respectively. Only 83%, 52%,

27% and no decolorization was observed at temperature dye concentration respectively.

4.6) Effect of carbon source on decolorization: In LB medium, only 70% decolorization of

BPB was observed in 24 h. In an attempt to enhance decolorization performance with extra

supplements of carbon source in LB medium, found 92% decolorization was observed with

glucose in 12h. Only 58% and 46% decolorization observed at fructose and starch respectively.

Conclusions:

The present research indicates the potential aspects of Bacillus subtilis to 88% decolorize and

degrade bromophenol blue. The culture has ability to decolorize bromophenolblue in optimizing

the various parameters like pH, temperature and initial dye concentration within less time, which

is significant for its commercial and industrial application. The results showed that the

decolorization depend on dye concentration, pH and temperature. These results indicate that the

color removal by Bacillus subtilis may be largely attributed to biodegradation. Overall results

suggested the ability of Bacillus subtilis for the decolorization of azo dye and ensured the

ecofriendly degradation of bromophenolblue. It is proposed that Bacillus subtillis ETL-1982 has

a remarkable practical application potential in the biotransformation of various dye effluents

Acknowledgements: Authors are highly thankful to the management of Enviro Technology

Limited., Ankleshwar, Gujarat, India for allowing us to carry our such a noble work for the

sustainable environment.

References:

1. Ajibola, V.O., Oniye, S.J., Odeh, C.E., Olugbodi, T., Umeh, U.G., 2005. Biodegradation

of Indigo containing textile effluent using some strains of bacteria. Journal of Applied

Sciences 5, 853–855.

2. An SY, Min SK, Cha IH, Choi YL, Cho YS, Kim CH, Lee YC(2002) Decolorization of

triphenylmethane and azo dyesby Citrobacter sp. Biotechnol Lett 24:1037–1040.

doi:10.1023/A:1015610018103

3. Cha CJ, Doerge DR, Cerniglia CE (2001) Biotransformation of malachite green by the

fungus Cunninghamella elegans. Appl Environ Microbiol 67:4358–4360.

doi:10.1128/AEM.67.9.4358- 4360.2001

4. El-Naggar MA, El Aasar SA, Barakat KI (2004) Bioremediation of crystal violet using

air bubble bioreactor packed with Pseudomonas aeruginosa. Water Res 38:4313–4322.

doi:10.1016/j.watres. 2004.06.034

5. Ewing, B., Hillier, L., Wendl, M., Green, P., 1998. Basecalling of automated sequencer

traces using phred. I. Accuracy assessment. Genome Research 8, 175–185.

6. Franciscon Elisangela, Zille Andrea, Dias Guimaro Fabio, Ragagnin de Menezes

Cristiano,Durrant Lucia Regina, Cavaco-Paulo Artur .Biodegradation of textile azo dyes

by a facultative Staphylococcus arlettae strain VN-11 using a sequential

microaerophilic/aerobic process.

7. G.K. Parshetti, A.A. Telke, D.C. Kalyani, S.P. Govindwar. Decolorization and

detoxification of sulfonated azo dye methyl orange by Kocuria rosea MTCC 1532.

8. Ganesh Parshetti, Satish Kalme, Ganesh Saratale, and Sanjay Govindwar. Biodegradation

of Malachite Green by Kocuria rosea MTCC 1532

9. Godon, J.J., Zumstein, E., Dabert, P., Habouzit, F., Moletta, R.,1997. Molecular

microbial diversity of an anaerobic digestor as determined by small-subunit rDNA

sequence analysis. Applied and Environmental Microbiology 63, 2802–2813.

10. Jadhav JP, Govindwar SP (2006) Biotransformation of malachite green by

Saccharomyces cerevisiae MTCC 463.Yeast 23:315–323. doi:10.1002/yea.1356

11. Kimura, M., 1980. A simple method for estimating evolutionary rates of base

substitutions through comparative studies of nucleotide sequences. Journal of Molecular

Evolution 16, 111– 120.

12. Kumar, S., amura, K., Nei, M., 2004. MEGA 3: integrated software for molecular

evolutionary genetics analysis and sequence alignment. Briefings in Bioinformatics 5,

150–163.

13. Lamia Ayed, Kamel Chaieb, Abdelkarim, Cheref Amina Bakhrouf Biodegradation of

triphenylmethane dye Malachite Green by Sphingomonas paucimobilis.

14. Liu W, Chao Y, Yang X, Bao H, Qian S (2004) Biodecolorization of azo, anthraquinonic

and triphenylmethane dyes by white-rot fungi and a laccase-secreting engineered strain. J

Ind Microbiol Biotechnol 31:127–132. doi:10.1007/s10295-004-0123-z

15. M.T. Madigan, J.M. Martinko, J. Parker, Brock Biology of microorganisms, Ed. Prentice

Hall, New Jersey, USA (2000).

16. Maulin P Shah, Kavita A Patel, Sunu S Nair, Darji AM (2013) Optimization of

Environmental Parameters on Microbial Degradation of Reactive Black Dye. Journal of

Bioremediation & Biodegradation 4:3

17. Maulin P Shah, Kavita A Patel, Sunu S Nair, Darji AM (2013) Bioremoval of Azo dye

Reactive Red by Bacillus spp. ETL-1982. Journal of Bioremediation & Biodegradation

4:3

18. Mittal A, Kurup L, Gupta VK (2005) Use of waste materials bottom ash de-oiled soya, as

potential adsorbents for the removal of Amaranth from aqueous solution. J Hazard Mater

B 117:171–178. doi:10.1016/j.jhazmat.2004.09.016

19. Mohan SV, Roa CN, Prasad KK et al (2002) Treatment of simulated reactive yellow 22

(Azo) dye effluents using Spirogyra species. Waste Manag 22:575–582. Doi:

10.1016/S0956-053X (02)00030-2.

20. Oranusi, N.A. and Ogugbue, C.J. 2005. Effect of pH and nutrient starvation on

biodegradation of Azodyes by Pseudomonas sp. J. Appl. Sci. Environ. Manage. 9(1):39-

43.

21. Oranusi, N.A.and Mbah, A.N. 2005. Utilization of Azo and Triphenylmethane dyes as a

sole source of carbon, energy and nitrogen by Bacillus sp. African J.Appl. Zool. Environ.

Biol. 7: 87- 94.

22. Rania M.A.Abedin. Decolorization and Biodegradation of Crystal Violet and Malachite

Green by Fusarium solani (Martius) Saccardo. A Comparative study on Biosorption of

Dyes by the Dead Fungal Biomass.

23. Ren SZ, Guo J, Zeng GQ, Sun GP (2006b) Decolorization of triphenylmethane, azo, and

anthraquinone dyes by a newly isolated Aeromonas hydrophila strain. Appl Microbiol

Biotechnol 72:1316–1321. doi:10.1007/s00253-006-0418-2

24. Saitou, N., Nei, M., 1987. The neighbor-joining method: a new method for reconstructing

phylogenetic trees. Molecular Biology and Evolution 4, 406–425.

25. Sani RK, Banerjee UC (1999) Decolorization of triphenylmethane dyes and textile and

dye- stuff effluent by Kurthia sp. Enzyme Microb Technol 24:433–437.

doi:10.1016/S0141- 0229(98)00159-8

26. Saratale, G.D., Kalme, S.D. and Govindwar, S.P., 2006. Decolourization of textile dyes

by Aspergillus ochraceus. Ind. J. Biotechnol., 5, 407–410.

27. Stolz, A., 2001. Basic and applied aspects in the microbial degradation of azo dyes.

Applied Microbiology and Biotechnology 56, 69–80.

28. Tekere, M., Mswaka, A.Y., Zvauya, R., Read, J.S., 2001. Growth, dye degradation and

ligninolytic activity studied on Zimbabwean white rot fungi. Enzyme Microbial

Technology 28, 420–426.

29. Thompson, J.D., Higgins, D.J., Gibson, T.J., 1994. CLUSTAL W: improving the

sensitivity of progressive multiple sequence alignment through sequence weighting,

position specific gap penalties and weight matrix choice. Nucleic Acids Research 22,

4673–4680.

30. Utkarsha S, Rhishikesh D, Jyoti J (2008) Biodegradation of triphenylmethane dye cotton

blue by Penicillium ochrochloron MTCC 517. J Hazard Mater 157:472–479.

doi:10.1016/j.jhazmat.2008.01.023

31. Wamik A, Rajesh KS, Uttam CB (1988) Biodegradation of triphenylmethane dyes.

Enzyme Microb Technol 22:185–191

32. Zollinger, H., 1991. Color Chemistry: Syntheses, Properties and Applications of Organic

Dyes and Pigments, second ed. VHC Publishers, New York.

Figure 1: OA G1.TIF

Figure 2: OA G2.1.TIF

Figure 3: OAG2.TIF

Figure 4: OA.G3.TIF

Figure 5: OA.G4.TIF

Figure 6: OA.G5.TIF

Figure 7: OA.G6.TIF