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Journal of Environmental Biology �January, 2009�
© Triveni Enterprises, Lucknow (India) J. Environ. Biol.
ISSN : 0254-8704 30(1), 89-92 (2009)
http : //www.jeb.co.in [email protected]
A simple method to screen for azo-dye-degrading bacteria
M.A. Syed, H.K. Sim, A. Khalid and M.Y. Shukor*
Department of Biochemistry, Faculty of Biotechnology and Biomolecular Sciences,
University Putra Malaysia, 43400 Serdang, Selangor, Malaysia
(Received: December 18, 2007; Revised received: June 10, 2008; Accepted: June 20, 2008)
Abstract: A stab-culture method was adapted to screen for azo dyes-decolorizing bacteria from soil and water samples. Decolorized azo
dye in the lower portion of the solid media indicates the presence of anaerobic azo dyes-decolorizing bacteria, while aerobic decolorizing
bacteria decolorizes the surface portion of the solid media. Of twenty soil samples tested, one soil sample shows positive results for the
decolourisation of two azo dyes; Biebrich scarlet (BS) and Direct blue 71 (DB) under anaerobic conditions. A gram negative and oxidase
negative bacterial isolate was found to be the principal azo dyes degrader. The isolate was identified by using the BiologTM identification
system as Serratia marcescens.
Key words: Azo dyes, Stab-culture, Screening method
PDF of full length paper is available with author (*[email protected])
Introduction
Many of the known bacterial isolates that could decolorize
azo dyes can only do so under static or anaerobic conditions. In
addition, growth and decolorizing ability are often a two-stage
process. Here, we report on a simple method to screen for azo dye
decolorizing bacteria in soil and water samples. Since the first
commercial synthetic dye, Mauveine, was discovered in 1856, more
than 100 000 dyes have been generated worldwide with an annual
production in excess of 7 x 105 metric tonnes (Zollinger, 1987). Azo
dyes are the largest class of commercially available synthetic dyes
and have found wide applications in textiles, food, cosmetics, plastic,
laboratories, leather, paper printing, colour photography,
pharmaceutical and toy industries (Chung, 1983; Garrigós et al.,
2002; Mathur and Bhatnagar, 2007; Pant et al., 2008; Laowansiri et
al., 2008). Their widespread use, especially in textile and dyestuff
industries, coupled with the fact that azo dyes are not readily degraded
in conventional aerobic treatment systems makes this class of
xenobiotic a significant environmental problem. Until today there is
still no effective and economical treatment of azo dyes that contain
effluents. The employment of physico-chemical methods, although
effective, is cost prohibitive and often lead to extra solid wastes. As a
viable alternative, biological processes have received increasing
interest owing to their cost effectiveness (Banat et al., 1996). Over
the past decades, many microorganisms that are capable of
degrading azo dyes, including bacteria (Haug et al., 1991), fungi
(Demir et al., 2007; Singh et al., 2007), yeasts (Banat et al., 1996),
actinomycetes (Zhou and Zimmerman, 1993) and algae (Jinqi and
Houtian, 1992) have been documented.
The anaerobic reductive cleavage of azo bonds is often
always favoured over the aerobic condition as oxygen molecules
will compete with the azo group for electrons in the oxidation of
reduced electron carrier, i.e. NADH (Banat et al., 1996). The low
specificity of azoreductase often involves low molecular weight redox
mediators (e.g., flavins or quinones) and allows for unspecific
reduction, thus providing a wider range of dyes reduction (Keck et
al., 1997). Many bacteria have been reported to readily decolourise
azo dyes under anaerobic conditions. Bacteroides sp., Eubacterium
sp., Clostridium sp., Proteus vulgaris and Streptococcus faecalis
(Bragger et al., 1997; Rafii et al., 1990). However, the aerobic
decolourisation of azo dyes can also be carried out in the presence
of external carbon sources and, presumably, does not use azo dyes
as the sole carbon or energy sources (Padmavathy et al., 2003).
Zissi et al. (1997) had reported on the cometabolic reductive cleavage
of p-aminoazobenzene to aniline during aerobic growth on glucose.
The reductive decolourisation of sulfonated azo dyes by Bacillus
sp., Pseudomonas sp., Sphingomonas sp. and Xanthomonas sp.
under aerobic conditions in the presence of additional carbon sources
has also been reported (Dykes et al., 1994; Sugiura et al., 1999).
Most of the known azo dye decolourizer was isolated from sludge,
soil and water samples. In all of the studies, the initial screening was
carried out either in shake flasks or solid azo dye media. Since the
majority of azo dye-degrading bacteria occur under anaerobic
conditions, isolation of azo dye-degrading bacteria is rather slow. It
can require up to four days to observe the decolourization zone due
to the high oxygen tension surrounding the bacterial colonies. The
usage of stab culture, as far as microbes are concerned, is usually
restricted to bacterial maintenance (Wise et al., 2006). To our
knowledge, the use of stab culture to screen for anaerobic and
aerobic xenobiotics-degrader has never been reported. Here, we
report on a simple method to screen for azodye-decolorizing bacteria,
using the well-known stab-culture method often used for culture
storage. Using this method, results can be obtained within 24 hours.
Materials and Methods
Isolation of bacteria: Soil samples, each measuring approximately
10 grams were taken randomly to a depth of 5 cm from the topsoil
using sterile spatula and stored in sterile screw-capped polycarbonate
Journal of Environmental Biology �January, 2009�
Syed et al.
Table - 1: Growth of S. Marcescens on various carbon sources in
BIOLOG’S GN microplates
Carbon sourceGrowth
(After 24 hr incubation)
Water (control) -
α-Cyclodextrin -
Dextrin +
Glycogen +/-
Tween 40 +
Tween 80 +
N-Acetyl-D-Galactosamine +
N-Acetyl-D-Glucosamine +
Adonitol +
L-Arabinose -
D-Arabitol -
D-Cellobiose -
i-Erythritol +
D-Fructose +
L-Fructose +
D-Galactose +
Gentiobiose -
α-D-Glucose +
m-Inositol +
α-D-Lactose -
Lactulose -
Maltose +
D-Mannitol +
D-Mannose +
D-Melibiose -
β-Methyl-D-Glucoside +
D-Psicose +
D-Raffinose -
L-Rhamnose -
D-Sorbitol +/-
Sucrose +
D-Trehalose +
Turanose -
Xylitol +
Pyruvic Acid Methyl Ester +
Succinic Acid-Mono-Methyl-Ester +
Acetic Acid -
Cis-Aconitic Acid +
Citric Acid +
Formic Acid +
D-Galactonic Acid Lactone -
D-Galacturonic Acid +/-
D-Gluconic Acid +
D-Glucosaminic Acid -
D-Glucuronic Acid -
α-Hydroxy butyric Acid -
β-Hydroxy butyric Acid -
γ- Hydroxy butyric Acid +/-
p-Hydroxy Penylacetic Acid +
Itaconic Acid -
α-Keto Butyric Acid -
α-Keto Glutaric Acid +
α-Keto Valeric Acid -
D,L-LacticAcid +
Malonic Acid -
Propionic Acid -
Quinic Acid -
D-Saccharic Acid -
Sebacic Acid -
Succinic Acid +
Bromosuccinic Acid +
Succinamic Acid -
Glucuronamide +/-
L-Alaninamide -
D-Alanine +
L-Alanine +
L-Alanyl-Glycine +
L-AsparticAcid +
L-Glutamic Acid +
Glycyl-L-Aspartic Acid -
Glycyl-L-Glutamic Acid +/-
L-Histidine +
Hydroxyl-L-Proline +/-
L-Leucine -
L-Ornithine -
L-Phenylalanine -
L-Proline +
L-Pyroglutamic Acid -
D-Serine -
L-Serine +
L-Threonine +/-
D,L-Carnitine -
γ-Amino Butyric Acid -
Urocanic Acid -
Inosine +
Uridine +
Thymidine +
Phenylethyl-amine -
Putrecine +/-
2-Aminoethanol -
2,3-Butanediol -
Glycerol +
D,L-α-Glycerol Phosphate +
α-D-Glucose-1-Phosphate +
+ = positive reaction
- = negative reaction
-/+ = borderline
90
Journal of Environmental Biology �January, 2009�
A simple method to screen for azo-dye-degrading bacteria
tubes. The soil samples were taken from several locations in Malaysia
near textile or dye-based industries. The exact locations were; (1)
Taman Tanjung and Lengkok Burmah, Penang, (2) Taman Miharja,
Selangor and (3) Bintulu, Sarawak. The samples were immediately
placed on ice until returned to University Putra Malaysia for further
examinations. Approximately one gram of each soil sample was
serially diluted ten times in 0.85% saline. Bacterial samples from
diluted soils were inoculated via stab inoculation into loosely-capped
microbiological test tubes containing the screening medium, semi-
solidified with 5.0 g l-1 of agar. The medium used was a modified
version of Hayase et al. (2000) and contained 2.34 g K2HPO
4, 1.33
g KH2PO
4, 0.20 g of MgSO
4.7H
2O, 1.00 g of (NH
4)2SO
4, 0.50 g of
NaCl, 0.10 g of yeast extract, 1.00 g of glucose and 1.0 ml of trace
element solution per liter, adjusted to the final pH of 7.0 with 3 M
NaOH and HCl. The trace element solutions contained 11.90 mg l-1
of CoCl2.6H
2O, 11.80 mg l-1 of NiCl
2, 6.30 mg l-1 of CrCl
2, 15.70 mg l-1 of
CuSO4.5H
2O, 0.97 g l-1 of FeCl
3, 0.78 g l-1 of CaCl
2.2H
2O and 10.00
mg l-1 of MnCl2.4H
2O. Bacteria from soil samples were tested for their
abilities to decolourise four azo dyes; Orange G (OG), Ponceau 2R
(P2R), Biebrich Scarlet (BS) and Direct blue 71 (DB). The final dye
concentration was 0.10 g l-1. Colour changes were qualitatively
observed.
Identification of bacteria: Isolates exhibiting distinct colonial
morphologies were isolated by repeated sub culturing into basal salt
medium and solidified basal salt medium until purified strains were
obtained. Identification at species level was performed by using
Biolog GN microplate (Biolog, Hayward, CA, USA) according to the
manufacturer’s instructions. Briefly,a pure culture of the bacterium
was grown on a Biolog Universal Growth agar plate. The bacteria
were swabbed from the surface of the agar plate, and suspended to
a specified density in GN/GP inoculating fluid. A hundred fifty µl of a
bacterial suspension was pipetted into each well of the micro-plate.
The micro-plate was incubated at 30oC or 35oC depending upon the
nature of the organism for 4-24 hr according to manufacturer’s
specification. The micro-plate was read with the Biolog MicroStationTM
system and compared to database.
Results and Discussion
Figure 1 shows typical screening results using the stab culture
method after 24 hr of incubation at room temperature. Of the twenty
soil samples tested, one soil sample showed the ability to decolorize
two azo dyes- Biebrich scarlet (BS) and Direct blue 71 (DB). The
location of the decolourization zones in both dye tubes indicates the
requirement for anoxic conditions. We were unable to isolate aerobic
azo dyes-decolorizing bacteria. If we had done so, the decolourization
zone would occur near or on the surface of the agar. The selective
advantage of a low oxygen tension environment on the
decolourisation of azo dyes, as observed, was probably due to the
electron-withdrawing properties of the azo bond itself that produces
electron deficiency at the site of cleavage and could only thus be
cleaved in the presence of reducing agents, generated during
anaerobic metabolism on static incubation (Rieger et al., 2002). After
further screening, a gram and oxidase negative bacterial isolate
was found to be the principal azo dyes degrader. The isolate was
identified by using the 95 carbon Biolog GN Microplate biochemical
tests (Table 1) as Serratia marcescens with 99% probability (0.749
similarity; 3.80 distance).
Methyl red degradation occurs both under aerobic and
anaerobic conditions. Aerobic reduction has been reported to
occur in several bacteria such as Pseudomonas sp. (Kulla et al.,
1983), Bacillus sp. (Maier et al. (2004) and Klebsiella pneumoniae
(Wong and Yuen, 1996). On the other hand, anaerobic conditions
are required in S. cerevisiae (Jadhav et al., 2007). Acetobacter
liquefaciens is unique in that decolourisation of methyl red occurs
efficiently under both aerobic and anaerobic (static) conditions
(So et al., 1990). The aerobic azoreductases from the carboxy-
orange-degrading Pseudomonas strains K22 and KF46
reductively cleave several sulfonated azo dyes (Kulla et al.,
1983). Therefore, the presence of an aerobic azoreductase in
Isolate 13 was probably responsible for the aerobic degradation
of methyl red dye. Under aerobic conditions, Biebrich Scarlet is
not readily metabolized by the isolate. However, under anaerobic
conditions, many bacteria reduce the highly electrophilic azo
bond in the dye molecule, reportedly by the activity of low
specificity cytoplasmic azoreductases, to produce colourless
aromatic amines (Pearce et al., 2003). The clear zone observed
at the bottom of the test tube suggests that the degradation of
Biebrich Scarlet occurs under anaerobic conditions. The
decolourization of azo dyes can easily be seen from the side of
the tube. The overall design is simple, yet effective, in screening
for aerobic and anaerobic azo dye-degrading bacteria.
Acknowledgments
This project was supported by funds from The Ministry of
Science, Technology and Innovation, Malaysia (MOSTI) under
IRPA-EA grant no: 09-02-04-0854-EA001.
Fig. 1: Stab inoculation of soil isolate in agar deep tubes containing BS (left)
and DB (right). The control uninoculated dye tubes are at the left of each pair
of tubes
BIEBRICH SCARLET (BS) DIRECT BLUE (DB)
Control Innoculated Control Innoculated
91
Journal of Environmental Biology �January, 2009�
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