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Azo dye biodegradation by microbial cultures immobilized in alginate beads
Environment International 31 (2005) 201 – 205 www.elsevier.com/locate/envint
Azo dye biodegradation by microbial cultures immobilized in alginate beads Silvia Steffana, Laura Bardib, Mario Marzonaa,* a
Dipartimento di Chimica Generale ed Organica Applicata-Universita` degli Studi di Torino C.so M. D’Azeglio, 48-10125 Torino, Italy b Istituto Sperimentale per la Nutrizione delle Piante, Via Pianezza, 115-10151 Torino, Italy Available online 21 November 2004
Abstract Microbial degradation of azo dyes usually starts in anaerobic conditions with a reductive cleavage of the azo bond, followed by an aerobic step necessary for the degradation of the aromatic amines formed. Because some reductive processes take place also in presence of molecular oxygen, a one-step azo dye degrading process has been investigated. A microbial consortium able to degrade ethyl orange in aerobic conditions has been selected and immobilized in alginate beads coated with polyacrylamide resin. Different concentrations of ethyl orange have been completely degraded in the presence of 1% glucose or starch as cosubstrates, and different beads preparation procedures have been studied to determine the best condition for microbial degradation. The catalytic activity of the immobilized consortium improved during five serial processes carried out for 30 days at room temperature. Three pure cultures were then isolated from the consortium. The one with the greatest degrading activity, a filamentous fungus, had a degradative capacity similar to that of the whole consortium. D 2004 Elsevier Ltd. All rights reserved. Keywords: Azo dye; Aerobic condition; Degradation kinetic
1. Introduction Azo dyes are the most important and diffuse group of synthetic dyes. It had been estimated that about 10% of the dye stuff used during the dyeing processes does not bind to the fibers and is released into the effluents (McMullan et al., 2001). Because all the industrially produced azo dyes are xenobiotic compounds, it is not surprising that they are recalcitrant in conventional sewage-treatment plants (Stolz, 2001). The decolorization process is achieved by reductive cleavage of the azo bond in anaerobic conditions, but the end-products of this reaction (aromatic amines) are often more dangerous than parent compounds (Idaka et al., 1987; Wong and Yuen, 1996). Moreover, their complete mineralization only occurs in the presence of molecular oxygen, so that a further step is required (Brown and Laboureur, 1983;
* Corresponding author. Tel.: +39 11 670 7597; fax: +39 011 670 7591. E-mail address: [email protected] (M. Marzona). 0160-4120/$ - see front matter D 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.envint.2004.09.016
Ekici et al., 2001; McMullan et al., 2001; Zeyer, 1985). Azo dyes decolorization can however be achieved by lignindegrading fungi and aerobic bacteria in aerobic conditions (Banat et al., 1996; Cripps et al., 1990; Hu, 2001; Paszczynski et al., 1992; Spadaro et al., 1992; Stolz, 2001; Wesemberg et al., 2001). In this case, one-step degradation process can be carried out, resulting in azo dye decolorization and at the same time aerobic degradation of the metabolites formed. In the present research, to protect microbial cells from possible toxic effects due to pollutant metabolites or changes in the environment conditions, immobilization of the biocatalyst was applied. Entrapment in alginate beads was chosen as the mildest immobilization method to preserve viability of cells and to reach all the advantages of heterogeneous catalysis (Sriamornsak, 1998; Tal et al., 2001). Using an immobilized biocatalyst in fact, the process can be carried out continuously allowing better process control, reduced operational cost, and washout of cells, continuous removal of toxic metabolites (Brodelius and Vandamme, 1987; Hartmeier, 1988). To this purpose,
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different immobilization processes were tested dealing with a polyacrylamide coating (Eudragit RL 100), to strengthen the beads (Ruggeri et al., 1991), and with the storage in CaCl2. Moreover, the effect of the presence and kind of an additional carbon source was investigated together with the persistence of the degradation activity in five serial degradation processes. Three strains were isolated from the selected consortium, and their azo dye degrading activity was compared to identify the most effective microorganism. Ethyl orange [4-[4-(diethylamino)-phenylazo]benzenesulfonic acid, sodium salt, (C2H5)2NC6H4N=NC6H4SO3Na, CAS 62758-12-7] was used as azo dye model.
2. Materials and methods 2.1. Microorganisms selection and culture conditions Azo dye-resistant microorganisms were obtained from a petroleum-polluted soil and from an aerated tank for the biological treatment of the effluents from a dye works. One gram soil sample and 1 ml water sample, respectively, were used as inoculum in 100 ml mineral medium [0.8 g/l K2HPO4, 0.2 g/l KH2PO4, 0.05 g/l CaSO4.2H2O, 0.5 g/l MgSO4.7H2O, 0.09 g/l FeSO4.7H20, 1g/l (NH4)2SO4] with 56.28 AM ethyl orange and 1% glucose as carbon and energy source. After an incubation at 28 8C for 5 days in a Dubnoff shaker at 130 rpm, 100 Al of each liquid culture was spreaded onto the same agar-solidified medium to isolate azo dye resistant strains. Each of these strains separately and a pool of them were used to prepare liquid cultures, in the same mineral medium, that were used as inoculum for the biodegradation tests. 2.2. Immobilization technique One gram sodium alginate was dissolved in 50 ml sterile liquid mineral medium and thoroughly stirred with 1 ml of the mixed culture of the selected consortium. The final mixture was extruded through a needle into 200 ml of 2% CaCl2 solution to yield 3- to 4-mm diameter alginate beads. After 20 min, they were filtered out and stored in 0.5% CaCl2 solution in the refrigerator for a time ranging from 24 to 300 h. Before use, the alginate matrix was coated with polyacrylamide resin by immersion in 100 ml of 96% ethyl alcohol with 15% w/w Eudragit RL 100 (Rofarma Italia) for 5 or 30 s. The beads were then dried by air for 2.5 h and washed with sterile water. 2.3. Biodegradation kinetic studies. Influence of the carbon source Four sets of five flasks were prepared to study the influence of the carbon source in the biodegradation kinetic of ethyl orange. In the first set, five 500-ml flasks were filled with 200 ml mineral medium, 56.28 AM ethyl orange, and 1% glucose (2
g) and sterilized at 120 8C for 20 min. Three of them were inoculated with 13 g of bead-entrapped microorganisms, one was inoculated with 2 ml of nonimmobilized microorganisms liquid culture, and the last one was not inoculated and used as control to evaluate possible photodegradation of ethyl orange. In the second set, the flasks were prepared as in the first set, but without glucose. In the third set, the flasks were prepared as in the first set, but with 1% starch instead of glucose. The fourth set of flasks was prepared as previously described, but glucose (1%) was added during the immobilization process, in concentration of 1% of the volume of that solution (0.5 g). All the flasks were incubated in a Dubnoff shaker at 28 8C and 130 rpm. Periodically, 10 ml of solution of each flask was filtered and analyzed at 513 nm (ethyl orange isosbestic point) by UV–Vis spectrometer (ATI unicam). The solution was also analyzed for the detection of sulfanilic acid, one of the degradation metabolites formed by azo group symmetric cleavage of ethyl orange. The analysis was carried out by thin layer chromatography (TLC); the eluent used was the organic phase of BAW (40% buthanol, 10% acetic acid, 50% demineralized water). 2.4. Biodegradation kinetic studies. Persistence of biocatalyst’s activity One more set of flasks was prepared in the same way as the first set was prepared for testing the influence of carbon sources, but at the end of the biodegradation kinetic, the beads were removed by filtration and poured in a new set of flasks for a new biodegradation kinetic. This process was repeated five times, and the times required for full degradation (when beads were recovered to start a new degradation kinetic) were different: 211 h in the first degradation kinetic, 165 h in the second degradation kinetic, 119 h in the third degradation kinetic, 119 h in the fourth degradation kinetic. 2.5. Azo dye degrading activity of the strains isolated from the consortium One set of four flasks was prepared to evaluate the azo dye degrading activity of the three strains isolated from the selected consortium. Each flask was filled with 200 ml mineral medium, 56.28 AM ethyl orange, and 1% glucose, sterilized at 120 8C for 20 min and inoculated with one of the three strains (1 ml of a suspension prepared as described in Section 2.1). All flasks were incubated at 28 8C in oscillatory shaker for 135 h, and the solution was analyzed at this time for the ethyl orange residual concentration, as previously described. An additional ethyl orange biodegradation test was carried out to compare the activity of the consortium and of the most effective strain. This set of flasks was analyzed as described above.
S. Steffan et al. / Environment International 31 (2005) 201–205
3. Results and discussion 3.1. Immobilization technique The selected consortium showed ethyl orange decolorization activity in presence of molecular oxygen, promoting degradation of metabolites, such as aromatic amines, formed during this process. TLC analysis for sulfanilic acid showed, as expected, that no aromatic amines were accumulated in the decolorization process. In aerobic condition, in fact, they are quite easily mineralized to simple compounds, such as CO2, H2O, and NH3, so that the polluted solution is biodegraded, not only decolorized (Ekici et al., 2001; Zeyer, 1985). The obtained ethyl orange degradation kinetics fitted with a first-order curve, as illustrated in Fig. 1. Whereas a half-degradation time (t 1/2) of 57.7 h was observed for the nonimmobilized biocatalyst system, a faster degradation kinetic was showed by the biocatalyst entrapped in alginate beads coated with polyacrylamide resin by immersion for 5 s (t 1/2=35.5 h) and 30 s (t 1/2=33.6 h) in the Eudragit solution. Entrapment is the most widely used technique for immobilization of whole cells, and alginate is a suitable matrix material because it is nontoxic and the method used for its gelation is mild towards the microorganisms (Sriamornsak, 1998; Cohen, 2001; Kourkoutas et al.,
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2004). Nevertheless, it leads to relatively soft products, and because it is bed on ionotropic gel formation, products are not stable in presence of ions with charges opposite to those required for the formation of the gel (Ruggeri et al., 1991). Phosphate ions, used in this work as buffer substances, can also be responsible for a gradual dissolution of the alginate spheres, because phosphate reacts with calcium, which is withdrawn from the beads (Hartmeier,1988). Immersion in Eudragit solution resulted in an increment of mechanical properties of alginate beads, which were coated with an external polyacrylamide resin layer giving strength to their structure and avoiding biomass leakage (Ruggeri et al., 1991). These advantages were greater for immersion time of 30 s than 5 s, but practically no differences were observed between the corresponding biodegradation curves (Fig. 1). In comparison with the nonimmobilized system, the both of them showed shorter half degradation times (t 1/2), meaning that better conditions were achieved inside the bead, where partition effects lead to a modified microenvironment (Brodelius and Vandamme, 1987). Besides, the coating always allows azo dye, glucose, and other nutrients to move inside the beads and be metabolized. A longer time in the Eudragit solution allowed a better external layer and a stronger bead structure, without hindering compound diffusion through the coating towards biocatalysts. Different storage times in CaCl2 seem to slightly influence the ethyl orange degradation kinetics, as supported by t 1/2 values of biodegradation kinetics by beads stored in CaCl2 for 24 h (t 1/2=33.6 h), 65 h (t 1/2=30.9 h), and 300 h (t 1/2=26.9 h). These results also show that azo dye degradation activity is maintained by the selected consortium entrapped in alginate beads at least up to 300 h storage in CaCl2 solution. 3.2. Biodegradation kinetic studies. Influence of the carbon source
Fig. 1. (a) Ethyl orange biodegradation kinetics by nonimmobilized biocatalyst (4, broken line), biocatalyst immobilized in alginate beads coated by immersion in Eudragit solution for 5 s ( w, dotted line) and for 30 s (5, continuous line). (b) Half degradation time of the previously described kinetics.
Azo dye biodegradation usually starts with the reduction of the azo bond by specific enzymes. For their activation, an additional energy source is required (Nigam et al., 1996), otherwise only sorption of ethyl orange into the beads can occur, as illustrated in Fig. 2. In the flask carried out with nonimmobilized consortium and without glucose, ethyl orange persisted for more than 100 h; in the flask with the immobilized biocatalyst and without glucose, the residual percentage of the dye rapidly decreased to 68.3% in the first 18 h and then remained constant until the end of the test, due to physical interaction between the bead matrix and the dye. Ethyl orange sorption into the beads is necessary to allow the contact between the dye and the microorganisms, but if an additional carbon source is not available, the suitable enzymes are not induced and the ethyl orange is not degraded (Nigam et al., 1996). When glucose is present, instead, sorbed dye is metabolized by immobilized consortium, so that new molecules of the ethyl orange can be
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S. Steffan et al. / Environment International 31 (2005) 201–205 Table 1 Ethyl orange (EO) degradation by the selected consortium, FUS, BIA, and NER after 135 h kinetic
Fig. 2. Ethyl orange biodegradation kinetics by immobilized biocatalyst in presence of (a) 0% glucose, (b) 1% glucose (in alginate beads), (c) 1% glucose (in liquid medium), (d) 1% starch, and (e) by non-immobilized biocatalyst with any additional energy source.
sorbed and degraded, which is why its total residue percentage is decreased proportionally to the glucose amount. Because glucose easily got inside the bead, best results were obtained when glucose was directly added to the liquid medium (t 1/2=30.9 h) and not during the immobilization process (t 1/2=58.7 h). In Fig. 2, azo dye biodegradation kinetic with 1% starch is also reported; its t 1/2 (27.9 h) points out that the used consortium may cleave the starch molecule using the formed products as source of carbon. Starch, that is cheaper than glucose, in the tested conditions is therefore a good alternative to glucose for aerobic degradation of azo dyes (Brown and Laboureur, 1983). 3.3. Biodegradation kinetic studies. Persistence of biocatalyst’s activity Subsequent use of alginate beads showed that increment in ethyl orange biodegradation activity was achieved with time. This activity was maintained for more than 2 months, when biodegradation rate is increased in five serial azo dye
Microorganisms
Degraded EO (%)
Consortium FUS BIA NER
80.1 89.5 38.7 70
degradation processes carried out at room temperature. This result was also confirmed by t 1/2 values, which were decreased from 47.6 to 14.5 h, as shown in Fig. 3. No release of microorganisms from beads into the liquid growth medium was observed; as inside the beads, cell multiplication is hindered and biomass can only be indirectly quantified (Kourkoutas et al., 2004), we supposed that faster degradation was probably caused by metabolic adaptation of the microorganisms (Cohen, 2001; Kourkoutas et al., 2004). Several activities can lead to the adaptation of a microbial community toward toxic or recalcitrant compounds, and this is very useful to improve biodegradation processes (Baldi et al., 1997). Cell immobilization is the best way to recover, when possible, at the end of a treatment the activated biocatalyst to be reused in a new process (Kourkoutas et al., 2004). 3.4. Azo dye degrading activity of the strains isolated from the consortium Pure strains were isolated from the microbial consortium to find out the strains with the higher degradative activity and to start a metabolic characterisation of them. The final aim is to assure that they do not produce toxic metabolites and are safe if used in bioremediation processes; moreover, entrapment of desired microbial species allows to maintain its activity without competition with other microorganisms present in the waste fluid (Cohen, 2001). Following a morphologic observation of colonies grown in agar-solified selective medium, three strains were isolated as the probable more representative and were called FUS, BIA, and NER. The results of the comparison of their azo dye degrading activity are reported in Table 1. The residual percentage of ethyl orange after 135 h shows that FUS, a filamentous fungus, seems to be the one with the greatest degrading activity (89.5% ethyl orange degradation instead of 38.7% observed for BIA and 70.0% for NER). Table 2 Residual ethyl orange (EO) concentration in the kinetics carried out with FUS strain and with the selected consortium
Fig. 3. Half-degradation time in degradation kinetics serially carried out with the same bead-entrapped microbial inoculum. Total time required for full degradation in each kinetic was I—211 h, II—165 h, III—119 h, IV— 119 h, and V—45 h.
Time (h)
Residual EO concentration with FUS strain (%)
Residual EO concentration with consortium (%)
0 24 47 95 125
100 93.3 42.4 14.4 12.2
100 92.4 49.4 11.9 14.4
S. Steffan et al. / Environment International 31 (2005) 201–205
The comparison of the ethyl orange degradation kinetic by the selected consortium and by FUS strain (Table 2) points out that all values are quite similar, confirming the prevailing degradative rule of this strain inside the consortium. Some filamentous fungi are able to synthesize lignin peroxidase (LiP; Kumari et al., 2002), and this enzyme is known for its ability to degrade several xenobiotics, such as PAHs or azo dyes. The oxidized form of LiP abstracted two electrons from the substrates, starting a radical process which is specific towards xenobiotics (Stolz, 2001). According to these observation, ethyl orange degradation by the selected filamentous fungus probably occurred with the abovementioned pathway, through an asymmetric cleavage of the azo bond without producing aromatic amines, as confirmed by the detection of no sulfanilic acid concentration in the solutions of the kinetics.
4. Conclusion Azo dyes’ biodegradation often leads to aromatic amines more toxic than parent compounds. In this work, a microbial consortium was isolated, which is able to decolorize ethyl orange and mineralize metabolites in aerobic conditions. Using the selected consortium, ethyl orange was completely degraded in the presence of molecular oxygen, and faster degradation rate was observed for the consortium entrapped in alginate beads coated with Eudragit (immersion time 30 s) and stored in CaCl2 (t 1/2=26.9 h). Additional energy source had to be added to activate cometabolism responsible for ethyl orange degradation; to this purpose, both 1% glucose or starch can be used. Fastest biodegradation kinetic was observed after five serial processes carried out with the same set of beads (t 1/2=14.5 h), which allowed best adaptation of microorganisms to the environment and the activation of the degrading metabolic pathway. The comparison of the azo dye degrading activity has shown that the selected filamentous fungus FUS strain is the most active among the three strains isolated from the consortium in metabolizing ethyl orange. Its activity is probably due to the ability of this organism to synthesize lignin peroxidase, which causes the asymmetric cleavage of the azo bond without forming aromatic amines, often toxic and cancerogenic.
References Baldi F, Janssen DB, Reineke W. Adaptation: dynamics of genes, enzyme activities and populations. In: Wild JR, editor. Perspectives in bioremediation, p. 93 – 105.
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Banat IM, Nigam P, Singh D, Marchant R. Microbial decolorization of textile-dye-containing effluents: a review. Bioresour Technol 1996;58:217 – 27. Brodelius P, Vandamme EJ. Immobilized cell systems. Biotechnology Enzyme Technology, vol. 7a. Weinheim, Federal Republic of Germany7 VCH Verlagsgesellschaft; 1987. p. 405 – 64. Brown D, Laboureur P. The aerobic biodegradability of primary aromatic amines. Chemosphere 1983;12:405 – 14. Cohen Y. Biofiltration—the treatment of fluids by microorganisms immobilized into the filter bedding material: a review. Bioresour Technol 2001;77:257 – 74. Cripps C, Bumpus JA, Aust SD. Biodegradation of azo and heterocyclic dyes by Phanerochaete chrysosporium. Appl Environ Microbiol 1990;56:1114 – 8. Ekici P, Leupold G, Parlar H. Degradability of selected azo dye metabolites in activated sludge system. Chemosphere 2001;44:721 – 8. Hartmeier W. Immobilized biocatalysts An introduction. Berlin, Germany7 Spriger-Verlag; 1988. Hu TL. Kinetics of azoreductase and assessment of toxicity of metabolic products from azo dyes by Pseudomonas luteola. Water Sci Technol 2001;43:261 – 9. Idaka E, Ogawa T, Horitsu H. Reductive metabolism of aminoazobenzenes by Pseudomonas cepacia. Bull Environ Contam Toxicol 1987; 39:100 – 7. Kourkoutas Y, Bekatorou A, Banat IM, Marchant R, Koutinas AA. Immobilization technologies and support materials suitable in alcohol beverages production: a review. Food Microbiol 2004;21:377 – 97. Kumari M, Yadav RS, Yadav KD. Secretion of ligninperoxidase by Penicillum citrinum, Fusarium oxysporum and Aspergillus terreus. Indian J Exp Biol 2002;40:802 – 6. McMullan G, Meehan C, Conneely A, Kirby N, Robinson T, Nigam P, et al. Microbial decolourisation and degradation of textile dyes. Appl Microbiol Biotechnol 2001;56:81 – 7. Nigam P, McMullan G, Banat IM, Marchant R. Decolourisation of effluent from the textile industry by a microbial consortium. Biotechnol Lett 1996;18:117 – 20. Paszczynski A, Pasti.Grigsby MB, Goszczynski S, Crawford RL, Crawford DL. Mineralization of sulfonated azo dyes and sulfanilic acid by Phanerochaete chrysosporium and Streptomyces chromofuscus. Appl Environ Microbiol 1992;58:3598 – 604. Ruggeri B, Sassi G, Specchia V, Bosco F, Marzona M. Alginate beads coated with polyacrylamide resin: potential as a biocatalyst. Process Biochem 1991;26:331 – 5. Spadaro JT, Gold MH, Renganathan V. Degradation of azo dyes by the lignin-degrading fungus Phanerochaete chrysosporium. Appl Environ Microbiol 1992;58:2397 – 401. Sriamornsak P. Preliminary investigation of some polysaccharides as a carrier for cell entrapment. Eur J Pharm Biopharm 1998;46:233 – 6. Stolz A. Basic and applied aspects in the microbial degradation of azo dyes. Appl Microbiol Biotechnol 2001;56:69 – 80. Tal Y, Schwartsburd B, Nussinovitch A, van Rijn J. Enumeration and factors influencing the relative abundance of a denitrifier, Pseudomonas sp JR12, entrapped in alginate beads. Environ Pollut 2001;112:99 – 106. Wesemberg D, Vanhulle S, Buchon F, Lucas M, Caillou S, Mertens V, et al. Decolorization and detoxification of dye-industry wastewater using novel white-rot fungi. Proceedings of the First European Bioremediation Conference, Chania, Crete, Greece, p. 209 – 12. Wong PK, Yuen PY. Decolorization and biodegradation of methyl red by Klebsiella pneumoniae RS-13. Water Res 1996;30:1736 – 44. Zeyer J, Wasserfallen A, Timmis KN. Microbial mineralization of ringsubstituted anilines through an ortho-cleavege pathway. Appl Environ Microbiol 1985;50:447 – 53.
Azo dye biodegradation by microbial cultures immobilized in alginate beads Silvia Steffana, Laura Bardib, Mario Marzonaa,* a
Dipartimento di Chimica Generale ed Organica Applicata-Universita` degli Studi di Torino C.so M. D’Azeglio, 48-10125 Torino, Italy b Istituto Sperimentale per la Nutrizione delle Piante, Via Pianezza, 115-10151 Torino, Italy Available online 21 November 2004
Abstract Microbial degradation of azo dyes usually starts in anaerobic conditions with a reductive cleavage of the azo bond, followed by an aerobic step necessary for the degradation of the aromatic amines formed. Because some reductive processes take place also in presence of molecular oxygen, a one-step azo dye degrading process has been investigated. A microbial consortium able to degrade ethyl orange in aerobic conditions has been selected and immobilized in alginate beads coated with polyacrylamide resin. Different concentrations of ethyl orange have been completely degraded in the presence of 1% glucose or starch as cosubstrates, and different beads preparation procedures have been studied to determine the best condition for microbial degradation. The catalytic activity of the immobilized consortium improved during five serial processes carried out for 30 days at room temperature. Three pure cultures were then isolated from the consortium. The one with the greatest degrading activity, a filamentous fungus, had a degradative capacity similar to that of the whole consortium. D 2004 Elsevier Ltd. All rights reserved. Keywords: Azo dye; Aerobic condition; Degradation kinetic
1. Introduction Azo dyes are the most important and diffuse group of synthetic dyes. It had been estimated that about 10% of the dye stuff used during the dyeing processes does not bind to the fibers and is released into the effluents (McMullan et al., 2001). Because all the industrially produced azo dyes are xenobiotic compounds, it is not surprising that they are recalcitrant in conventional sewage-treatment plants (Stolz, 2001). The decolorization process is achieved by reductive cleavage of the azo bond in anaerobic conditions, but the end-products of this reaction (aromatic amines) are often more dangerous than parent compounds (Idaka et al., 1987; Wong and Yuen, 1996). Moreover, their complete mineralization only occurs in the presence of molecular oxygen, so that a further step is required (Brown and Laboureur, 1983;
* Corresponding author. Tel.: +39 11 670 7597; fax: +39 011 670 7591. E-mail address: [email protected] (M. Marzona). 0160-4120/$ - see front matter D 2004 Elsevier Ltd. All rights reserved. doi:10.1016/j.envint.2004.09.016
Ekici et al., 2001; McMullan et al., 2001; Zeyer, 1985). Azo dyes decolorization can however be achieved by lignindegrading fungi and aerobic bacteria in aerobic conditions (Banat et al., 1996; Cripps et al., 1990; Hu, 2001; Paszczynski et al., 1992; Spadaro et al., 1992; Stolz, 2001; Wesemberg et al., 2001). In this case, one-step degradation process can be carried out, resulting in azo dye decolorization and at the same time aerobic degradation of the metabolites formed. In the present research, to protect microbial cells from possible toxic effects due to pollutant metabolites or changes in the environment conditions, immobilization of the biocatalyst was applied. Entrapment in alginate beads was chosen as the mildest immobilization method to preserve viability of cells and to reach all the advantages of heterogeneous catalysis (Sriamornsak, 1998; Tal et al., 2001). Using an immobilized biocatalyst in fact, the process can be carried out continuously allowing better process control, reduced operational cost, and washout of cells, continuous removal of toxic metabolites (Brodelius and Vandamme, 1987; Hartmeier, 1988). To this purpose,
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different immobilization processes were tested dealing with a polyacrylamide coating (Eudragit RL 100), to strengthen the beads (Ruggeri et al., 1991), and with the storage in CaCl2. Moreover, the effect of the presence and kind of an additional carbon source was investigated together with the persistence of the degradation activity in five serial degradation processes. Three strains were isolated from the selected consortium, and their azo dye degrading activity was compared to identify the most effective microorganism. Ethyl orange [4-[4-(diethylamino)-phenylazo]benzenesulfonic acid, sodium salt, (C2H5)2NC6H4N=NC6H4SO3Na, CAS 62758-12-7] was used as azo dye model.
2. Materials and methods 2.1. Microorganisms selection and culture conditions Azo dye-resistant microorganisms were obtained from a petroleum-polluted soil and from an aerated tank for the biological treatment of the effluents from a dye works. One gram soil sample and 1 ml water sample, respectively, were used as inoculum in 100 ml mineral medium [0.8 g/l K2HPO4, 0.2 g/l KH2PO4, 0.05 g/l CaSO4.2H2O, 0.5 g/l MgSO4.7H2O, 0.09 g/l FeSO4.7H20, 1g/l (NH4)2SO4] with 56.28 AM ethyl orange and 1% glucose as carbon and energy source. After an incubation at 28 8C for 5 days in a Dubnoff shaker at 130 rpm, 100 Al of each liquid culture was spreaded onto the same agar-solidified medium to isolate azo dye resistant strains. Each of these strains separately and a pool of them were used to prepare liquid cultures, in the same mineral medium, that were used as inoculum for the biodegradation tests. 2.2. Immobilization technique One gram sodium alginate was dissolved in 50 ml sterile liquid mineral medium and thoroughly stirred with 1 ml of the mixed culture of the selected consortium. The final mixture was extruded through a needle into 200 ml of 2% CaCl2 solution to yield 3- to 4-mm diameter alginate beads. After 20 min, they were filtered out and stored in 0.5% CaCl2 solution in the refrigerator for a time ranging from 24 to 300 h. Before use, the alginate matrix was coated with polyacrylamide resin by immersion in 100 ml of 96% ethyl alcohol with 15% w/w Eudragit RL 100 (Rofarma Italia) for 5 or 30 s. The beads were then dried by air for 2.5 h and washed with sterile water. 2.3. Biodegradation kinetic studies. Influence of the carbon source Four sets of five flasks were prepared to study the influence of the carbon source in the biodegradation kinetic of ethyl orange. In the first set, five 500-ml flasks were filled with 200 ml mineral medium, 56.28 AM ethyl orange, and 1% glucose (2
g) and sterilized at 120 8C for 20 min. Three of them were inoculated with 13 g of bead-entrapped microorganisms, one was inoculated with 2 ml of nonimmobilized microorganisms liquid culture, and the last one was not inoculated and used as control to evaluate possible photodegradation of ethyl orange. In the second set, the flasks were prepared as in the first set, but without glucose. In the third set, the flasks were prepared as in the first set, but with 1% starch instead of glucose. The fourth set of flasks was prepared as previously described, but glucose (1%) was added during the immobilization process, in concentration of 1% of the volume of that solution (0.5 g). All the flasks were incubated in a Dubnoff shaker at 28 8C and 130 rpm. Periodically, 10 ml of solution of each flask was filtered and analyzed at 513 nm (ethyl orange isosbestic point) by UV–Vis spectrometer (ATI unicam). The solution was also analyzed for the detection of sulfanilic acid, one of the degradation metabolites formed by azo group symmetric cleavage of ethyl orange. The analysis was carried out by thin layer chromatography (TLC); the eluent used was the organic phase of BAW (40% buthanol, 10% acetic acid, 50% demineralized water). 2.4. Biodegradation kinetic studies. Persistence of biocatalyst’s activity One more set of flasks was prepared in the same way as the first set was prepared for testing the influence of carbon sources, but at the end of the biodegradation kinetic, the beads were removed by filtration and poured in a new set of flasks for a new biodegradation kinetic. This process was repeated five times, and the times required for full degradation (when beads were recovered to start a new degradation kinetic) were different: 211 h in the first degradation kinetic, 165 h in the second degradation kinetic, 119 h in the third degradation kinetic, 119 h in the fourth degradation kinetic. 2.5. Azo dye degrading activity of the strains isolated from the consortium One set of four flasks was prepared to evaluate the azo dye degrading activity of the three strains isolated from the selected consortium. Each flask was filled with 200 ml mineral medium, 56.28 AM ethyl orange, and 1% glucose, sterilized at 120 8C for 20 min and inoculated with one of the three strains (1 ml of a suspension prepared as described in Section 2.1). All flasks were incubated at 28 8C in oscillatory shaker for 135 h, and the solution was analyzed at this time for the ethyl orange residual concentration, as previously described. An additional ethyl orange biodegradation test was carried out to compare the activity of the consortium and of the most effective strain. This set of flasks was analyzed as described above.
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3. Results and discussion 3.1. Immobilization technique The selected consortium showed ethyl orange decolorization activity in presence of molecular oxygen, promoting degradation of metabolites, such as aromatic amines, formed during this process. TLC analysis for sulfanilic acid showed, as expected, that no aromatic amines were accumulated in the decolorization process. In aerobic condition, in fact, they are quite easily mineralized to simple compounds, such as CO2, H2O, and NH3, so that the polluted solution is biodegraded, not only decolorized (Ekici et al., 2001; Zeyer, 1985). The obtained ethyl orange degradation kinetics fitted with a first-order curve, as illustrated in Fig. 1. Whereas a half-degradation time (t 1/2) of 57.7 h was observed for the nonimmobilized biocatalyst system, a faster degradation kinetic was showed by the biocatalyst entrapped in alginate beads coated with polyacrylamide resin by immersion for 5 s (t 1/2=35.5 h) and 30 s (t 1/2=33.6 h) in the Eudragit solution. Entrapment is the most widely used technique for immobilization of whole cells, and alginate is a suitable matrix material because it is nontoxic and the method used for its gelation is mild towards the microorganisms (Sriamornsak, 1998; Cohen, 2001; Kourkoutas et al.,
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2004). Nevertheless, it leads to relatively soft products, and because it is bed on ionotropic gel formation, products are not stable in presence of ions with charges opposite to those required for the formation of the gel (Ruggeri et al., 1991). Phosphate ions, used in this work as buffer substances, can also be responsible for a gradual dissolution of the alginate spheres, because phosphate reacts with calcium, which is withdrawn from the beads (Hartmeier,1988). Immersion in Eudragit solution resulted in an increment of mechanical properties of alginate beads, which were coated with an external polyacrylamide resin layer giving strength to their structure and avoiding biomass leakage (Ruggeri et al., 1991). These advantages were greater for immersion time of 30 s than 5 s, but practically no differences were observed between the corresponding biodegradation curves (Fig. 1). In comparison with the nonimmobilized system, the both of them showed shorter half degradation times (t 1/2), meaning that better conditions were achieved inside the bead, where partition effects lead to a modified microenvironment (Brodelius and Vandamme, 1987). Besides, the coating always allows azo dye, glucose, and other nutrients to move inside the beads and be metabolized. A longer time in the Eudragit solution allowed a better external layer and a stronger bead structure, without hindering compound diffusion through the coating towards biocatalysts. Different storage times in CaCl2 seem to slightly influence the ethyl orange degradation kinetics, as supported by t 1/2 values of biodegradation kinetics by beads stored in CaCl2 for 24 h (t 1/2=33.6 h), 65 h (t 1/2=30.9 h), and 300 h (t 1/2=26.9 h). These results also show that azo dye degradation activity is maintained by the selected consortium entrapped in alginate beads at least up to 300 h storage in CaCl2 solution. 3.2. Biodegradation kinetic studies. Influence of the carbon source
Fig. 1. (a) Ethyl orange biodegradation kinetics by nonimmobilized biocatalyst (4, broken line), biocatalyst immobilized in alginate beads coated by immersion in Eudragit solution for 5 s ( w, dotted line) and for 30 s (5, continuous line). (b) Half degradation time of the previously described kinetics.
Azo dye biodegradation usually starts with the reduction of the azo bond by specific enzymes. For their activation, an additional energy source is required (Nigam et al., 1996), otherwise only sorption of ethyl orange into the beads can occur, as illustrated in Fig. 2. In the flask carried out with nonimmobilized consortium and without glucose, ethyl orange persisted for more than 100 h; in the flask with the immobilized biocatalyst and without glucose, the residual percentage of the dye rapidly decreased to 68.3% in the first 18 h and then remained constant until the end of the test, due to physical interaction between the bead matrix and the dye. Ethyl orange sorption into the beads is necessary to allow the contact between the dye and the microorganisms, but if an additional carbon source is not available, the suitable enzymes are not induced and the ethyl orange is not degraded (Nigam et al., 1996). When glucose is present, instead, sorbed dye is metabolized by immobilized consortium, so that new molecules of the ethyl orange can be
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S. Steffan et al. / Environment International 31 (2005) 201–205 Table 1 Ethyl orange (EO) degradation by the selected consortium, FUS, BIA, and NER after 135 h kinetic
Fig. 2. Ethyl orange biodegradation kinetics by immobilized biocatalyst in presence of (a) 0% glucose, (b) 1% glucose (in alginate beads), (c) 1% glucose (in liquid medium), (d) 1% starch, and (e) by non-immobilized biocatalyst with any additional energy source.
sorbed and degraded, which is why its total residue percentage is decreased proportionally to the glucose amount. Because glucose easily got inside the bead, best results were obtained when glucose was directly added to the liquid medium (t 1/2=30.9 h) and not during the immobilization process (t 1/2=58.7 h). In Fig. 2, azo dye biodegradation kinetic with 1% starch is also reported; its t 1/2 (27.9 h) points out that the used consortium may cleave the starch molecule using the formed products as source of carbon. Starch, that is cheaper than glucose, in the tested conditions is therefore a good alternative to glucose for aerobic degradation of azo dyes (Brown and Laboureur, 1983). 3.3. Biodegradation kinetic studies. Persistence of biocatalyst’s activity Subsequent use of alginate beads showed that increment in ethyl orange biodegradation activity was achieved with time. This activity was maintained for more than 2 months, when biodegradation rate is increased in five serial azo dye
Microorganisms
Degraded EO (%)
Consortium FUS BIA NER
80.1 89.5 38.7 70
degradation processes carried out at room temperature. This result was also confirmed by t 1/2 values, which were decreased from 47.6 to 14.5 h, as shown in Fig. 3. No release of microorganisms from beads into the liquid growth medium was observed; as inside the beads, cell multiplication is hindered and biomass can only be indirectly quantified (Kourkoutas et al., 2004), we supposed that faster degradation was probably caused by metabolic adaptation of the microorganisms (Cohen, 2001; Kourkoutas et al., 2004). Several activities can lead to the adaptation of a microbial community toward toxic or recalcitrant compounds, and this is very useful to improve biodegradation processes (Baldi et al., 1997). Cell immobilization is the best way to recover, when possible, at the end of a treatment the activated biocatalyst to be reused in a new process (Kourkoutas et al., 2004). 3.4. Azo dye degrading activity of the strains isolated from the consortium Pure strains were isolated from the microbial consortium to find out the strains with the higher degradative activity and to start a metabolic characterisation of them. The final aim is to assure that they do not produce toxic metabolites and are safe if used in bioremediation processes; moreover, entrapment of desired microbial species allows to maintain its activity without competition with other microorganisms present in the waste fluid (Cohen, 2001). Following a morphologic observation of colonies grown in agar-solified selective medium, three strains were isolated as the probable more representative and were called FUS, BIA, and NER. The results of the comparison of their azo dye degrading activity are reported in Table 1. The residual percentage of ethyl orange after 135 h shows that FUS, a filamentous fungus, seems to be the one with the greatest degrading activity (89.5% ethyl orange degradation instead of 38.7% observed for BIA and 70.0% for NER). Table 2 Residual ethyl orange (EO) concentration in the kinetics carried out with FUS strain and with the selected consortium
Fig. 3. Half-degradation time in degradation kinetics serially carried out with the same bead-entrapped microbial inoculum. Total time required for full degradation in each kinetic was I—211 h, II—165 h, III—119 h, IV— 119 h, and V—45 h.
Time (h)
Residual EO concentration with FUS strain (%)
Residual EO concentration with consortium (%)
0 24 47 95 125
100 93.3 42.4 14.4 12.2
100 92.4 49.4 11.9 14.4
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The comparison of the ethyl orange degradation kinetic by the selected consortium and by FUS strain (Table 2) points out that all values are quite similar, confirming the prevailing degradative rule of this strain inside the consortium. Some filamentous fungi are able to synthesize lignin peroxidase (LiP; Kumari et al., 2002), and this enzyme is known for its ability to degrade several xenobiotics, such as PAHs or azo dyes. The oxidized form of LiP abstracted two electrons from the substrates, starting a radical process which is specific towards xenobiotics (Stolz, 2001). According to these observation, ethyl orange degradation by the selected filamentous fungus probably occurred with the abovementioned pathway, through an asymmetric cleavage of the azo bond without producing aromatic amines, as confirmed by the detection of no sulfanilic acid concentration in the solutions of the kinetics.
4. Conclusion Azo dyes’ biodegradation often leads to aromatic amines more toxic than parent compounds. In this work, a microbial consortium was isolated, which is able to decolorize ethyl orange and mineralize metabolites in aerobic conditions. Using the selected consortium, ethyl orange was completely degraded in the presence of molecular oxygen, and faster degradation rate was observed for the consortium entrapped in alginate beads coated with Eudragit (immersion time 30 s) and stored in CaCl2 (t 1/2=26.9 h). Additional energy source had to be added to activate cometabolism responsible for ethyl orange degradation; to this purpose, both 1% glucose or starch can be used. Fastest biodegradation kinetic was observed after five serial processes carried out with the same set of beads (t 1/2=14.5 h), which allowed best adaptation of microorganisms to the environment and the activation of the degrading metabolic pathway. The comparison of the azo dye degrading activity has shown that the selected filamentous fungus FUS strain is the most active among the three strains isolated from the consortium in metabolizing ethyl orange. Its activity is probably due to the ability of this organism to synthesize lignin peroxidase, which causes the asymmetric cleavage of the azo bond without forming aromatic amines, often toxic and cancerogenic.
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