Decolorization of 1-amino-4-bromoanthraquinone-2-sulfonic acid by a newly isolated strain of Sphingomonas herbicidovorans

Decolorization of 1-amino-4-bromoanthraquinone-2-sulfonic acid by a newly isolated strain of Sphingomonas herbicidovorans

International Biodeterioration & Biodegradation 63 (2009) 88–92 Contents lists available at ScienceDirect International Biodeterioration & Biodegrad...

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International Biodeterioration & Biodegradation 63 (2009) 88–92

Contents lists available at ScienceDirect

International Biodeterioration & Biodegradation journal homepage: www.elsevier.com/locate/ibiod

Decolorization of 1-amino-4-bromoanthraquinone-2-sulfonic acid by a newly isolated strain of Sphingomonas herbicidovoransq Li Fan a, b, Shunni Zhu a, b, Dongqi Liu a, b, Jinren Ni a, b, * a b

Shenzhen Graduate School, Peking University, The Key Laboratory for Environmental and Urban Sciences, Guang Dong 518055, China Department of Environmental Engineering, Peking University, The Key Laboratory of Water and Sediment Sciences, Ministry of Education, Beijing 100871, China

a r t i c l e i n f o

a b s t r a c t

Article history: Received 15 May 2007 Received in revised form 12 June 2008 Accepted 30 July 2008 Available online 17 September 2008

A novel isolate of Sphingomonas herbicidovorans could decolorize 1-amino-4-bromoanthraquinone-2sulfonic acid (bromoamine acid, BAA), an intermediate of anthraquinone dyes, and grow with it as the sole source of carbon. The strain was identified by 16S rRNA gene sequencing and physiologicalbiochemical test. The optimal condition for both decolorization and cell growth was found at temperature of 30  C and pH 7.0, respectively. Furthermore, the decolorization efficiency could be enhanced with higher shaking speed. The percentage of BAA decolorization could be over 98% within 24 h even for the initial concentration greater than 1000 mg l1. The decolorization kinetics could be reasonably described by the Monod equation. Additional carbon sources such as glucose could enhance the decolorization rate. During the decolorization process, the molecular of BAA cleaved, releasing phthalic acid and an end product which might be benzene derivative substituted by amino, bromo, hydroxyl and sulfonate groups according to the infrared spectral analysis. Ó 2008 Elsevier Ltd. All rights reserved.

Keywords: Decolorization Anthraquinone Sphingomonas herbicidovorans Synthetic dyes

1. Introduction Synthetic dyes are extensively used in textile dyeing, paper, color photography, cosmetics and other industries. The amount of dyes produced in the world is estimated to be over 7  105 tons per year, while the quantity of dyes discharged in the environment is assumed to be 1–2% in production and 1–10% in use (Forgacs et al., 2004). Besides the unpleasant appearance of the dyepolluted wastewater, most dyes and their potential breakdown products are toxic. Therefore, the dyes pose an environmental threat. Microbial decolorization is an environment-friendly and costcompetitive alternative to chemical decomposition process. Much of the work undertaken in biodecolorization has involved the azo

q Statement of the scientific relevance: Anthraquinone based compounds are difficult to be biodegraded under aerobic conditions due to their fused aromatic structure. This paper presents a successful use of Sphingomonas herbicidovorans for biodegradation of 1-amino-4-bromoanthraquinone-2-sulfonic acid (bromoamine acid, BAA), which is a major intermediate of anthraquinone dyes harmful to aqueous environments due to its high water solubility. This is the first report on BAA biodegradation by S. herbicidovorans under aerobic conditions, although this strain has been used for degradation of phenoxyalkanoic acid herbicides in previous studies. * Corresponding author. Shenzhen Graduate School, Peking University, The Key Laboratory for Environmental and Urban Sciences, Guang Dong 518055, China. Tel.: þ86 10 62751185; fax: þ86 10 62756526. E-mail address: [email protected] (J. Ni). 0964-8305/$ – see front matter Ó 2008 Elsevier Ltd. All rights reserved. doi:10.1016/j.ibiod.2008.07.004

dyes (Brown, 1987; Banat et al., 1996; McMullan et al., 2001; Stolz, 2001; Pearcea, 2003; Forgacs et al., 2004), while there are limited reports on biodegradation of the anthraquinone dyes and their intermediates (Itoh et al., 1993; Walker and Weatherley, 2000; Chen et al., 2003; Dong et al., 2003; Qu et al., 2005a; Ren et al., 2006; Xu et al., 2006). Anthraquinone dyes are the second most common class of synthetic dyes after azo dyes, and they are highly resistant to decolorization under aerobic conditions due to their fused aromatic structure (Itoh et al., 1993). BAA is a major synthetic intermediate of the acid and reactive anthraquinone dyes. Its high water solubility permits its persistence in aqueous environments. Approximately 20 cubic meters of wastewater are discharged per ton of BAA produced, whereas the traditional activated sludge treatment proved to be ineffective and the current physical or chemical treatments uneconomical. Therefore, screening of efficient BAA-degrading bacterial strains and application of bioaugmented reactors (Qu et al., 2005b) have become popular research fields. Several bacterial strains have been isolated which can decolorize BAA such as Flavobacterium BX26 (Xin et al., 2000), Zoogloea HP3 (Huang et al., 2001) and Sphingomonas xenophaga (Li et al., 2003; Qu et al., 2005a), and some of them could grow with BAA as the sole source of carbon and nitrogen (Huang et al., 2001; Qu et al., 2005c). This paper deals with biodecolorization of BAA by a newly isolated strain of Sphingomonas herbicidovorans. The optimal decolorization and cell growth conditions were investigated, and the process of decolorization was analyzed.

L. Fan et al. / International Biodeterioration & Biodegradation 63 (2009) 88–92 2. Materials and methods 2.1. Chemicals and growth conditions BAA of commercial purity was obtained from the DanKong Industry & Trade Group Co., Ltd (Taizhou, Zhejiang, China) with an appearance of red acicular crystal. Phthalic acid was purchased from the Sigma–Aldrich Corp. (Saint Louis, Missouri, USA). Acetonitrile of chromatographical purity was purchased from the Merck Corp. (Darmstadt, Germany). All other chemicals were of analytical grade. Basal salts medium (BSM) was used for the isolation of the BAA-degrading strain. It contained (g l1): 2.2 Na2HPO4, 0.8 KH2PO4, 0.4 (NH4)2SO4, 0.01 MgSO4$7H2O, 0.01 FeSO4$7H2O, 0.005 CaCl2$2H2O. The pH of the medium was adjusted to 7.0. The medium was autoclaved at 121  C for 20 min. 2.2. Microbial strain The microbial strain designated as FL was isolated from a microbial consortium provided by the Department of Microbiology, Nankai University. The consortium was obtained from BAA-contaminated soil collected from a chemical plant by an enrichment culture technique using a mineral salts medium amended with BAA as the sole source of carbon (Li et al., 2003). The consortium was then inoculated on an agar plate containing BSM added with BAA, and the colonies were transferred to fresh agar plates several times in order to obtain a pure strain which showed consistent decolorization. The physiological and biochemical properties of the strain FL were characterized according to Bergey’s Manual of Determinative Bacteriology (Buchanan and Gibbons, 1984) by the Guangdong Detection Center of Microbiology (under ID: (2006) ZD0332). 2.3. Decolorization assays The strain FL was cultivated aerobically at 30  C in a shaker at 150 rpm. The stock cultures were prepared by growing a single colony in a conical flask (500 ml) containing 200 ml BSM with 500 mg l1 BAA and then subcultured 10% (v/v) to the fresh BSM with 100 mg l1 BAA. The optimal decolorization conditions were investigated under different pH (5.0–9.0), temperature (20–40  C) and shaking speed (0–150 rpm). The effect of initial concentration of BAA was determined under different concentrations (100–1000 mg l1). The effect of external carbon sources was investigated by adding 500 mg l1 glucose, sucrose and starch in the BSM containing 100 mg l1 BAA, while the cultivating solution without external carbon sources was the blank control. 2.4. Analytical methods During the reaction, samples were taken at regular intervals and the biomass concentration was measured spectrophotometrically at OD660. Furthermore, the corresponding bacterial dry weight could be obtained from a predetermined calibration curve describing the relationship between OD660 and dry weight. The samples were then centrifuged at 10,000 rpm for 10 min, and the concentration of BAA was measured at its maximum absorbance wavelength (485 nm) with the supernates. The UV–vis spectra of the supernates were scanned using a UV–vis spectrophotometer (Shimadzu UV1700, Japan). The metabolites of BAA decolorization were investigated using HPLC (Agilent 1200, USA) directly and GC/MS (Agilent 6890N/5975, USA) after extraction. A ZORBAX Eclipse XDB-C18 column (150  4.6 mm) was used as the separation column during the HPLC analysis. Gradient elution was used with a mobile phase consisting of acetonitrile (A) and 0.01 mol l1 ammonium acetate aqueous solution (B). The gradient elution started after holding at 0% A for 1 min, and increased linearly to 40% A over 2 min and held 2 min, and then increased to 100% A over 1.5 min and held 3.5 min. The flow rate was 1.0 ml min1 and the oven temperature was set at 30  C. The detection was carried out at the wavelength of 240 nm. Samples for GC/MS analysis were acidified to pH 2.0 and extracted three times with ethyl acetate. The extracts were combined and concentrated by evaporation in a water bath at 40  C. The prepared samples were analyzed by GC/MS equipped with HP-35 capillary column (30 m  250 mm  0.25 mm) in the EI mode (70 eV). The infrared spectra were obtained for BAA and the end product of its decolorization by FT-IR spectrophotometer (SHIMAZU 8400, Japan). After the degradation of 1000 mg l1 BAA, the supernates were filtrated through 0.45 mm membrane and freeze-dried under vacuum. A quantity of 2.0 mg of each sample was compressed with KBr, and the pellets obtained were analyzed covering a frequency range of 7800–350 cm1.

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denaturation period of 5 min at 94  C; 28 cycles of 94  C for 30 s, 57  C for 45 s, and 72  C for 2 min; 72  C for 10 min; and incubation at 4  C until further processing. The PCR amplicon was purified with agarose gel DNA purification kit and TA cloned onto pMD 18-T vector (TaKaRa Biotechnology, Dalian, Shandong, China). The ligated DNA was introduced into Escherichia coli JM109 cells by the standard transformation method (Sambrook and Russel, 2001). Plasmids were extracted by alkaline lysis method. After checking the sequence heterogeneity by PCR amplification, the nucleotide sequence of the 16S rRNA gene clone was determined by the SinoGenoMax Co., Ltd (Beijing, China). The nucleotide sequence was compared with the sequences in the GenBank nucleotide sequence databases by the BLASTN program (http://www.ncbi.nlm.nih. gov/BLAST/). The sequences were aligned by the Clustal X 1.8 program, and the phylogenetic analysis was performed with the neighbor-joining method (MEGA 3.1). The reference nucleotide sequences used in the phylogenetic tree construction were obtained from the GenBank nucleotide sequence database.

3. Results and discussion 3.1. Characteristics and phylogenetic analysis of the microbial strain A pure strain which could grow with BAA as the sole source of carbon was obtained from soil samples of a chemical plant. It is a mobile Gram-negative, rod-shaped, yellow-pigmented aerobic bacteria, 0.8–1.3 mm long and 0.5–0.6 mm wide. This strain forms rounded and mucoid colonies when cultivated on solid BAA–BSM medium. It is oxidase and catalase positive, but ornithine decarboxylase and lysine decarboxylase negative. It ferments glucose, maltose, lactose, and sucrose but not inositol. It lyses esculin but not gelatin. It does not use nitrate as an electron acceptor, does not utilize acetate, and does not produce indole when cultivated in the peptone solution. A 1457 bp amplicon was obtained by 16S rDNA PCR amplification. Its sequence was deposited in GenBank with the accession number EF065102. Phylogenetically, the genus Sphingomonas clusters within the alpha subclass of the Proteobacteria. On the basis of 16S rRNA gene sequence comparisons, the selected strains showed high homology to the strain FL, among which S. herbicidovorans (AB022428) showed 99% homology. Fig. 1 shows the phylogenetic tree based on 16S rRNA gene sequence comparisons.

2.5. 16S rRNA gene cloning, nucleotide sequencing, and phylogeny Genomic DNA was rapidly prepared by the boiling method (Sambrook and Russel, 2001). The 16S rDNA PCR amplifications were performed by using primers 27F (50 -AGAGT TTGAT CATGG CTCAG-30 ) and 1492R (50 -TACGG TTACC TTGTT ACGAC TT-30 ). The final 50 ml reaction mixture contained 0.5 mM each primer, 200 mM each dNTP, 10–70 ng genomic DNA, PCR buffer (50 mM KCl, 10 mM Tris–HCl and 1.5 mM MgCl2) and 0.1 U Pyrobest DNA Polymerase. The PCR protocol included an initial

Fig. 1. Unrooted phylogenetic tree based on 16S rRNA gene sequence comparisons.

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Fig. 2. Effects of temperature, pH and shaking speed on decolorization (column) and cell growth (line).

The genus Sphingomonas is becoming increasingly interesting in environmental microbiology because various xenobiotic-degrading organisms belong to this group. Previously, Sphingomonas strains have been described to degrade compounds such as biphenyl, naphthalene, fluorene, phenanthrene, pyrene, diphenylether, furan, carbazole, polyethylene glycols, and chlorinated phenols (Basta et al., 2004). The strain S. herbicidovorans has been used for degradation of phenoxyalkanoic acid herbicides in previous studies (Zipper et al., 1996; Kohler, 1999). 3.2. Effects of temperature, pH and shaking speed on decolorization The effects of temperature, pH and shaking speed on both decolorization and biomass concentration are shown in Fig. 2. Maximum decolorization percentage of about 98% was observed at 30  C and pH 7.0 within 14 h. The strain FL could decolorize BAA under a relatively low temperature of 20  C showing a decolorization percentage of 59.4%. However, the decolorization percentage decreased rapidly with increasing temperature, e.g. from 37.2% at 35  C to 9% at 40  C. The inhibition of decolorization at higher temperature was presumably due to the growth suppression of the cell. The strain FL grew between 20 and 40  C, however, beyond 41  C no growth was observed. The decolorization activity was high over a narrow pH range around 7.0. The decolorization percentage dropped by about 31.2% at pH 6.0 and 64.4% at pH 8.0, which indicated that the strain was more sensitive to the alkaline condition. BAA utilizing microorganisms were reported to grow and degrade BAA best at neutral pH and 30  C (Huang et al., 2000; Li et al., 2003; Qu et al., 2005a). With the increase of shaking speed, higher decolorization percentage and biomass concentration were obtained. However, there was little difference between the decolorization percentages at 100 and 150 rpm, both of which were above 98% within 14 h. It was indicated that the oxygen concentration under 100 rpm was enough to maintain high activities of decolorization enzyme. Moreover, decolorization also occurred under static condition with the percentage of 69.9%, which suggested the strain FL did not need much oxygen to survive.

Fig. 3. Effect of BAA concentration on specific decolorization rate.

decolorization rate of BAA per unit biomass (specific decolorization rate) at the exponential growth phase (v) increased with the elevation of initial BAA concentration, and the v of 100 and 1000 mg l1 BAA was 191.20 and 807.46 mg BAA g cell1 h1, respectively. The BAA decolorization kinetics could be described by the Monod equation (v ¼ vmax S=KS þ S), where vmax is the maximum specific decolorization rate (mg BAA g cell1 h1), S is the BAA concentration (mg l1) and Ks is the half-velocity coefficient for decolorization (mg l1). Fig. 3 shows a close fit between the experimental and predicted results (R2 ¼ 0.99), which implied that BAA of the tested concentration did not have an inhibitory effect on decolorization. The vmax and Ks estimated from the experimental data was 1144.74 mg BAA g cell1 h1 and 408.71 mg l1, respectively. Huang et al. (2000) described similar tolerance for BAA up to 1000 mg l1 of Zoogloea HP3. Comparatively, the study with S. xenophaga QYY showed that BAA with an initial concentration over 654.0 mg l1 had an inhibitory effect, and the vmax and Ks estimated from the Haldane model was 335.96 mg BAA g cell1 h1 and 299.42 mg l1 (Xing et al., 2006). 3.4. Effect of external carbon sources on decolorization The decolorization rate could be enhanced by the external carbon sources, of which glucose was the best for such a purpose

3.3. Decolorization kinetics under different initial BAA concentrations The strain FL could tolerate high concentration of BAA, and the decolorization percentages of different initial concentrations were all above 98% within 24 h. With the increase of initial BAA concentration, the cell grew faster. The maximum biomass concentration of 100 and 1000 mg l1 BAA could reach 36.7 and 114.6 mg l1, respectively. With the restriction of biomass concentration, the reaction time needed to the termination of 100 mg l1 BAA was longer than that of 200 mg l1 BAA. The average

Fig. 4. Effect of external carbon sources on decolorization.

L. Fan et al. / International Biodeterioration & Biodegradation 63 (2009) 88–92

Fig. 5. UV–vis spectra during the decolorization of BAA (initial BAA 100.0 mg l1).

(Fig. 4). The decolorization percentage of the test group supplied with glucose could reach 98% within 4 h. The experiment also revealed the enhanced cell growth, which indicated that glucose, starch, and sucrose are ideal carbon sources for the growth of the strain. When the strain FL was inoculated, it preferred to utilize glucose, starch and sucrose as carbon sources and energy, and the decolorization enzyme was induced by BAA simultaneously. Since the biomass concentration was enhanced, the secretion of decolorization enzyme increased and the decolorization of BAA was accelerated. Furthermore, the external carbon sources were not substrates of the decolorization enzyme, and did not have competitive inhibition effect towards BAA decolorization. Huang et al. (2000) also reported the enhancement of BAA decolorization by Zoogloea HP3 using glucose, sucrose, fructose, xylose and mannose. 3.5. Analysis of the decolorization process The color of reaction mixture changed from red to colorless as a function of time. The contribution of biosorption on the removal of BAA was neglectable, since the biomass concentration was low in the BSM and the color of the cell pellets was their inherent yellow at the end of the reaction. Fig. 5 shows that during the decolorization of BAA, the absorbance peak at 485 nm disappeared completely after 14 h cultivation, and there was a significant decrease between 230 and 300 nm. The great changes occurring both in UV and visible spectra indicated that the molecular structure of BAA changed evidently after decolorization. However, the UV–vis absorbance spectra of the blank control which was not added with the strain FL did not change throughout the test period.

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The red color of BAA was caused by the conjugated structure of anthraquinone nuclear and amino group. Since the strain FL grew with BAA as the sole carbon source, it could be presumed that the anthraquinone nuclear cleaved during the reaction and the primary chromophore was destroyed. The absorbance peak at UV spectra did not disappear in the end of decolorization, which indicated that BAA was not completely mineralized. The HPLC analysis (Fig. 6) shows the general decrease of the BAA (tR ¼ 6.2 min) concentration with incubation time in the process of decolorization, accompanied with two metabolites produced. Metabolite ‘‘1’’ was an intermediate and its retention time, tR ¼ 1.7 min, was the same as that of phthalic acid. Further analysis by GC/MS demonstrated its agreement with phthalic acid in terms of mass spectra (m/z 148, 104, 76, 50). During the degradation of 892.2 mg l1 BAA, phthalic acid was produced and then experienced two variation stages, i.e. firstly increased from zero to a maximum of 16.2 mg l1 after 18 h and then decreased. Furthermore, metabolite ‘‘2’’ (tR ¼ 4.7 min) was proved as the end product. During the degradation of BAA, the concentration of metabolite ‘‘2’’ continuously increased before it reached to an unchanged value. However, the end product and BAA were not detected by GC/MS probably due to their nonvolatility. Fig. 7 shows the infrared spectra of BAA and the end product. The significant infrared peaks of BAA were due to the carbonyl (C]O) stretch at 1670 cm1, the amino (N–H) stretches at 3286.5 and 3379.1 cm1, the C–Br stretch at 624.9 cm1, as well as the S]O stretches between 1014.5 and 1238.2 cm1 (Yang, 1987). Other peaks are related to the breathing mode of anthraquinone ring at 1527 and 1593.1 cm1, and the C–H deformation mode of anthraquinone ring at 721.3 and 740.6 cm1. The infrared spectrum of the end product was influenced by the existence of salt in the sample to some extent. However, the loss of the carbonyl vibrational peak at 1670 cm1, and the appearance of the skeleton stretching peaks of benzene at 1488.9, 1577.7 and 1620.1 cm1 suggested that BAA was ring-cleaved and the chromophore destroyed. The end product probably had amino, bromo, hydroxyl and sulfonate groups, since the related vibrational peaks of N–H (3332.8, 3417.6 cm1), C–Br (624.9 cm1), O–H (3541.1 cm1), and S]O (1041.5–1230.5 cm1) were observed. The existence of the peak at around 860.2 cm1 owing to the aromatic C–H deformation supported the assumption that the end product was multi-substituted benzene derivative. Further degradation of the final product by the strain FL proved to be difficult, perhaps due to the molecular steric effect, which inhibited the contact between the enzymes and the substrates. Previous studies have shown that fungi can degrade anthraquinone derivatives through the phthalate route (Hammel, 1991; Itoh, 1998). In the present study, the detected phthalic acid as the only intermediate in the reaction mixture of BAA degradation indicated that BAA might be attacked by the oxygenase on the carbonyl carbon and ring-cleavaged, releasing phthalic acid and an end product. More studies are needed to clarify the degradation mechanism of BAA.

Fig. 6. HPLC analysis of metabolites of BAA decolorization (initial BAA 892.2 mg l1).

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References

Fig. 7. FT-IR analysis of BAA and the end product (a: BAA; b: the end product).

4. Conclusions This study demonstrated the ability of a new strain of S. herbicidovorans to decolorize BAA and grow with it as the sole source of carbon. The idealized decolorization occurred at temperature of 30  C and pH 7.0. The decolorization was robust even under high concentration of BAA up to 1000 mg l1. External carbon sources such as glucose could enhance the decolorization rate. The primary mechanism for decolorization was the ring cleavage of BAA through the phthalate route. However, BAA was not completely mineralized and one end product was formed which might be multi-substituted benzene derivative. Acknowledgements This work was supported by the National Basic Research Program of China under grant No. 2006BAB04A14 and the Presidential Foundation of the Shenzhen Graduate School of Peking University under grant No. 2006013. We are grateful to Professor B.L. Cai (Nankai University, China) for his help to isolate the strain. Sincere thanks are also to Professor A.G.L Borthwick (University of Oxford, UK) for his English editing of the manuscript.

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