Biochemical characterization of a novel azoreductase from Streptomyces sp.: Application in eco-friendly decolorization of azo dye wastewater

International Journal of Biological Macromolecules 140 (2019) 1037–1046

Contents lists available at ScienceDirect

International Journal of Biological Macromolecules journal homepage: http://www.elsevier.com/locate/ijbiomac

Biochemical characterization of a novel azoreductase from Streptomyces sp.: Application in eco-friendly decolorization of azo dye wastewater Hao Dong a, Tianyuan Guo a, Wenxue Zhang a, Hanjie Ying b, Ping Wang c, Yibing Wang a,⁎, Yong Chen b,⁎⁎ a State Key Laboratory of Bioreactor Engineering, Biomedical Nanotechnology Center, Shanghai Collaborative Innovation Center for Biomanufacturing Technology, School of Biotechnology, East China University of Science and Technology, Shanghai 200237, China b National Engineering Research Center for Biotechnology, College of Biotechnology and Pharmaceutical Engineering, Nanjing Tech University, Nanjing 211816, China c Department of Bioproducts and Biosystems Engineering, University of Minnesota, St Paul, MN 55108, USA

a r t i c l e

i n f o

Article history: Received 1 July 2019 Received in revised form 21 August 2019 Accepted 22 August 2019 Available online 23 August 2019 Keywords: Azoreductase Whole-cell biocatalyst Coenzyme regeneration

a b s t r a c t Azo dyes are the most widely applied chemical dyes that have also raised great concerns for environmental contamination and human health issues. There has been a growing interest in discovering bioremediation methods to degrade azo dyes for environmental and economic purposes. Azoreductases are key enzymes evolved in nature capable of degrading azo dyes. The current work reports the identification, expression, and properties of a novel azoreductase (AzoRed2) from Streptomyces sp. S27 which shows an excellent stability against pH change and organic solvents. To overcome the requirements of coenzyme while degrading azo dyes, we introduced a coenzyme regeneration enzyme, Bacillus subtilis glucose 1-dehydrogenase (BsGDH), to construct a recycling system in living cells. The whole-cell biocatalyst containing AzoRed2 and BsGDH was used to degrade a representative azo dye methyl red. The degradation rate of methyl red was up to 99% in 120 min with high substrate concentration (250 μM) and no external coenzyme added. The degradation rate was still 98% in the third batch trial. To sum up, a novel azoreductase with good properties was found, which was applied to construct whole-cell biocatalyst. Both the enzymes and whole-cell biocatalysts are good candidates for the industrial wastewater treatment and environmental restoration. © 2019 Published by Elsevier B.V.

1. Introduction Wastewater generated from traditional dyeing industries is a major source of environmental contamination, seriously threatening native living organisms and causing human health risks [1–3]. Synthetic dyes are widely used in industries such as paper, leather, cosmetics and pharmaceuticals [4,5]. Over 10 million tonnes of dyes are produced and applied annually [6–9]. It is estimated that N50% of all dyes are azo dyes, compounds that can be expressed as R-N=N-R' (where -N=N- is the azo group, and the R or R' can be either aryl or alkyl compounds) [10,11]. Azo dyes are particularly desired in the textile industry [12]. Many azo dyes are considered highly toxic and carcinogenic. For example, benzidine, the decomposition product of azo dyes, has been

⁎ Correspondence to: Y. Wang, School of Biotechnology, East China University of Science and Technology, No. 130 Meilong Road, Xuhui District, Shanghai 200237, China. ⁎⁎ Correspondence to: Y. Chen, National Engineering Research Center for Biotechnology, Nanjing Tech University, NO. 30 Puzhu Road, Nanjing 211816, China. E-mail addresses: [email protected] (Y. Wang), [email protected] (Y. Chen).

https://doi.org/10.1016/j.ijbiomac.2019.08.196 0141-8130/© 2019 Published by Elsevier B.V.

reported to be associated with human bladder and kidney cancers [13–15]. A variety of approaches have been developed to treat dyecontaminated wastewater. They can be largely classified as physical, chemical and biological methods [3,16]. Physical methods apply technologies such as adsorption, coagulation-flocculation, membrane filtration. Chemical treatment may include chemical oxidation, electrochemistry, ion-exchange, chemical precipitation, catalytic degradation. Biological method, or bioremediation, applies microorganisms that can uptake and degrade dyes from water. Although physical and chemical methods can effectively remove dyes from wastewater, they can be expensive and may involve formation of secondary hazardous wastes [17]. In recent years, bioremediation has drawn growing interests since it promises high performance, lower cost, and environmentally friendly. Recent studies on degradation of azo dyes have revealed promising enzymes found in fungi and bacteria [18–20]. Azo dyes are stable in nature and hard to degrade only if the azo linkage is first reduced [21]. Azoreductase (EC 1.7.1.6) is the key enzyme expressed in azo-dyedegrading microorganisms to catalyze the reductive cleavage of azo bond under the ping-pong mechanism [22,23]. Based on the cofactor

1038

H. Dong et al. / International Journal of Biological Macromolecules 140 (2019) 1037–1046

requirements, azoreductases are categorized as either flavin-dependent or flavin-independent [24]. Each of that is further divided into subgroups according to their preference to NADH or NADPH utilized as the electron donor. The requirement of coenzyme (NADH or NADPH) for enzymatic degradation of azo dyes is generally a limiting factor for its application in wastewater treatment [25]. One way to overcome this problem is to introduce in situ coenzyme regeneration by incorporating NAD(P)H regeneration enzymes. The most common used enzymes for coenzyme regeneration are glucose dehydrogenases (GDHs) and formate dehydrogenases (FDHs) [26,27]. Yang et al. constructed an integrated enzyme system containing azoreductase and glucose 1-dehydrogenase, which showed efficient dye removal [28]. Rathod et al. used the formate dehydrogenase fdh as the coenzyme regeneration enzyme to reduce NAD+ in Escherichia coli (E. coli), and the whole-cell biocatalyst showed 30% decolorization toward violet 5R (RV5R) in 12 h [20]. Compared to GDHs, FDHs usually exhibits low enzymatic activity and stability, which limits its wide use in the coenzyme regeneration system [28,29]. In addition to the requirement of coenzymes, low substrate tolerance of the enzyme systems could also be substantially limiting [30,31]. At present, most of the identified azoreductases have low substrate tolerance and low stability, which restricts their use in wastewater treatment. Thus, screening a novel azoreductase with higher substrate tolerance and higher stability toward environment change can fulfill its wide use in azo dye wastewater treatment from different sources. In this work, a novel azoreductase (AzoRed2) from Streptomyces sp. S27 was identified and showed good stability against harsh conditions. By constructing a coupled system consisting of AzoRed2 and Bacillus subtilis glucose 1-dehydrogenase (BsGDH), the whole-cell biocatalysts for degradation of azo dyes (specifically, methyl red as a model dye) is examined. Our study demonstrates that AzoRed2 is a novel azoreductase with excellent properties, which could be a promising candidate to degrade azo dyes efficiently in the industry wastewater. 2. Materials and methods 2.1. Materials Phanta super-fidelity DNA polymerase was purchased from Vazyme Biotech (Nanjing, China). Restriction enzymes (EcoRI, HindIII, XhoI) were from New England BioLabs (Ipswich, MA, USA). T4 DNA ligase was purchased from TAKARA (Japan). A genomic DNA extraction kit was purchased from QIAGEN (Germany). Other chemicals and reagents were purchased from local markets. 2.2. Screening and sequence analysis of azoreductase genes Through searching of the complete genome sequence of Streptomyces sp. S27 (unpublished), a gene (named azored2) coding a putative azoreductase was found. A phylogenetic tree of AzoRed2 and other azoreductases was constructed using MEGA 7.0 software. Multiple sequence alignments were carried out using the Clustal X program and exported using ESPript 3.0 (http://espript.ibcp.fr/ESPript/ESPript/).

using standard methods [32]. In this section, the expression vector and host strain were pET28a(+) and BL21(DE3), respectively. The constructed recombinant plasmid was sent to be sequenced (Sangon Biotech). E. coli BL21(DE3) containing the recombinant plasmid (pET28aazored2) was cultivated at 37 °C in 100 mL LB liquid medium (+50 μg/mL Kanamycin) until the OD600 reached 0.4–0.7. Isopropyl-β-D-1thiogalactopyranoside (IPTG) with a final concentration of 100 μM was added to the medium at 20 °C for 20 h to induce the AzoRed2 protein expression. The cells were harvested and washed with PBS buffer (pH 7.4) once, and then the cells were resuspended in 10 mL PBS buffer (pH 7.4). Induced cells were ruptured by sonication. After centrifugation (8000 ×g) at 4 °C for 15 min, the supernatant of the cell lysis solution was loaded onto a Ni-NTA column (1 mL, GE Healthcare, USA). Unbound proteins were washed with PBS buffer (pH 7.4) containing 20 mM imidazole. The bound AzoRed2 protein was eluted using a linear imidazole gradient (20–500 mM) in PBS buffer (pH 7.4). Finally, the purified AzoRed2 was analyzed by SDS-PAGE. 2.4. Enzyme characterization of AzoRed2 The azoreductase activity of AzoRed2 was determined using the method reported by Nakanishi et al. with minor modification [23]. Briefly, the assay system contained 25 μM methyl red, 250 μM NADH, and 5 μM FMN in phosphate buffer (100 mM) at 30 °C. The reaction was initiated by adding the AzoRed2 protein and was terminated by adding SDS with a final concentration of 1.0%. The decrease in the value of the absorbance at 430 nm was recorded. One unit of azoreductase activity was defined as the amount of enzyme required to degrade 1 μmol methyl red per minute. The molar adsorption coefficient of methyl red is 23.36 mM−1 cm−1. 2.4.1. Cofactor identification The non-covalently bound flavin in AzoRed2 was identified using UV–vis adsorption spectra. Native purified AzoRed2 protein was used as a control and was further treated using 1.0% SDS (final concentration) to compare the changes in absorption peaks [33]. To determine the stoichiometric binding of AzoRed2 with FMN molecules, the purified AzoRed2 was incubated with excess amount of FMN at 4 °C for 30 min. After removing the unbound FMN by filtering through a Millipore Amicon Ultra 10,000 cutoff filter, the contents of protein and protein-bound FMN molecules in reconstituted AzoRed2-FMN complex were quantified. 2.4.2. Coenzyme preference determination Azoreductases use NADH or NADPH as a coenzyme to degrade azo dyes, and they may exhibit preference toward NADH and NADPH. Based on the initial reaction system, NADH was replaced by NADPH to determine the coenzyme preferences of AzoRed2.

2.3. Heterologous expression and purification of azoreductase AzoRed2

2.4.3. Optimal temperature and thermostability To determine the optimal reaction temperature of AzoRed2, the reaction solution was incubated at temperatures ranging from 25 °C to 70 °C. The thermostability of AzoRed2 was determined by incubating AzoRed2 at different temperatures (25 °C–70 °C) for 1 h, and the residual activity was determined.

The Streptomyces sp. S27 stored at −20 °C in our laboratory was inoculated into the liquid medium (KNO3, 0.1%; K2HPO4, 0.05%; MgSO4·7H2O, 0.05%; NaCl, 0.05%; FeSO4·7H2O, 0.01%; Starch, 2.0%) and incubated at 37 °C with rotary shaking (180 rpm) for 24 h. The genomic DNA of S27 was extracted following the instructions of the genomic DNA extraction kit (QIAGEN, Germany). According to the nucleotide sequence of predicted azoreductase azored2, one pair of full-length primers was synthesized (azored2_F and azored2_R, Table S1). The recombinant plasmid was constructed

2.4.4. Optimal pH and pH stability The optimal reaction pH was studied over pH values ranging from 4.0 to 8.0 in different buffers (4.0–5.0, Na2HPO4-Citric acid buffer; 5.0–6.0, Sodium citrate buffer; and 6.0–8.0, Phosphate buffer). The procedure for examining pH stability was carried out by incubating the AzoRed2 protein in different pH buffers (pH 4.0 Na2HPO4-Citric acid buffer, pH 5.0 Sodium citrate buffer, pH 6.0 Sodium citrate buffer, pH 7.0 Phosphate buffer, pH 8.0 Tris-HCl buffer, pH 9.0 Tris-HCl buffer) at 25 °C for hours, and the residual activity was determined.

H. Dong et al. / International Journal of Biological Macromolecules 140 (2019) 1037–1046

1039

Scheme 1. Construction of recombinant plasmid coupled AzoRed2 and BsGDH.

2.4.5. Substrate specificity Substrate specificity was determined using different azo dyes (Congo Red, Methyl Orange, Trypan Blue, Sudan Black B, and Direct Black 38), and methyl red was used as a control (activity defined as 100%). 2.4.6. Effect of detergents on AzoRed2 activity The effect of detergents on AzoRed2 activity was determined by adding 0.5% or 1.0% of various detergents (Tween 20, Tween 80, Triton X-100, SDS) to the reaction system. The group without adding any detergents was used as a control, and its catalytic activity was defined as 100%. 2.4.7. Effect of metal ions on AzoRed2 activity To estimate the influence of metal ions on AzoRed2 activity, the reaction solution was mixed with different metal ions (Ni2+, Mg2+, Ca2 + , Mn2+, Cu2+, Zn2+, Co2+, Fe2+, and Fe3+) and the chelating agent Na2-EDTA. The activity of the solution without adding metal ions or Na2-EDTA was defined as 100%. 2.4.8. Effect of organic solvents on AzoRed2 activity To estimate the effect of organic solvents on AzoRed2 activity, different organic solvents (DMSO, methanol, ethanol, acetonitrile, isopropanol, n-propanol, n-butanol, isoamyl alcohol, chloroform, nhexane, and isooctane) were mixed with purified AzoRed2 solution, and the final organic solvent concentrations were 10% or 20%. The mixed solutions were incubated at 30 °C with rotary shaking (80 rpm)

for 1 h, and the residual activities were finally determined. In order to reduce the effect of organic solvents on the determination of azoreductase activity, the hydrophobic organic solvents were removed by centrifugation from the mixture after incubation [34,35]. For hydrophilic organic solvents, the concentration of organic solvents was diluted to a low content (5%) before determining the azoreductase activity [34,35]. 2.4.9. Specific activity of AzoRed2 Under the optimal reaction conditions (25 μM methyl red, 250 μM NADH, and 5 μM FMN in Na2HPO4-Citric acid buffer (pH 5.0, 100 mM) at 55 °C), the activity of AzoRed2 was determined. The protein content was assayed according to the method of Bradford using bovine serum albumin as the standard protein. The relative activity was defined as the total azoreductase activity per mg protein. 2.5. Construction of a co-expression plasmid and co-expression of AzoRed2 and BsGDH The co-expression plasmid (pET28a-azored2-bsgdh) containing the azored2 gene and the B. subtilis gdh gene (WP_003246720.1) was constructed as described in Scheme 1 [27]. A pair of oligonucleotide primers (sd-as-bsgdh_F and sd-as-bsgdh_R, Table S1) was designed and synthesized. The PCR product of sd-as-bsgdh was double digested and then ligated to the plasmid pET28a-azored2. The recombinant BL21(DE3) strain containing the co-expression pET28a-azored2-bsgdh plasmid was used to express AzoRed2 and BsGDH as described above. SDS-PAGE was used to determine the expression levels of AzoRed2 and BsGDH.

1040

H. Dong et al. / International Journal of Biological Macromolecules 140 (2019) 1037–1046

Fig. 1. Bioinformatic analysis of AzoRed2. (a) Neighbor-joining phylogenetic tree. Family I, FMN-dependent NADH preference azoreductase; Family II, FMN-dependent NADPH preference azoreductase; Family III, flavin-free azoreductase. (b) Multiple sequence alignments between AzoRed2 and other azoreductases from Family I.

The azoreductase activity (NADH as the coenzyme) of the cell lysate supernatant was determined as described above. The GDH activity (NAD+ as the coenzyme) of the cell lysate supernatant was assayed as in the report described [27]. The coupling enzymatic activity of the supernatant toward methyl red (NAD+ as the coenzyme) was also determined. After determining the protein content, the relative activities were calculated. 2.6. Methyl red degradation using the whole-cell biocatalyst The wet co-expression cells containing AzoRed2 and BsGDH were used to study the efficiency of degrading methyl red. The initial reaction system contained 2.5 mg/mL biocatalyst, 100 μM methyl red, 20 mM glucose, and pH 6.0 sodium citrate buffer (100 mM). The mixed reaction system was incubated under 30 °C with rotary shaking (80 rpm). The reaction solution was sampled, and the residual methyl red was quantified at various intervals, after which, the decolorization was determined. The decolorization of the solution represented the degradation rate of methyl red, and the decolorization determining formula was as follows: Residual methyl red Decolorization ¼ 1− Initial methyl red

2.6.1. Optimization of reaction conditions 2.6.1.1. Effect of additional concentration of NAD+. Based on the reaction conditions, the effect of additional NAD+ amounts was studied. The additional amounts were set as 0, 0.1, 0.2, 0.3, 0.4, 0.5, and 0.6 mM. 2.6.1.2. Effect of glucose concentration. To study whether glucose concentration had a significant effect on the degradation efficiency of methyl red, the glucose concentration was varied (5, 10, 20, 30, 40, and 60 mM), while other factors remained unchanged at the initial conditions. 2.6.1.3. Effect of biocatalyst loading. Under the initial reaction conditions, biocatalyst loading was further used as the variable to carry out the degradation reaction. The loading amounts of wet whole-cell biocatalysts were 2.5, 5.0, 7.5, 10.0, 12.5 and 15.0 mg/mL. 2.6.1.4. Effect of methyl red concentration. After evaluating the effects of additional amounts of NAD+, biocatalyst loadings, and glucose concentrations, the substrate tolerance of the whole-cell biocatalyst was tested under optimal conditions (no NAD+ addition, 5.0 mg/mL wet wholecell biocatalyst and 10 mM glucose). The concentrations of methyl red in the medium were increased to 125, 150, 175, 200, 225, and 250 μM.

H. Dong et al. / International Journal of Biological Macromolecules 140 (2019) 1037–1046

1041

Phylogenetic tree analysis was carried out using azoreductases from different families to classify AzoRed2. As shown in Fig. 1a, AzoRed2 showed amino acid sequence similarity to Family I azoreductases (FMN-dependent NADH preference azoreductase). Furthermore, multiple sequence alignment results (Fig. 1b) showed that AzoRed2 had the highly conserved FMN binding region but low similarity in the NAD (P)H binding region with other FMN-dependent NADH preference azoreductases. The changes in the NAD(P)H binding domains have been reported in other studies [36]. These results indicated that AzoRed2 may be a protein belonging to Family I. 3.2. Expression and purification of AzoRed2 The extracted genome DNA of Streptomyces sp. S27 (Fig. S1) was used as the template to amplify the azored2 gene. This gene was cloned into a pET28a(+) vector with a 6 × His tag at the N terminus (resulting in pET28a-azored2) and finally transformed to E. coli BL21(DE3) for protein expression. As shown in Fig. 2, the AzoRed2 protein was successfully expressed as a soluble fraction. Moreover, purified AzoRed2 protein was separated as a single band consistent with the predicted molecular weight (28.89 kDa). 3.3. Characterization of AzoRed2 Fig. 2. SDS-PAGE analysis of AzoRed2. Lane 0, protein marker; Lane 1, supernatant of AzoRed2 cell lysate; Lane 2, precipitation of AzoRed2 cell lysate; and lane 3, purified AzoRed2.

2.6.2. Batch biodegradation of methyl red using the whole-cell biocatalyst In this section, the wet whole-cell biocatalyst was used to determine its reusability, and the reaction system was as follows: 5.0 mg/mL biocatalyst, 250 μM methyl red and 10 mM glucose in pH 6.0 sodium citrate buffer (100 mM). The dye removal process was assessed during 2.0 h per batchwise cycle.

3. Results and discussion 3.1. Sequence analysis of azored2 Through genome searching of Streptomyces sp. S27, a putative azoreductase gene (named azored2) with 702 bp was identified. This gene codes a putative protein that had the highest sequence identity (77.1%) with FMN-dependent NADH-azoreductase from Streptomyces sp. XY431. These are all azoreductases that have not been expressed or characterized. The nucleotide sequence of azored2 was deposited in the GenBank database with the accession number MK894988.

Fig. 3. UV–vis adsorption spectra of native AzoRed2 and denatured AzoRed2.

3.3.1. Cofactor identification and coenzyme preference The flavin mononucleotide (FMN) dependence and NAD(P)H preferences are two important characteristics for azoreductase classification [37]. To identify the binding form of AzoRed2 and FMN, SDS was used to treat the AzoRed2 protein solution. The UV–vis result (Fig. 3) showed that a spectral blue-shift phenomenon occurred from 463 to 448 nm, which was attributed to the noncovalent bonds broken between FMN and the apoprotein [33,38]. To determine the contents of protein and FMN in AzoRed2-FMN complex, we measured the saturation of protein by FMN. As a result, the ratio between FMN and AzoRed2 was calculated to be 1.941, which suggested that 1 mol AzoRed2 could non-covalently bind with 2 mol FMN molecules. In the determination of AzoRed2 activity, excess amount of FMN was used to maintain AzoRed2 in the saturation form by FMN. Coenzyme preference of AzoRed2 was verified by using NADH or NADPH as electron donors. The results (not shown) indicated that AzoRed2 could use either NADH or NADPH as electron donors to catalyze the cleavage of the -N=N- bond, and the activities toward methyl red were nearly the same, which was different from Family I azoreductases (FMN-dependent NADH preference azoreductases). Combining the results of the phylogenetic tree analysis and coenzyme characteristics, we can conclude that AzoRed2 was a novel FMN-dependent azoreductase that had no preference toward NADH and NADPH. Considering economic factors, we chose NADH as the electron donor for the next experiments. 3.3.2. Effects of temperature and pH on enzymatic activity and stability The effect of temperature on the enzymatic activity of AzoRed2 was investigated over a broad range, from 25 °C to 70 °C. As shown in Fig. 4a, AzoRed2 displayed the maximum activity at 55 °C. The azoreductase activity increased linearly from 25 °C to 55 °C; however, the azoreductase activity decreased rapidly beyond 55 °C. The thermostability results for AzoRed2 (Fig. 4a) showed that after incubation at various temperatures, the enzymatic activity was nearly unchanged from 25 °C to 30 °C. AzoRed2 was not stable under higher temperatures, and when the temperature increased to 50 °C, the AzoRed2 lost all enzymatic activity after incubation. We chose 30 °C as the temperature for the whole-cell biocatalytic reactions due to the strong thermostability, which may be beneficial to more batch reactions. Fig. 4b shows that AzoRed2 displayed the maximum activity at pH values between 4.5 and 5.5, and the optimal pH was 5.0. AzoRed2 displayed relatively high activity under acidic conditions, which was a desirable property for wastewater treatment in the industry. The pH

1042

H. Dong et al. / International Journal of Biological Macromolecules 140 (2019) 1037–1046

Fig. 4. Characterization of purified AzoRed2. (a) Optimal reaction temperature and thermostability of AzoRed2. (b) Optimal reaction pH of AzoRed2. (c) pH stability of AzoRed2. (d) Substrate specificity toward different azo dyes.

stability results are shown in Fig. 4c. AzoRed2 was unstable in pH 4.0 and pH 5.0 buffers. The residual activity of AzoRed2 decreased to 5.23% in pH 4.0 Na2HPO4-citric acid buffer after 3 h and to 5.49% in sodium citrate buffer after 12 h. However, AzoRed2 was stable at pH ≥ 6.0. At pH 7.0, AzoRed2 had the highest stability, and after 30 h incubation, AzoRed2 retained 39.6% enzymatic activity. In Yang's report, the azoreductase had the highest stability in pH 6.5 sodium phosphate buffer, however, the activity decreased significantly in other buffers with incubation for only 1 h [39]. In our study, after incubation for 1 h, AzoRed2 retained over 89% initial activities with pH ranging from 6.0 to 9.0. Especially in pH 7.0 phosphate buffer, the activity of AzoRed2 stayed nearly unchanged. Even incubation for 6 h, AzoRed2 could retain over 75% initial activities in various buffers with pH ranging from 6.0 to 9.0. In another report, Lu et al. [29] constructed a fusion enzyme GFA which showed more fragile toward pH changes. GFA showed the highest stability in pH 7.0 sodium phosphate buffer, however, the degradation activity of GFA toward methyl red highly decreased in other buffers after incubation for only 1 h. The residual activities of GFA were around 50%, 70%, and 20% in pH 5.0, pH 6.0, and pH 8.0 buffers,

respectively. From the above discussion, it could be concluded that AzoRed2 had a good tolerance toward pH changes. 3.3.3. Substrate specificity In order to determine the substrate specificity of AzoRed2, we tested its enzymatic activities toward different azo dyes. The results (Fig. 4d) showed that AzoRed2 had the highest activity toward methyl red, and the activity toward methyl orange was 12.07% compared with methyl red. AzoRed2 had relatively low activities toward trypan blue, sudan black B, and direct black 38, and no activity toward Congo red. The relatively low enzymatic activities might be attributed to the larger molecular weight of trypan blue, sudan black B, direct black 38, and Congo red. The relative activity of purified AzoRed2 was 1.173 U·mg−1 toward methyl red, which was higher than the reported azoH value (0.27 U·mg−1) [18] and lower than the reported AzoRo value (141.25 U·mg−1) [40].

Table 2 Effect of metal ions on AzoRed2 activity. Ion

Table 1 Effect of detergents on AzoRed2 activity. Detergents

Control Tween 20 Tween 80 Triton X-100 SDS a

ND not detectable.

Relative activity (%) 0.5%, m/v

1.0%, m/v

100 ± 3.17 84.90 ± 2.10 84.44 ± 2.06 65.90 ± 4.95 NDa

100 ± 6.85 72.21 ± 3.89 67.43 ± 4.00 56.26 ± 5.52 ND

Control Na2-EDTA Ni2+ Mg2+ Ca2+ Mn2+ Cu2+ Zn2+ Co2+ Fe2+ Fe3+

Relative activity (%) 1 mM

10 mM

100 ± 4.27 101.61 ± 8.53 94.62 ± 5.18 100 ± 3.23 97.31 ± 4.93 89.78 ± 8.28 39.25 ± 6.72 88.71 ± 7.03 101.08 ± 4.93 67.20 ± 7.62 80.11 ± 2.46

100 ± 5.67 96.73 ± 5.61 78.04 ± 5.67 97.66 ± 1.62 83.18 ± 6.62 69.63 ± 8.21 6.07 ± 0.81 44.86 ± 1.40 71.03 ± 2.14 15.42 ± 4.21 67.76 ± 2.92

H. Dong et al. / International Journal of Biological Macromolecules 140 (2019) 1037–1046

1043

Table 3 Effect of organic solvents on AzoRed2 activity. Organic solvents

Control DMSO Methanol Ethanol Acetonitrile Isopropanol n-propanol n-butanol Isoamyl alcohol Chloroform n-hexane Isooctane

log P

– −1.3 −0.76 −0.24 −0.15 0.1 0.28 0.88 1.02 2.0 3.5 4.5

Residual activity (%) at concentration (%, v/v) of 10

20

100 ± 0.46 85.28 ± 3.54 80.98 ± 12.07 91.41 ± 5.62 48.47 ± 9.44 87.73 ± 7.66 84.05 ± 3.83 13.06 ± 2.59 13.06 ± 3.93 10.82 ± 1.29 103.73 ± 3.42 69.03 ± 5.29

100 ± 1.47 100 ± 6.25 100 ± 5.29 105.93 ± 15.53 17.80 ± 9.17 109.32 ± 2.54 23.73 ± 1.47 6.27 ± 3.77 9.76 ± 1.60 12.20 ± 2.18 92.33 ± 3.96 91.99 ± 2.09

3.3.4. Effects of detergents, metal ions, and organic solvents Different detergents (0.5% and 1.0%) all displayed inhibitory effects on the enzymatic activity of AzoRed2 (Table 1). SDS, an ionic detergent, inactivated AzoRed2 activity completely. In other words, no azoreductase activity was determined when adding SDS to the reaction system. This high inactivation effect of SDS to enzymatic activity has been reported in the literatures [34,39]. With the concentration of other detergents (Tween 20, Tween 80, and Triton X-100) increased from 0.5% (m/v) to 1.0% (m/v), the inhibitory effects became stronger. And they decreased the AzoRed2 activity by 15–34% at 0.5% (m/v) and 28–44% at 1.0% (m/v). Wastewater always contains various metal ions, and these might influence the activity of AzoRed2 toward azo dyes [29,41]. In order to detect the metallic dependence of AzoRed2, Na2-EDTA was used to combine the possible binding metal ions in enzymes. As shown in Table 2, Na2-EDTA (1 and 10 mM) had little effect on AzoRed2 activity, meaning that AzoRed2 was a nonmetallic dependent enzyme. Mg2+ (1 and 10 mM) also had little effect on AzoRed2 activity. Co2+ had almost no effect on AzoRed2 activity at 1 mM; however, AzoRed2 enzymatic activity decreased to 71.03% at 10 mM Co2+. Cu2+ had the highest inhibitory effect on AzoRed2 activity, and the relative activity was 39.25% at 1 mM and 6.07% at 10 mM. Other ions (Ni2+, Ca2+, Mn2+,

Fig. 5. SDS-PAGE analysis of the coupled system co-expressing AzoRed2 and BsGDH. Lane 1, supernatant of cell lysate; Lane 2, precipitation of cell lysate; and lane 3, protein marker.

Zn2+, Fe2+, and Fe3+) decreased the AzoRed2 activity by 3–23% at 1 mM and 17–85% at 10 mM. Often, wastewater contains a mixture of dyes, organic solvents, ions, organic matter, and humic substances [42,43]. Therefore, we determined the influence of organic solvents on AzoRed2 activity. Table 3 shows that DMSO and methanol inhibited AzoRed2 activity at a concentration of 10% (v/v); however, these solvents had no inhibitory effect at 20% (v/v) concentration. Surprisingly, ethanol and isopropanol

Scheme 2. Diagram of methyl red degradation.

1044

H. Dong et al. / International Journal of Biological Macromolecules 140 (2019) 1037–1046

Table 4 Enzymatic activities determination of coupled system. NO.

Coenzyme

Containing glucose

Substrate

Relative activity (U/g)

1 2 3

NADH NAD+ NAD+

No Yes Yes

Methyl red Methyl red NAD+

13.92 ± 0.29 15.59 ± 0.13 162.58 ± 5.48

inhibited AzoRed2 activity at 10% (v/v) concentration and activated AzoRed2 activity at concentrations up to 20% (v/v). Yang et al. isolated an organic solvent-tolerant azoreductase from Shewanella oneidensis MR-1, and they also found ethanol and isopropanol with low concentrations had slight activation effects on the azoreductase activities, which was similar to our AzoRed2 [39]. Other organic solvents tested all had inhibitory effects with different degrees, especially n-butanol, isoamyl alcohol, and chloroform. Above all, AzoRed2 had good tolerances toward some organic solvents (e.g. DMSO, methanol, ethanol, and isopropanol), which might be beneficial to the degradation of azo dyes in wastewater containing organic solvents. 3.4. Biodegradation of methyl red using a whole-cell biocatalyst The reductive cleavage of the -N=N- bond of azoreductase requires a high and continuous pool of electron donors (Scheme 2) such as NADH and NADPH [20]. The additional coenzyme would highly increase the treatment price, which would limit the widespread use of azoreductases in the industry. In situ coenzyme regeneration was carried out to solve this problem. The integrated enzyme system consisted of Streptomyces sp. S27 azoreductase (AzoRed2) and B. subtilis glucose 1-dehydrogenase (BsGDH). After induction, the co-expression cells were collected and lysed, and the SDS-PAGE results are shown in Fig. 5. This result indicated that AzoRed2 and BsGDH could both be

highly expressed as soluble fractions with predicted molecular weights. The supernatant of the cell lysate from the coupled enzyme system was used to determine its azoreductase activity and GDH activity. As shown in Table 4, the relative azoreductase activity was 13.92 U/g using NADH as a coenzyme (NO.1), while the degradation activity toward methyl red was up to 15.59 U/g using NAD+ as a coenzyme (NO.2). This phenomenon that degrading activity toward methyl red was higher using NAD+ as coenzyme may be ascribed to the high BsGDH activity (162.58 U/g, NO.3) in the supernatant, where BsGDH reduced NAD+ to NADH quickly to maintain the NADH concentration at a high level. Importantly, BsGDH was not the limiting rate enzyme in the coupled enzyme system, which could fulfill the supply of NADH toward azo dye decolorization in real time. Our result shows the high expression level of both AzoRed2 and BsGDH, which may contribute to the efficient decolorization of azo dye wastewater. In Rathod's report [20], the authors constructed a coupled system containing an azoreductase azoA and a formate dehydrogenase fdh. However, the expression of azoA was much low, which led to a low level of decolorization. External addition of coenzyme NAD(P)+ has been reported to greatly increase the catalytic efficiency and improve the conversion rate [27,44]. The loading of external NAD+ was examined, and the results are shown in Fig. 6a. Adding additional NAD+ could significantly improve the conversion rate of the whole-cell biocatalyst in 90 min. Furthermore, the degradation rate improved slightly with increasing amounts of NAD+. After incubation for 90 min, the conversions were all over 98% when external NAD+ was added. However, the decolorization rate was much lower with no additional NAD+ in 90 min. In the coenzyme regeneration reaction, glucose acted as the hydrogen donor to reduce NAD+ to NADH. It was reported that the concentration of glucose can influence the activities of coupled systems [28]. Fig. 6b shows that with increasing glucose concentration from 5 to 60 mM, the reaction rate increased slightly. With a glucose

Fig. 6. Optimization of methyl red decolorization using whole-cell biocatalyst. (a) Effects of external NAD+ addition amount. (b) Effects of the glucose concentrations. (c) Effects of the loading of the whole-cell biocatalyst. (d) Substrate tolerance of the whole-cell biocatalyst.

H. Dong et al. / International Journal of Biological Macromolecules 140 (2019) 1037–1046

concentration of 5 mM, the decolorization was 97.46% in 120 min. With glucose concentrations exceeding 10 mM, the decolorization of the mixed solution was N98% (98.42% at 10 mM glucose). In the end, we chose 10 mM as the optimal glucose concentration to remove methyl red in order to reduce the treatment price. Yang et al. reported an integrated enzyme system for dye removal, however, the reaction system contained 250 mM glucose, which was far less economical than ours [28]. It has been accepted that the dosages of biocatalysts highly influenced the reaction speed. In this section, the results indicated that the decolorization increased significantly with the dosages of whole-cell biocatalysts increasing from 2.5 to 15.0 mg/mL (Fig. 6c). When the dosages were over 12.5 mg/mL, the decolorization was up to 97% in 20 min. Moreover, with a loading of over 5.0 mg/mL biocatalyst, the decolorization percentages of methyl red all reached 99% in 120 min, which emphasized that the high decolorization efficiency (N99%) could be achieved by increasing the biocatalyst loading with no external NAD+ added. Considering the economic factors, we finally used a relatively lower dosage of 5.0 mg/mL as the optimal biocatalyst loading. Substrate tolerance is an important factor in evaluating the performances of biocatalysts, and biocatalysts with high substrate tolerance are promising in the industries. As shown in Fig. 6d, when the methyl red concentrations increased from 100 to 250 μM, the decolorization efficiency decreased slightly. However, the decolorization still reached 99.11% in 120 min, even at a substrate concentration of 250 μM, meaning that no substrate inhibition occurred. In Yang's report [28], when the methyl red concentration was up to 200 μM, the activity of the integrated enzyme system decreased with the substrate inhibition. And this indicated that our biocatalysts had a higher substrate tolerance than Yang's [28]. Compared with free enzyme biocatalysts, reusability is an attractive feature for whole-cell biocatalysts because of their easy recovery. We set 98% as the decolorization baseline that we wished to achieve, i.e., the goal being that the methyl red was almost completely degraded. As shown in Fig. 7, in three batch trials, the methyl red was decolored efficiently, and the decolorization remained at a high level (99.39%, 98.50%, and 97.91%) per batchwise cycle. Rathod et al. used the azoA and fdh coupled system to degrade reactive violet 5R (RV5R), and the decolorization efficiency was only 30% in 12 h with the final RV5R concentration of 2 μM [20]. Gao et al. constructed a fed-batch reactor by coupling GtAZR and LsGDH-DS255, and under a relative low flow rate at 60 mL·h−1, the decolorization was highly efficient (99.3%) [31]. However, this reaction system contained 200 mM glucose and external addition of 1.0 mM NAD+, which increased the cost of methyl red treatment [31]. Furthermore, there are some reports of using the whole-cell biocatalysts containing azoreductases and no coenzyme regeneration system to degrade azo dyes. For example, Feng et al. [22] used the recombinant E. coli cell

Fig. 7. Batch experiments of the whole-cell biocatalyst.

1045

BL21(DE3)plysS consisting of azoreductase AzoA to degrade azo dyes. The final decolorization efficiency was around 96% with low methyl red concentration (25 μM) and external NADH added, which was much less efficient than our system [22]. From the above discussion, we finally conclude that our whole-cell biocatalyst is efficient for methyl red degradation. 4. Conclusion In this work, a novel azoreductase (AzoRed2) from Streptomyces sp. S27 was identified and expressed for dye decolorization in the industrial wastewater. Wastewater always contains a mixture of dyes, organic solvents, ions, organic matter, and humic substances that may influence the activities of biocatalysts. Notably, AzoRed2 showed excellent stability against pH changes, organic solvents, and methyl red, indicating that AzoRed2 could be stable against harsh conditions. By introducing an in situ coenzyme regeneration system, an efficient whole-cell biocatalyst containing AzoRed2 and BsGDH was constructed, achieving the goal of complete azo dye degradation (N 99%) without external NAD+ addition. The overall results suggest that AzoRed2 could be a promising candidate in the treatment of industry wastewater containing azo dyes. Acknowledgements This work was supported by the National Natural Science Foundation of China (21636003 and 21672065). We thank LetPub (www. letpub.com) for its linguistic assistance during the preparation of this manuscript. Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi. org/10.1016/j.ijbiomac.2019.08.196. References [1] B. Gupta, M. Rani, R. Salunke, R. Kumar, In vitro and in vivo studies on degradation of quinalphos in rats, J. Hazard. Mater. 213-214 (2012) 285–291. [2] T.S. Natarajan, M. Thomas, K. Natarajan, H.C. Bajaj, R.J. Tayade, Study on UV-LED/ TiO2 process for degradation of Rhodamine B dye, Chem. Eng. J. 169 (1) (2011) 126–134. [3] U. Shanker, M. Rani, V. Jassal, Degradation of hazardous organic dyes in water by nanomaterials, Environ. Chem. Lett. 15 (4) (2017) 623–642. [4] M. Rafatullah, O. Sulaiman, R. Hashim, A. Ahmad, Adsorption of methylene blue on low-cost adsorbents: a review, J. Hazard. Mater. 177 (1) (2010) 70–80. [5] F.P. Van der Zee, F.J. Cervantes, Impact and application of electron shuttles on the redox (bio)transformation of contaminants: a review, Biotechnol. Adv. 27 (3) (2009) 256–277. [6] N.Ž. Šekuljica, N.Ž. Prlainović, A.B. Stefanović, M.G. Žuža, D.Z. Čičkarić, D.Ž. Mijin, D.Z. Knežević-Jugović, Decolorization of anthraquinonic dyes from textile effluent using horseradish peroxidase: optimization and kinetic study, Sci. World J. 2015 (2015) 12. [7] A. Gürses, M. Açıkyıldız, K. Güneş, M.S. Gürses, Dyes and pigments: their structure and properties, in: A. Gürses, M. Açıkyıldız, K. Güneş, M.S. Gürses (Eds.), Dyes and Pigments, Springer International Publishing, Cham 2016, pp. 13–29. [8] N.M. Julkapli, S. Bagheri, S.B.A. Hamid, Recent advances in heterogeneous photocatalytic decolorization of synthetic dyes, Sci. World J. 2014 (2014) 25. [9] S.K. Sen, S. Raut, P. Bandyopadhyay, S. Raut, Fungal decolouration and degradation of azo dyes: a review, Fungal Biol. Rev. 30 (3) (2016) 112–133. [10] A. Ajmal, I. Majeed, R.N. Malik, H. Idriss, M.A. Nadeem, Principles and mechanisms of photocatalytic dye degradation on TiO2 based photocatalysts: a comparative overview, RSC Adv. 4 (70) (2014) 37003–37026. [11] H. Zhang, D. Chen, X. Lv, Y. Wang, H. Chang, J. Li, Energy-efficient photodegradation of azo dyes with TiO2 nanoparticles based on photoisomerization and alternate UV −visible light, Environ. Sci. Technol. 44 (3) (2010) 1107–1111. [12] F.M.D. Chequer, G.A.R.d. Oliveira, E.R.A. Ferraz, J. Carvalho, M.V.B. Zanoni and D.P.d. Oliveir, Textile dyes: Dyeing process and environmental impact, in: M. Günay (Eds.), Eco-friendly Textile Dyeing and Finishing. IntechOpen, (2013). [13] K. Golka, S. Kopps, Z.W. Myslak, Carcinogenicity of azo colorants: influence of solubility and bioavailability, Toxicol. Lett. 151 (1) (2004) 203–210. [14] K. Golka, W. Weistenhofer, P. Jedrusik, F. Geller, M. Blaszkewicz, H.M. Bolt, Nacetyltransferase 2 phenotype in painters with bladder cancer and controls, Ann. Acad. Med. Singap. 30 (5) (2001) 464–467. [15] K. Golka, A. Wiese, G. Assennato, H.M. Bolt, Occupational exposure and urological cancer, World J. Urol. 21 (6) (2004) 382–391.

1046

H. Dong et al. / International Journal of Biological Macromolecules 140 (2019) 1037–1046

[16] S.S. Vaghela, A.D. Jethva, B.B. Mehta, S.P. Dave, S. Adimurthy, G. Ramachandraiah, Laboratory studies of electrochemical treatment of industrial azo dye effluent, Environ. Sci. Technol. 39 (8) (2005) 2848–2855. [17] R.L. Singh, P.K. Singh, R.P. Singh, Enzymatic decolorization and degradation of azo dyes – a review, Int. Biodeterior. Biodegradation 104 (2015) 21–31. [18] M. Eslami, M.A. Amoozegar, S. Asad, Isolation, cloning and characterization of an azoreductase from the halophilic bacterium Halomonas elongata, Int. J. Biol. Macromol. 85 (2016) 111–116. [19] S. Mishra, A. Maiti, The efficacy of bacterial species to decolourise reactive azo, anthroquinone and triphenylmethane dyes from wastewater: a review, Environ. Sci. Pollut. R. 25 (9) (2018) 8286–8314. [20] J. Rathod, S. Dhebar, G. Archana, Efficient approach to enhance whole cell azo dye decolorization by heterologous overexpression of Enterococcus sp. L2 azoreductase (azoA) and Mycobacterium vaccae formate dehydrogenase (fdh) in different bacterial systems, Int. Biodeterior. Biodegradation 124 (2017) 91–100. [21] A. Moutaouakkil, Y. Zeroual, F.Z. Dzayri, M. Talbi, K. Lee, M. Blaghen, Purification and partial characterization of azoreductase from Enterobacter agglomerans, Arch. Biochem. Biophys. 413 (1) (2003) 139–146. [22] J. Feng, T. M. Heinze, H. Xu, C. E. Cerniglia, H. Chen, Evidence for significantly enhancing reduction of azo dyes in Escherichia coli by expressed cytoplasmic azoreductase (AzoA) of Enterococcus faecalis, Protein Pept. Lett. 17 (5) (2010) 578–584. [23] M. Nakanishi, C. Yatome, N. Ishida, Y. Kitade, Putative ACP phosphodiesterase gene (acpD) encodes an azoreductase, J. Biol. Chem. 276 (49) (2001) 46394–46399. [24] K.i. Matsumoto, Y. Mukai, D. Ogata, F. Shozui, J.M. Nduko, S. Taguchi, T. Ooi, Characterization of thermostable FMN-dependent NADH azoreductase from the moderate thermophile Geobacillus stearothermophilus, Appl. Microbiol. Biotechnol. 86 (5) (2010) 1431–1438. [25] K. Verma, G. Saha, L.M. Kundu, V.K. Dubey, Biochemical characterization of a stable azoreductase enzyme from Chromobacterium violaceum: application in industrial effluent dye degradation, Int. J. Biol. Macromol. 121 (2019) 1011–1018. [26] Q. Ye, P. Ouyang, H. Ying, A review—biosynthesis of optically pure ethyl (S)-4chloro-3-hydroxybutanoate ester: recent advances and future perspectives, Appl. Microbiol. Biotechnol. 89 (3) (2011) 513–522. [27] Y. Zhang, H. Wang, L. Chen, K. Wu, J. Xie, D. Wei, Efficient production of ethyl (R)-4chloro-3-hydroxybutanoate by a novel alcohol dehydrogenase from Lactobacillus curieae S1L19, J. Mol. Catal. B Enzym. 134 (2016) 51–60. [28] Y. Yang, B. Wei, Y. Zhao, J. Wang, Construction of an integrated enzyme system consisting azoreductase and glucose 1-dehydrogenase for dye removal, Bioresour. Technol. 130 (2013) 517–521. [29] L.L. Lu, Y.Y. Yang, H. Lin, F. Gao, Y.H. Zhao, Construction, expression and characterization of fusion enzyme containing azoreductase and glucose 1-dehydrogenase for dye removal, Int. Biodeterior. Biodegradation 87 (2014) 81–86.

[30] D. Cui, G. Li, D. Zhao, X. Gu, C. Wang, M. Zhao, Purification and characterization of an azoreductase from Escherichia coli CD-2 possessing quinone reductase activity, Process Biochem. 47 (3) (2012) 544–549. [31] F. Gao, H. Ding, L. Shao, X. Xu, Y. Zhao, Molecular characterization of a novel thermal stable reductase capable of decoloration of both azo and triphenylmethane dyes, Appl. Microbiol. Biotechnol. 99 (1) (2015) 255–267. [32] T. Mamiatis, E.F. Fritsch, J. Sambrook, J. Engel, Molecular cloning–a laboratory manual. New York: Cold Spring Harbor Laboratory. 1982, 545 S., 42 $, Acta Biotechnol. 5 (1) (1985) 104. [33] J. Feng, T. Han, M.Q. Zhang, Y. Zhou, Q.Q. Wu, Application of 2D fluorescence correlation method to investigate the dilution-induced heterogeneous distribution of the bound FMN in azoreductase, Chin. Chem. Lett. 26 (2) (2015) 210–214. [34] H. Dong, F. Secundo, C. Xue, X. Mao, Whole-cell biocatalytic synthesis of cinnamyl acetate with a novel esterase from the DNA library of Acinetobacter hemolyticus, J. Agric. Food Chem. 65 (10) (2017) 2120–2128. [35] W. Gao, K. Wu, L. Chen, H. Fan, Z. Zhao, B. Gao, H. Wang, D. Wei, A novel esterase from a marine mud metagenomic library for biocatalytic synthesis of short-chain flavor esters, Microb. Cell Factories 15 (1) (2016) 41. [36] H. Chen, J. Feng, O. Kweon, H. Xu, C.E. Cerniglia, Identification and molecular characterization of a novel flavin-free NADPH preferred azoreductase encoded by azoB in Pigmentiphaga kullae K24, BMC Biochem. 11 (1) (2010) 13–23. [37] H. Chen, Recent advances in azo dye degrading enzyme research, Curr. Protein Pept. Sci. 7 (2) (2006) 101–111. [38] A. Ryan, N. Laurieri, I. Westwood, C.J. Wang, E. Lowe, E. Sim, A novel mechanism for azoreduction, J. Mol. Biol. 400 (1) (2010) 24–37. [39] Y. Yang, L. Lu, F. Gao, Y. Zhao, Characterization of an efficient catalytic and organic solvent-tolerant azoreductase toward methyl red from Shewanella oneidensis MR1, Environ. Sci. Pollut. Res. 20 (5) (2013) 3232–3239. [40] J. Qi, M. Schlömann, D. Tischler, Biochemical characterization of an azoreductase from Rhodococcus opacus 1CP possessing methyl red degradation ability, J. Mol. Catal. B Enzym. 130 (2016) 9–17. [41] N. Adhoum, L. Monser, N. Bellakhal, J.E. Belgaied, Treatment of electroplating wastewater containing Cu2+, Zn2+ and Cr(VI) by electrocoagulation, J. Hazard. Mater. 112 (3) (2004) 207–213. [42] G.A. Epling, C. Lin, Photoassisted bleaching of dyes utilizing TiO2 and visible light, Chemosphere 46 (4) (2002) 561–570. [43] V. García, E. Pongrácz, P.S. Phillips, R.L. Keiski, From waste treatment to resource efficiency in the chemical industry: recovery of organic solvents from waters containing electrolytes by pervaporation, J. Clean. Prod. 39 (2013) 146–153. [44] L.F. Chen, Y.P. Zhang, H.Y. Fan, K. Wu, J.P. Lin, H.L. Wang, D.Z. Wei, Efficient bioreductive production of (R)-N-Boc-3-hydroxypiperidine by a carbonyl reductase, Catal. Commun. 97 (2017) 5–9.