Achromobacter denitrificans strain SP1 produces an intracellular esterase upon utilizing di(2ethylhexyl)phthalate

International Biodeterioration & Biodegradation 105 (2015) 160e167

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Achromobacter denitrificans strain SP1 produces an intracellular esterase upon utilizing di(2ethylhexyl)phthalate Selvanesan Pradeep a, b, Moolakkariyil Sarath Josh a, Erandapurathukadumana Sreedharan Hareesh a, Sunil Kumar c, Sailas Benjamin a, * a Enzyme Technology Laboratory, Biotechnology Division, Department of Botany, School of Bioscience, University of Calicut, Malappuram, 673635, Kerala, India b Environmental Technology Division, CSIR-National Institute for Interdisciplinary Science & Technology, Thiruvananthapuram, 695019, Kerala, India c Solid and Hazardous Waste Management Division, CSIR-NEERI, Nagpur, 440020, India

a r t i c l e i n f o

a b s t r a c t

Article history: Received 1 June 2015 Received in revised form 10 September 2015 Accepted 10 September 2015 Available online xxx

Production of an esterase by Achromobacter denitrificans strain SP1 e a di(2-ethylhexyl)phthalate (DEHP) degrading novel bacterium e in a modified basal salt medium supplemented with DEHP as an inducercum-additional carbon source was studied. The PlacketteBurman and BoxeBehnken designs were applied to statistically optimize the production parameters, which resulted in an increase of esterase production by 24%. For the production of the maximum (30.5 U) intracellular esterase, 10 mM DEHP and 72 h incubation at pH 8.0 were found as optimum conditions; while the predicted value was 28.8 U with a correlation coefficient of 0.932; which signifies the fitness of the model. The optimum activity of the 2.5 folds purified esterase was 89.5 U with 20 mM para-nitrophenyl acetate as substrate (50
C, pH 8.0 and for 30 min), and various metal ions were found to retarde the esterase activity. The approximate MW of partially purified esterase was 53 kDa, and the activity of esterase was also confirmed by native-PAGE. The Km and Vmax values of esterase were 1.308 mM and 62.52 mmol min1 mg1, respectively. Briefly, this was the first report on an enzyme from the DEHP degrading A. denitrificans SP1, which in comparison with esterase from other phthalate degrading bacteria and fungi showed better Km and Vmax. © 2015 Elsevier Ltd. All rights reserved.

Keywords: Achromobacter denitrificans strain SP1 Esterase Response surface methodology Native-PAGE SDS-PAGE

1. Introduction Owing to a wide array of applications, modern life of humans greatly depends on petroplastics; their rampant use and improper disposal often create serious threat to the environment and biota. The fat soluble phthalates (or phthalic acid esters) encompass a major class of plasticizers used to blend with the rigid polyvinylchloride (PVC) to transform it into flexible products of consumer interest (Hauser and Calafat, 2005). Di(2-ethylhexyl)phthalate (DEHP) is the prime phthalate produced globally, mainly to plasticize the PVC. PVC biomedical devices being used in blood transfusion, dialysis, extracorporeal membrane oxygenation, etc. are also composed of phthalates, especially DEHP (Hauser and Calafat, 2005; Heudorf et al., 2007). Due to the lack of covalent linkage with PVC, DEHP blended in it easily leaches out of the polymer mesh into the surrounding environments (Sarath Josh et al., 2012).

* Corresponding author. E-mail address: [email protected] (S. Benjamin). http://dx.doi.org/10.1016/j.ibiod.2015.09.007 0964-8305/© 2015 Elsevier Ltd. All rights reserved.

A portion of DEHP leachate from these PVC plastics including medical devices subsequently reaches into the biota including humans, thereby causing various dysfunctions like disruption of the endocrine system, cancer, nervous disorders, anomalies in sexual differentiation, etc. (Koch et al., 2006; Latini et al., 2010), and that the main routes of phthalate exposure to humans maybe ingestion via food, drinks, personal care products, inhalation, house dust, wall covering materials, etc. (Koch et al., 2006; Heudorf et al., 2007). Biodegradation through aerobic or anaerobic (or both) microbial intervention would significantly influence the environmental fate of phthalates. Many bacterial strains (Chatterjee and Dutta, 2008; Pradeep et al., 2014), fungi (Luo et al., 2009; Pradeep et al., 2013), and a few yeasts (Gartshore et al., 2003) and algae (Yan and Pan, 2004) emerged as the major degraders of phthalates. Esterases (EC 3.1.1.60) play a crucial role in the degradation process of xenobiotic compounds like phthalates, which hydrolyze them into phthalic acid and free alcohols, i.e., the initial rate-limiting step in hydrolysis of phthalates (Benjamin et al., 2015). Due to their

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acceptability in the food, pharmaceutical, textile, paper and leather industries, screening and identification of novel microorganisms producing esterases are of great importance. Characterization of esterases produced by microbes using phthalate as both inducer and substrate is addressed hardly (Luo et al., 2012). Very recently, our group demonstrated a pharmaceutically active 25C prodigiosin produced by Achromobacter denitrificans strain SP1 (a new strain) upon utilizing DEHP as the sole source of carbon (Pradeep et al., 2014), and we partly deduced the DEHP biodegradation pathway (Pradeep et al., 2015). Based upon this background; as a part of the ongoing studies on the A. denitrificans strain SP1, the present study addresses: (a) production of esterase by A. denitrificans SP1 in modified basal salt medium (BSM) containing DEHP as both inducer-cum-additional carbon source; (b) statistical optimization of growth parameters for the enhanced production of esterase; and (c) partial purification and characterization of the esterase produced. 2. Materials and methods 2.1. Esterase production medium The BSM (Pradeep and Benjamin, 2012) supplemented with DEHP was used for the production of esterase by the addition of 1 /4th the strength of nutrient broth. The composition of modified BSM used in the present study was (g L1): 2.25 NaCl, 1.0 K2HPO4,0.5 NH4Cl, 0.4 MgSO4, 1.25 peptone and 0.75 beef extract (pH 7.2). The 12 h old inoculum of A. denitrificans strain SP1 contained ~3  108 cfu (in100 mL BSM); which was incubated in a temperature-controlled shaker (Scigenics Biotech, India), and set at different conditions as described in the succeeding sections. 2.2. Cell-free extract for the estimation of esterase production Cells of A. denitrificans SP1 grown in the afore-described medium were harvested by centrifugation (8000g for 10 min at 4
C) after incubation for required time, as described in the following experimental designs. The pellet was washed twice with 50 mM potassium phosphate buffer (pH 7.5), and re-suspended in the same buffer (1 g mL1), which was disrupted by sonication (output wattage 15 for 5 min) at 4
C. The resulting cell homogenate was centrifuged (10,000g for 25 min at 4
C), and the clear supernatant so obtained was used as source of crude esterase. Production of extracellular esterase was found to be negligible (0.13 U mL1 before optimization and 0.19 U mL1 after optimization); hence only intracellular esterase was focused in this study. Furthermore, neither extracellular nor intracellular esterase activity was detected in the absence of DEHP in the growth medium. 2.3. Statistical optimization for esterase production 2.3.1. PlacketteBurman experimental design Five independent variables were selected for the initial screening studies; i.e., temperature, pH, agitation, concentration of DEHP supplied in the medium, and incubation time. Twenty different combinations were generated to estimate the combined effects of these parameters on esterase production (Table 1). Minitab version 14 was used to generate the experimental design with high (þ1) and low (1) levels for each variable selected. 2.4. BoxeBehnken model and response surface methodology Based on the results of PlacketteBurman experiment, three parameters (concentration of DEHP, incubation time and pH of the medium) were found significant for the production of esterase by

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A. denitrificans SP1. Employing these three parameters, different combinations of experiments were designed (Table 2). Each variable was analyzed at low (1), medium (0) and high (þ1) levels, and the results were analyzed by applying second order polynomial equation. Validation experiments for the quadratic model were conducted as predicted by the point prediction software Minitab 14 (Supplementary Table 1). The difference between the estimated and predicted values of esterase activity was compared.

2.5. Esterase assay Modified method of Lee et al. (1999) was employed for the assay of esterase activity using para-nitrophenyl acetate (p-NPA) as substrate. Briefly, the reaction mixture contained 1710 mL of 50 mM potassium phosphate buffer (pH 7.5), 72 mL absolute alcohol, 600 mL esterase solution and 18 mL substrate (10 mM p-NPA in acetonitrile). Control samples were also set without esterase solution. The reaction mixture was incubated at 37
C for 15 min, subsequently the absorbance (optical density, OD) of the samples was measured at 405 nm. One unit of esterase activity (U) corresponds to the esterase present in 1 g wet cell pellet (in the case of intracellular esterase) or 1 ml culture broth (in the case of extracellular esterase) required for the liberation of 1 mmol of para-nitrophenol per min from p-NPA under the assay conditions described elsewhere. Esterase activity was calculated using the formula,

Esterase activity ðUÞ ¼

DE  Vf ; Dt  ε  Vs  d

where DE (absorbance at 405 nm), Vf (final volume), Vs (volume in mL of esterase used), Dt (time of hydrolysis), ε (Extinction coefficient, 0.017), d (diameter of cuvette, 1 cm for standard cuvette).

2.6. Partial purification and characterization of esterase A. denitrificans SP1 was cultivated under the statistically optimized conditions (according to the BoxeBehnken design), and the cells harvested were disrupted by sonication for obtaining the crude esterase extract in 50 mM potassium phosphate buffer (pH 7.5), which was further subjected to step-wise ammonium sulphate fractionation until it reached 80% saturation (i.e., 0e20, 20e40, 40e60 and 60e80%). The precipitated protein in each step was pelleted by centrifugation (2400g for 10 min at 4
C). The pellet was re-suspended in a minimum volume of 50 mM potassium phosphate buffer (pH 7.5); dialyzed against the same buffer for 24 h at 4
C with two buffer changes. The crude enzyme sample obtained after dialysis was subjected to Vivaspin column purification (Vivaspin 6, Sweden); packed with polyethersulfone; followed by sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDSPAGE). The Vivaspin column would demarcate the sample into two fractions, i.e., upper fraction with MW higher than 45 kDa and the lower fraction with MW below 45 kDa. Both fractions were tested for esterase activity.

2.7. SDSePAGE The partially purified enzyme was subjected to SDSePAGE using PAGE (12% resolving gel), and stained with Coomassie Brilliant Blue R-250 (CBB) to determine the approximate molecular weight (MW) of the purified protein. SDS-PAGE was performed on a vertical mini gel (8  7 cm) slab with notched glass plate system. Gels of 1.5 mm thickness were prepared for the entire study. Running voltage for stacking was 70 and for resolving, it was 100.

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Table 1 Results of PlacketteBurman experimental design for the production of esterase by A. denitrificans strain SP1.


Run order

Temp ( C)

pH

Agitation (rpm)

DEHP (mM)

Time (h)

Esterase (U)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

32 32 28 32 28 32 28 32 28 32 28 28 32 32 28 32 28 32 28 28

7.5 7.5 7.5 5.5 7.5 5.5 5.5 7.5 5.5 5.5 5.5 7.5 7.5 5.5 5.5 5.5 7.5 7.5 5.5 7.5

200 200 200 200 100 200 100 100 200 100 100 200 100 200 200 100 200 100 100 100

5.0 15 5.0 15 15 15 5.0 5.0 5.0 15 15 15 5.0 5.0 15 5.0 5.0 15 5.0 15

30 30 50 30 30 50 30 30 50 50 30 50 50 50 30 30 30 50 50 50

19.5 12.0 23.2 11.0 13.85 16.1 12.2 21.5 20.4 15.8 11.49 16.83 30.5 17.13 12.51 15.3 18.5 18.0 19.6 20.1

Table 2 Results of BoxeBehnken design for the production of esterase by A. denitrificans strain SP1. Run order

DEHP (mM)

Time (h)

pH

Observed activity (U)

Predicted activity (U)

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15

10 10 15 20 15 20 20 10 15 20 15 15 15 10 15

72 60 72 72 48 48 60 48 72 60 48 60 60 60 60

6.75 8.0 8.0 6.75 8.0 6.75 8.0 6.75 5.5 5.5 5.5 6.75 6.75 5.5 6.75

26.2 24.5 22 11 15.2 7.8 10 17 13.8 8.93 14.7 20 21.6 18.5 17.6

25.3 24.7 22.6 10.2 14.2 8.6 10.2 17.7 14.8 8.6 14.0 19.7 19.7 18.3 19.7

2.8. Native-PAGE and activity staining Polyacrylamide gel system without SDS was used for nativePAGE, in which 5% stacking and 10% resolving gels were used (Niazi et al., 2001). Broad range protein MW marker (GeNei, India) was used to judge the approximate MW of esterase on the gel. After performing the native PAGE at 4
C and 100 V (for both stacking and resolving), the gel was cut into two halves, in such a way to retain the MW marker and the protein sample in one half (used for CBB staining), and the other half with protein sample only (for esterase activity staining), thus the migration of protein bands could be compared easily on both halves by differential staining. The esterase activity staining was performed by incubating the gel (impregnated with native esterase) in 100 mL of 50 mM TriseHCl buffer (pH 7.0) containing 0.5% (w/v) Fast Blue BB salt and 5.38 mM a-naphthyl acetate (substrate for esterase) for 20 min at 37
C. The esterase impregnated into the gel hydrolyzes the a-naphthyl acetate to a-naphthol, which would further react with the Fast Blue BB salt to form a complex, thereby giving intense brownish band at the sites of esterase activity (Niazi et al., 2001).

studies. In order to identify the characteristics of esterase, i.e., effects of pH, temperature, substrate (p-NPA) concentration, and different metal ions (Kþ, Naþ, Ca2þ, Mn2þ, and Zn2þ) on esterase activity were studied. 2.10. Effect of pH on esterase activity The optimum pH for esterase activity was determined by measuring the enzymatic activities in 50 mM phosphate buffer (pH 6.0, 6.5, 7.0 and 7.5), and 50 mM TriseHCl buffer (pH 8.0, 8.5 and 9.0) at 37
C for 15 min incubation. 2.11. Effect of temperature on enzyme activity The optimum temperature for esterase activity was determined by measuring the enzymatic activities upon incubation in 50 mM TriseHCl buffer (pH 8.0) at different temperatures, i.e., 25, 30, 35, 37, 40, 45, 50, 55, 60 and 65
C, with an incubation time for 15 min. 2.12. Effect of different metal salts on enzyme activity

2.9. Characterization of esterase The Vivaspin column fraction was used for the characterization

Effects of various metal ions on esterase activity was determined by incubating the reaction mixture with different metal salts, i.e., Kþ, Naþ, Mn2þ, Ca2þ, and Zn2þ to a final concentration of 0.1, 0.25,

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0.5 and 1.0 mM at 50
C and pH 8.0 for 15 min.

3.3. Enzyme characteristics

2.13. Calculation of Km and Vmax

3.3.1. Effect of pH on enzyme activity The upper molecular weight fraction (>45 kDa) from the Vivaspin column separation was used for the characterization studies. The optimal esterase activity (70 U) was obtained at pH 8.0, and the lowest (15.6 U) was at pH 6.0. Esterase activity was considerably decreased at low/acidic pH (Fig. 3A).

The enzyme solution was treated with p-NPA at a concentration of 1.0, 5.0, 10, 15 and 20 mM. The reaction mixture was incubated for 5e45 min with 5 min intervals; i.e., 5, 10, 15, 20, 25, 30, 35, 40, 45 min at 50
C and pH 8.0. The Km and Vmax values were calculated for esterase using the software, Hyper 32.

3.4. Effect of temperature on enzyme activity 3. Results 3.1. Statistical optimization of esterase production The results of PlacketteBurman experimental design are given in Table 1. The maximum esterase production (23.2 U) was attained at 28
C, pH 7.5, rpm 200, 5 mM DEHP and 50 h incubation, while the minimum production (11 U) was observed at 32
C, pH 5.5, rpm 200, 15 mM DEHP and 30 h incubation (Table 1). Based on the results of PlacketteBurman experimental design, a pareto chart was constructed (Supplementary Fig. 1). The effects of the standardized parameters (temperature, pH, agitation, incubation time and concentration of DEHP) on esterase production were estimated at 5% level of significance. This analysis showed that the esterase production was highly influenced by three parameters, i.e., the concentration of DEHP, incubation time and pH. Subsequently, these three parameters were considered for BoxeBenhnken experimental design by RSM to identify the optimized conditions for the maximum production of esterase by A. denitrificans SP1 (Fig. 1). Regression coefficient was estimated for the production of esterase (esterase versus concentration of DEHP, incubation time or pH). The regression model for the production of esterase was highly significant (p < 0.05) with an acceptable value of determination coefficient (R2 ¼ 96.95%). The regression equation obtained for esterase production was: Y ¼ 87.5815 þ 4.7X1þ1.0X2þ11.2 X30.1X210.01X221.0X23 0.02X1X20.2X1X3þ0.1X2X3; where X1, X2, X3 are DEHP, time and pH, respectively. Three parameters (concentration of DEHP, incubation time and pH) selected from the validation experiments falling within the range determined from BoxeBehnken were assessed in detail for the aptness of the model (Table 2). The correlation coefficient of the result was 0.932, which was in good agreement with the predicted and experimental values. The optimum esterase production was 30.5 U (i.e., 24% increase after optimization), when the medium was supplemented with 10 mM DEHP (pH 8.0, 72 h) (Supplementary Table 1). 3.2. Partial purification of esterase by (NH4)2SO4 fractionation Of various (NH4)2SO4 fractions, fraction 40e60% showed the maximum esterase activity, which represented a purification of 1.4 fold with 66.6% yield (Table 3). The (NH4)2SO4 fraction of protein at 40e60% saturation was subjected to Vivaspin column purification, which resulted in a 2.5 folds purified fraction with 27.9% yield (Table 3). This partially purified esterase showed that its approximate MW was 53 kDa, as revealed by the profiles on SDS-PAGE (Fig. 2A), and the esterase activity was also confirmed at nonreducing/native state (Fig. 2B). On native gel, the esterase active band was clear upon both CBB and Fast Blue BB salt staining techniques (Fig. 2B), this was done just to double check and compare the presence of active esterase, and not for judging the MW. In fact, esterase fraction showed more migration on the gel close to the 43 kDa marker protein, as evidenced from the migrated active band (Fig. 2B).

Results indicated that the temperature optimum for esterase was 50
C with an activity of 76 U (Fig. 3B), with comparable activities at 40
C and 45
C. However, at 65
C (7.1 U) and above, the esterase activity declined sharply. 3.5. Effect of different metal salts on enzyme activity at different concentrations All the metal ions (Naþ, Kþ, Ca2þ, Mn2þ or Zn2þ) tested were individually added to the reaction mixture to a final concentration of 0.1, 0.25, 0.5 and 1.0 mM and incubated at 50
C and pH 8.0. The results suggested that metal ions had no significant role in enhancing enzyme activity; however, the maximum esterase activity (70.5 U) was observed at 0.25 mM concentration of Mn2þ (Fig. 3C), which was less than the activity obtained without metal ion. It indicates that metal ion has no role in enhancing activity of esterase. 3.6. Enzyme kinetics Various concentrations (1.0, 5.0, 10, 15 and 20 mM) of the substrate (p-NPA) were used for the kinetic studies. The color reaction was found to increase upon the addition of higher concentrations of substrate. The reaction became steady at 30 min of incubation. The absorbance was measured at 405 nm (at 0, 5.0, 10, 15, 20, 25, 30, 35, 40 and 45 min) at 50
C and the esterase activity was calculated. The maximum esterase activity was 89.5 U with 20 mM p-NPA at 30 min incubation. At this state, the Km and Vmax values were found to be 1.308 mM and 62.52 mmol min1 mg1, respectively (Fig. 4). 3.7. The fold increase of activity The fold increase in esterase activity was compared with the initial activity (of crude esterase, i.e., 30.5 U) for all the factors tested (i.e., effects of pH, temperature, different metal salts and substrate concentration) (Fig. 5). The initial activity (at standard reaction conditions: 10 mM p-NPA, 7.5 pH, 37
C, 15 min) was fixed as 1. It was evident that 2.36 folds change/increase in esterase activity was observed at pH 8 (at 10 mM p-NPA, 37
C, 15 min); which was increased to 2.49 folds at 50
C (10 mM p-NPA, pH 8, 15 min); while 2.93 folds increase in esterase activity was observed at increased substrate concentration, i.e., 20 mM mM p-NPA (at pH 8.0, 50
C, 30 min). 4. Discussion A. denitrificans SP1 (MTCC No. 5710) is a new DEHP degrading bacterium; which was discovered by our group from heavily plastics contaminated sewage sludge (Pradeep et al., 2014, 2015). It utilized the hazardous DEHP supplemented in the BSM (both in situ, blended in PVC blood bag, and ex situ in liquid form) as the sole source of carbon and energy, and produced prodigiosin 25-C as a by-product (Pradeep et al., 2014). Moreover, very recently, we

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Fig. 1. Three dimensional response surface and contour plots showing the interactive effects of various parameters on the production of esterase. (A). Effects of incubation time (h) and DEHP concentration (mM); (B). Effects of pH and concentration of DEHP (mM); and (c). Effects of pH and incubation time (h).

Table 3 Summary of partial purification of esterase from A. denitrificans SP1. Purification

Total protein (mg g1 cell pellet)

Total activity (U)

Specific activity (U mg1 protein)

Yield (%)

Fold purification

Crude extract 40e60% (NH4)2SO4 fraction Spin column fraction

6.25 3.0 0.69

30.5 20.3 8.5

4.9 6.8 12.3

100 66.6 27.9

1.0 1.4 2.5

demonstrated the degradation pathway of DEHP by A. denitrificans SP1 (Pradeep et al., 2015), in which we could not detect the phthalic acid as a metabolite and protocatechuate dioxygenases responsible for the degradation of phthalic acid. Therefore, esterase activity was

explored in detail as reported herein; this is the first report on an enzyme produced by A. denitrificans SP1. The present study shows that DEHP acted as an inducer for the production of esterase. Density of cells is significantly less in BSM

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Fig. 2. A. SDS-PAGE (reducing) profile showing esterase active band with approximate MW of 53 kDa. Lane 1 shows the profile of standard protein MW marker. Lane 2 shows crude protein harvested after 72 h incubation supplemented with 10 mM DEHP in BSMþ 1/4 nutrient broth medium. Lane 3 shows the profile of 40e60% (NH4)2SO4 fraction, and lane 4 represents the profile of Vivaspin column fraction with MW cut-off >45 kDa. B. Native-PAGE profile (non-reducing) showing esterase activity on the same gel; wherein one half of the gel with MW markers (lane 1) and Vivaspin column fraction (lane 2) was stained with Coomassie Brilliant Blue (left half). Only Vivaspin column fraction was loaded on the other half (right half) of the gel (lane 3), which shows single characteristic band of esterase after the gel incubated in 50 mM TriseHCl buffer (pH 7.0) containing 0.5% (w/v) Fast Blue BB salt and 5.38 mM a-naphthyl acetate. The esterase would hydrolyze a-naphthyl acetate to a-naphthol, which further complexes with the Fast Blue BB salt to form brownish intense bands on the sites of activity. It also showed that the migration pattern of esterase was faster under non-reducing condition. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

supplemented with DEHP, therefore 1/4th the concentration of the ingredients in the nutrient broth was supplemented in the growth medium for increasing the cell density with the retention of esterase activity. Because, no esterase activity was noticed in the absence of DEHP in the medium; and on the contrary, the bacterium showed luxuriant growth in the full strength nutrient broth with no production of esterase. Intracellular bacterial esterases from di-methyl phthalate degrading bacterium Sphingomonas yanoikuyae DOS01 (Gu et al., 2009), and a di-methyl terephthalate degrading fungus Fusarium sp. DMT-5-3 were also reported (Luo et al., 2012). The initial step in the degradation pathway of phthalate esters in bacteria seems to be mediated by de-esterification reaction (Pradeep et al., 2015; Benjamin et al., 2015); for instance, Pseudomonas pseudoalcaligenes hydrolyzed di-n-butyl phthalate to mono-n-butylphthalate and phthalic acid (Benckiser and Ottow, 1982); Nocardia erythropolis hydrolyzed DEHP to its monoester and phthalic acid (Kurane et al., 1984), and an esterase from Bacillus sp. catalyzed the initial degradation of di-methyl phthalate (Niazi et al., 2001). Like esterase, this is the first report describing the application of RSM for esterase production from a phthalate degrading bacterium. As illustrated in the contour and three dimensional response surface plots, the optimum conditions such as concentration of DEHP, initial pH, incubation time were confined within the design boundary, thus well fit to the model. The validation data were statistically analyzed so as to correlate the observed and predicted values. The experimental results showed optimum esterase production of 30.5 U, when the medium was supplemented with 10 mM DEHP, whereas the predicted value was 28.8 U. The correlation coefficient value of 0.932 suggests that the experimental values were highly compatible with those of the predicted, a proof for the precision of the model. Statistically optimized conditions

were applied for the production of esterase from A. denitrificans SP1, and partially purified to 2.5 folds with 25% yield. Maruyama et al. (2005) described an esterase from Micrococcus sp.YGJ1 with 1.17 fold purification and 73.9% yield, which hydrolyzed mono-alkyl phthalates. From the SDS-PAGE profile, the approximate MW of esterase from A. denitrificans SP1 was 53 kDa. It seems that the MW of phthalate esterases produced by bacteria varies from 15 to 67 kDa (Table 4). The p-NPA is the substrate commonly used for assaying the activity of esterase (Prasad and Suresh, 2012). In the present study, the maximum esterase activity was noticed at pH 8.0, and the lowest activity was at pH 6.0, which indicated that the alkaline condition favors esterase activity. Likewise, the esterase produced by dimethyl phthalate degrading Ochrobactrum anthoropi (Xu et al., 2006), and a fungal esterase from a Fusarium sp. degrading dimethyl terephthalate (Luo et al., 2012) showed pH and temperature optima as pH 8.0 and 50
C, respectively. Present study also showed inhibitory or no effect of metal ions; similarly, presence of Ca2þ and Zn2þ in the reaction mixture inhibited the activity of esterase obtained from O. anthoropi by 75% and 47%, respectively (Xu et al., 2006). Very recently, using p-NPA as the substrate, Luo et al. (2012) showed that the activity of esterase produced by a Fusarium sp. was inhibited by Cr3þ, Cu2þ, Hg2þ, Zn2þ, Ni2þ and Cd2þ; whereas Mn2þ, Mg2þ, Ca2þ, Co2þ, Liþ, Kþ and Naþ showed almost no effect on the catalytic activity of the enzyme. From this, it seems that esterase does not require a metal ion as activator. The Km (1.308 mM) and Vmax (62.52 mmol min1 mg1) values were determined for the esterase produced by A. denitrificans strain SP1. The Km value represents the dissociation constant (affinity for substrate) of the enzymeesubstrate complex, i.e., low values of Km indicate that the enzymeesubstrate complex is held tightly and dissociates rarely before the substrate is converted to product,

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Fig. 3. A. Effect of pH on the esterase activity. Maximum esterase activity (70 U) was noticed at pH 8.0, while the lowest activity (15.6 U) was at (pH 6.0). B. Effect of temperature on esterase activity; 50
C was optimum for the maximum (75.2 U) esterase activity. C. Effect of different metal ions on enzyme activity. The maximum activity of esterase (70.5 U) was obtained with 0.25 mM of Mn2þ at 50
C and pH 8.0; while the influence of other metal ions on the activity of esterase at optimum pH and temperature was insignificant or lesser.

Fig. 4. MichaeliseMenten kinetics showing the effect of substrate concentration (pNPA) on the activity of esterase.

which in turn indicates the efficiency of the enzyme. High Vmax indicates the higher efficiency of the enzyme, i.e., the number of substrate molecules converted into product per unit time when the enzyme is fully saturated with the substrate. Various studies demonstrated that the phthalates or their degradation products (corresponding monoesters) induced the production of esterase; for instance, the dimethyl phthalte induced the production of esterase in Sphingomonas yanoikuyae DOS01, which showed a Km of 2.225 mM and Vmax of 22.691 mmol min1; mono-methyl phthalate also induced the production of esterase by the same organism, which showed a Km of 1.973 mM and Vmax of 10.772 mmol min1 (Gu et al., 2009). Mono-propyl and mono-butyl, mono-pentyl phthalates induced esterases in Micrococcus sp.YGJ1 with the Km of

Fig. 5. Fold activity in comparison to the crude esterase fraction. Compared to the initial activity of crude esterase, 1.8 folds increase was observed at pH 8.0; 2 folds increase was observed at temperature 50
C; metal ions did not influence the activity significantly, while esterase activity increased to 2.4 folds in the presence of and 20 mM p-NPA. The reaction condition of each stage is given on the X e axis.

5.88, 1.67 and 1.39 mM, respectively; and the corresponding Vmax values were: 7.81, 40.1 and 15.6 mmol min1 mg1, respectively (Maruyama et al., 2005). In comparison to these reports, it is evident that the esterase reported from A. denitrificans SP1 is much

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Table 4 Comparison of the molecular weight of phthalate esterases produced by certain bacteria. Microorganism

Type of phthalate

MW

Reference

Achromobacter denitrificans SP1 Gordonia sp. P8219 Micrococcus sp. YGJ1 Micrococcus sp. YGJ1 Ochrobactrum anthoropi 6-2b Pseudomonas sp.054 Rhodococcus erythropolis

DEHP DEHP DAPs MAPs DETP DMTP DAPs

53 31 56 60 40 67 15

Present study Nishioka et al. (2006) Akita et al. (2001) Maruyama et al. (2005) Xu et al. (2006) Tserovska et al. (2006) Kurane (1997)

efficient in activity. 5. Conclusions Statistical optimization of the production parameters resulted in an increase of esterase production by 24%. At natural environments (water or soil), this statistical optimization may not have direct impact; however, this information would be useful to judge the tolerance level of A. denitrificans SP1 once it is deployed in the DEHP (likely other phthalates too) contaminated environment. In comparison to other bacterial esterases induced by various phthalates, the esterase produced by A. denitrificans SP1 is highly promising in an industrial perspective, especially in lipid industry. Since this is the first report on an enzyme from A. denitrificans SP1, more studies are required on this esterase including the characterization of its isoforms which may be expressed upon induction by different phthalates or their metabolites. Acknowledgments The authors would like to thank the Ministry of Environment and Forests, government of India for a research Grant no. 19/62/ 2005-RE and Department of Biotechnology, Government of India for a research Grant no. BT/PR7521/BCE/8/1026/2013. Appendix A. Supplementary data Supplementary data related to this article can be found at http:// dx.doi.org/10.1016/j.ibiod.2015.09.007. References Akita, K., Naitou, C., Maruyama, K., 2001. Purification and characterization of an esterase from Micrococcus sp. YGJ1 hydrolyzing phthalate esters. Biosci. Biotechnol. Biochem. 65, 1680e1683. Benckiser, G., Ottow, J., 1982. Metabolism of the plasticizer di-n-butylphthalate by Pseudomonas pseudoalcaligenes under anaerobic conditions, with nitrate as the only electron acceptor. Appl. Environ. Microbiol. 44, 576e578. Benjamin, S., Pradeep, S., Sarath Josh, M.K., Kumar, S., Masai, E., 2015. A monograph on the remediation of hazardous phthalates. J. Hazard. Mater. 298, 58e72. Chatterjee, S., Dutta, T.K., 2008. Complete degradation of butyl benzyl phthalate by a defined bacterial consortium: role of individual isolates in the assimilation pathway. Chemosphere 70, 933e941. Gartshore, J., Cooper, D.G., Nicell, J.A., 2003. Biodegradation of plasticizers by Rhodotorula rubra. Environ. Toxicol. Chem. 22, 1244e1251. Gu, J.G., Han, B., Duan, S., Zhao, Z., Wang, Y., 2009. Degradation of the endocrinedisrupting dimethyl phthalate carboxylic ester by Sphingomonas yanoikuyae DOS01 isolated from the South China Sea and the biochemical pathway. Int. Biodeterior. Biodegr. 63, 450e455. Hauser, R., Calafat, A., 2005. Phthalates and human health. Occup. Environ. Med. 62,

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