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Characterization of a novel (R)-mandelate dehydrogenase from Pseudomonas putida NUST506
Journal of Molecular Catalysis B: Enzymatic 120 (2015) 23–27
Contents lists available at ScienceDirect
Journal of Molecular Catalysis B: Enzymatic journal homepage: www.elsevier.com/locate/molcatb
Characterization of a novel (R)-mandelate dehydrogenase from Pseudomonas putida NUST506 Jizhong Wang, Jing Feng, Weile Li, Chengli Yang, Xing Chen, Bingxin Bao, Junfang Yang, Peng Wang, Dali Li ∗ , Ruofu Shi Department of Bioengineering, Nanjing University of Science & Technology, Nanjing 210094, Jiangsu, People’s Republic of China
a r t i c l e
i n f o
Article history: Received 23 January 2015 Received in revised form 29 April 2015 Accepted 30 April 2015 Available online 11 May 2015 Keywords: Characterization (R)-Mandelate dehydrogenase Biocatalysis Pseudomonas putida Purification
a b s t r a c t (R)-Mandelate dehydrogenase (RMDH) has the potential to produce chiral mandelic acid. The present work reports isolation and identification of a Pseudomonas putida NUST506 contained RMDH. The strain was rod shaped with polar flagella, approximately 0.6–0.9 m wide and 1.7–2.3 m long. The RMDH was purified from the strain. The molecular weight of the enzyme was calculated to be 61 kDa. The optimal conditions for RMDH were pH 8.5 and 30 ◦ C. At the optimum condition, the Km and kcat of the RMDH were 2.0 × 10−2 mM and 0.9 s−1 for (R)-mandelic acid, 1.8 × 10−2 mM and 0.9 s−1 for NAD+ , as well as 1.5 × 10−2 mM and 0.3 s−1 for NADP+ , respectively. The enzyme activity was increased by K+ and dithiothreitol (DTT), but obviously inhibited by Zn2+ , Hg2+ , sodium dodecyl sulfate (SDS) and ethylenediamine tetra-acetic acid (EDTA). © 2015 Published by Elsevier B.V.
1. Introduction
2. Materials and methods
During the past few years, considerable attention has been focused on the field of chiral mandelic acid, because of its important role as building blocks widely used in the synthesis of chiral pharmaceutical and biochemical materials [1–3]. Many methods have been reported for the production of chiral mandelic acid. The chiral mandelic acid can be obtained from racemic cyanohydrins by enzyme-catalyzed enantioselective degradation [4,5], from methyl mandelate by lipase-catalyzed hydrolysis [6], or through selectively consuming (S)- or (R)-mandelic acid from the racemic mandelic acid by biotransformation and leaving (R)- or (S)-mandelic acid in reaction system [7–9]. The microbial cells enantioselective degradation of (R)-mandelic acid have been reported with Alcaligenes bronchisepticus [10], Pseudomonas polycolor [11], Gibberella fujikuroi [12], and Pseudomonas putida [8,9,13]. However, to the best of our knowledge, little is known about the characterization of RMDH. In this study, we isolated a strain P. putida NUST506 containing NAD+ -dependent RMDH and presented the purification and characterization the enzyme from the strain.
2.1. Chemicals
∗ Corresponding author. Tel.: +86 025 84315512; fax: +86 025 84315512. E-mail address: [email protected] (D. Li). http://dx.doi.org/10.1016/j.molcatb.2015.04.017 1381-1177/© 2015 Published by Elsevier B.V.
The chemicals used were of highest grade and purchased from Sinopharm Chemical Reagent Corporation (Nanjing, China). TOYOPEARL Butyl-650M was obtained from Tosoh Corporation (Osaka, Japan). The blue plus protein marker was purchased from Beijing TransGen Biotech Co. Ltd (Beijing, China). 2.2. Microorganism The microorganism used in the study was isolated from soil around the schoolyard of Nanjing University of Science and Technology (Nanjing, China). The isolates have been screened on agar plate containing (w/v) 0.4% mandelic acid, 0.4% peptone, 0.1% KH2PO4 , 0.06% MgSO4 , 0.02% FeSO4 , 0.02% MnSO4 , 0.04% CaC12 , 2% agar, and the pH was adjusted to 7.0 using 1 M NaOH. The samples were cultivated on plates and incubated at 30 ◦ C for 48 h. The bacterial colonies observed on the plates were considered as the microorganism we wanted and were used for further studies. The strain was identified as P. putida by 16S rDNA sequence. The sequence of the 16S rDNA was amplified using a polymerase chain reaction (PCR) with the forward primer (5 -AGAGTTTGATCCTGGCTCAG-3 ) and the reverse primer (5 TACGGCTACCTTGTTACGACTT-3 ). The PCR conditions used were an initial denaturation at 95 ◦ C for 3 min, followed by 35 cycles
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J. Wang et al. / Journal of Molecular Catalysis B: Enzymatic 120 (2015) 23–27
of denaturation at 94 ◦ C for 45 s, annealing at 55 ◦ C for 30 s, and extension at 72 ◦ C for 90 s, and a final extension at 72 ◦ C for 5 min. The DNA nucleotide sequence was determined by GenScript Corporation (Nanjing, China). The sequence obtained was analyzed for similarities to other known sequences found in the GenBank database using BLAST program of the NCBI database. The cells were cultivated on LB agar plates with 0.4% mandelic acid addition at 30 ◦ C for 12 h. Then the cells were collected and washed in 10 mM potassium phosphate buffer (pH 7.4). According to the similar method described previously [14], the cell morphology was examined with a transmission electron microscope (TEM, H-7650; Hitachi, Tokyo, Japan). 2.3. Partial optimization of culture conditions To determine the effect of culture time on the cell growth and on the RMDH product yield, two experiments were performed. One is recording the cell growth curve by detecting the changes of biomass concentration in fermentation broth, the other is analyzing the enzyme product yield by assaying total enzyme activity of cultivation samples. The stock culture was inoculated into 1 L Erlenmeyer flask containing 250 mL of medium (w/v, 1.0% NaCl, 1.0% peptone, 0.5% yeast extract, 0.4% mandelic acid, pH 7.0). After inoculation, the culture was incubated with shaking cultivation at 30 ◦ C and 120 rpm for up to 22 h. Two milliliters of medium were sampled at 1 h intervals and analyzed for biomass concentration by measuring the optical density value at a wavelength of 600 nm (OD600 ) using a spectrophotometer (U-1800; Hitachi, Tokyo, Japan). Fifty milliliters of fermentation broth were sampled at 2 h intervals from the fourteenth hour to the twenty-second hour of the culture broth, and rapidly cooled in an ice bath. Then the cells were collected by refrigerated centrifugation at 4 ◦ C (3000 × g, 5 min), and washed thrice with deionized water. Subsequently the crude enzyme was extracted from cell pellet with ultrasonic processor (VC 130PB, Sonics and Materials Inc., USA). The enzyme activity was determined at standard assay conditions. The activity was expressed as percent relative activity with respect to maximum activity, which was considered as 100%. 2.4. Culture conditions for enzyme production The bacterial strain was cultivated in medium containing (w/v) 1.0% NaCl, 1.0% peptone, 0.5% yeast extract, 0.4% mandelic acid (pH 7.0). After pre-cultivated for 12 h, the strain was inoculated into 500 mL of Erlenmeyer flasks containing 100 mL of medium, and shaken at 30 ◦ C with 120 rpm for 18 h. The cells were harvested by refrigerated centrifugation at 4 ◦ C (3000 × g, 5 min), washed with deionized water three times. 2.5. Purification of the enzyme All purification steps were carried out at 4 ◦ C, and deionized water was used unless otherwise stated. The cell pellet was resuspended in a minimum volume of deionized water and then broken by ultrasonic processor. Cell debris was removed by centrifuging for 20 min at 10,000 × g, and the supernatant was used as the crude enzyme. Ammonium sulfate was added by a slow addition to 70% saturation with stirring. The precipitate formed and was collected by centrifugation (10,000 × g, 20 min), then dissolved in 20% saturation of ammonium sulfate solution preliminarily prepared in an ice bath. After centrifugation (10,000 × g, 20 min), the supernatant was applied to a TOYOPEARL Butyl-650 M column (1.6 cm × 25 cm) equilibrated with 50 mM phosphate buffer (pH 7.0) containing 20% saturation of ammonium sulfate. After removing the unbound proteins through washing step, the column was eluted with the same
buffer containing 20% to 0% saturation of ammonium sulfate at a flow rate of 1 mL/min and the active fractions were collected. 2.6. Biochemical properties of the enzyme The effect of temperature on enzyme activity was studied from 20 ◦ C to 60 ◦ C in 50 mM sodium phosphate buffer (pH 8.5). The effect of pH on enzyme activity was studied in the pH range of 5.0–10.0 at 30 ◦ C. The maximum residue enzyme activity was considered as 100%. The pH stability of the enzyme was examined by incubating the enzyme at 30 ◦ C in various buffers from pH 6.0 to 9.0. The buffers were used as follows: 50 mM citric acid-disodium hydrogen phosphate buffer, pH 5.0–8.0; 50 mM Tris–HCl buffer, pH 8.0–9.0; 50 mM glycine-NaOH buffer, pH 9.0–10.0. For measuring the thermal stability, the enzyme was incubated at 30 ◦ C and 4 ◦ C in 50 mM Tris–HCl buffer (pH 8.5). The activity of the enzyme in the beginning of the test was considered as 100% activity. Kinetic parameters (Km and kcat ) of the enzyme for (R)-mandelic acid, NAD+ and NADP+ were determined using Lineweaver-Burk method at pH 8.5 and 30 ◦ C [15,16]. For (R)-mandelic acid, the activity assay was performed in a mixture containing various concentrations of (R)-mandelic acid ranging from 0.02 mM to 2.75 mM and 125.0 M of NAD+ . For NAD+ and NADP+ , 2.75 mM of (R)-mandelic acid and NAD+ in a range of 1.0 M to 125.0 M, as well as NADP+ in a range of 0.9 M to 112.5 M were used for assaying activity. The influence of metal ions on the enzyme activity was investigated using chloride salts of Li+ , Na+ , K+ , Mg2+ , Ca2+ , Mn2+ , Ni2+ , Zn2+ and Hg2+ in a final concentration of 1 mM and 10 mM. The effect of EDTA, DTT in a final concentration of 10 mM and 50 mM, and SDS, Triton X-100 in a final concentration of 0.25% and 0.75% on enzyme activity were separately studied. The activity of enzyme without any additive was taken as 100%. 2.7. Assay of the enzyme activity The enzyme activity was assayed spectrophotometrically at 30 ◦ C by monitoring the increase in the absorbance of NADH at 340 nm. The reaction mixture containing 100 L of 66 mM substrate, 40 L of 3 mM NAD+ , 2 mL of 50 mM sodium phosphate buffer (pH 8.0) and 100 L of enzyme solution. A molar absorption coefficient of NADH 340 = 6.22 × 103 /M cm was used [17]. One unit of enzyme activity was defined as the amount of enzyme reducing 1 mol of NADH generation per minute under the above reaction conditions [7]. An equal concentration of (R)-mandelate and (S)mandelate, instead of the substrate, were used to determine the substrate specificity. 2.8. Protein quantification and determination of molecular mass Protein concentrations were determined by the Bradford method using bovine serum albumin as a standard protein [18]. The molecular mass of the purified enzyme was determined by SDS–polyacrylamide gel electrophoresis (PAGE). SDS–PAGE was performed as described by Laemmli using a 5% stacking gel and a 10% separating gel [19]. The molecular mass of the enzyme was calculated with GelAnalyzer (software, GelAnalyzer.com) [20]. 2.9. High performance liquid chromatography (HPLC) High performance liquid chromatography (HPLC) (LC-10AT; Shimadzu, Kyoto, Japan) was used for detecting the concentration of mandelic acid, benzoylformic acid and mandelic acid enantiomers. All reaction liquids were centrifuged at 10,000 × g for 10 min prior to analyze. Mandelic acid and benzoylformic acid were detected by a Shimadzu VP-ODS C18 column (150 mm × 4.6 mm) with a mobile phase of methanol-phosphate buffer (6.6 g/L
J. Wang et al. / Journal of Molecular Catalysis B: Enzymatic 120 (2015) 23–27
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Na2 HPO4 , 6.8 g/L KH2 PO4 ) (1:9, v/v) at a flow rate of 1 mL/min, detected with a Shimadzu SPD-10A detector at 230 nm in room temperature. The mandelic acid enantiomers were separated on a chiral HPLC column ␥-CD (150 mm × 4.6 mm, YMC, Kyoto, Japan) with a mobile phase of phosphate buffer (6.6 g/L Na2 HPO4 , 6.8 g/L KH2 PO4 )-ethanol-acetonitrile (65:20:15, v/v) at a flow rate of 1 mL/min, detected at 230 nm in room temperature.
3. Results and discussion 3.1. Molecular and morphological identification The 16S rDNA sequence of the strain isolated has been deposited in NCBI GenBank with accession number KP204477. According to 16S rDNA sequence analysis, the strain we obtained was identified as P. putida. We named the strain as P. putida NUST506. The colony of the strain was convex with a rough undulate edge on nutrient agar plate. From the TEM image (Fig. 1), the cell was rod shaped (approximately 0.6–0.9 m wide and 1.7–2.3 m long) with polar flagella. The morphological identification of the strain isolated was in agreement with the description of Palleroni [21]. As shown in Fig. 1, there was some viscous mixture around the cell. The viscous mixture may be viscous exopolysaccharide produced by the P. putida [22,23].
3.2. The bacteria growth and enzyme production Fig. 2 showed the effect of culture time on bacteria growth and on enzyme product yield. From the zero to the eighth hour, no significant bacterial growth was observed. From the eighth to twentieth hour, the bacteria grew into a logarithmic phase. During the phase, the bacteria had an exponential growth last for 12 h with a final cell concentration value of 1.96. After 20 h of cultivation, the growth curves of the cell entered the stationary phase, with the cell density maintained at a level of 1.98 in OD600 value. The enzyme product yield was gradually increased from the fourteenth hour to the eighteenth hour with relative yield of 79% to 100%. However, the enzyme production yield began getting down after 18 h of cultivation. Finally, the product yield decreased to 61% at the twenty-second hour.
Fig. 1. TEM image of the identified strain.
Fig. 2. Effect of culture time on enzyme production.
3.3. Purification of the enzyme As summarized in Table 1, the crude enzyme with 87.5 mg protein showed a specific activity of 0.03 U/mg. After ammonium sulfate precipitation, a specific activity of 0.03 U/mg with 92.0% recovery was obtained. After the final purification step, the enzyme was purified 11.0 folds with a specific activity of 0.33 U/mg. The purified enzyme was demonstrated as apparent single protein band on SDS–PAGE and its molecular weight was estimated to be 61 kDa (Fig. 3). 3.4. Effects of pH and temperature on the enzyme activity and stability The effect of pH on the activity of enzyme was analyzed at 30 ◦ C in pH buffer range of 5.0 to 10.0. Maximum activity was measured at pH 8.5 (Fig. 4). The influence of temperature on the enzyme activity was investigated over a temperature range from 20 ◦ C to 70 ◦ C at
Fig. 3. SDS–PAGE following the various purification steps of the enzyme. Lane 1, purified enzyme from TOYOPEAEL butyl hydrophobic chromatography; Lane 2, enzyme after ammonium sulfate treatment; Lane 3, crude enzyme extract; Lane M, molecular standard marker.
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J. Wang et al. / Journal of Molecular Catalysis B: Enzymatic 120 (2015) 23–27
Table 1 Purification of the RMDH. Purification step
Volume (mL)
Total protein (mg)
Total activity (U)
Specific activity (U/mg)
Yield (%)
Purification fold
Crude extract Ammonium sulfate treatment Hydrophobic interaction chromatography
5.5 11.5 7.5
87.5 70.9 6.1
2.5 2.3 2.0
0.03 0.03 0.33
100.0 92.0 80.0
1.0 1.0 11.0
Fig. 4. Effect of pH on activity of the purified enzyme. The maximum residue enzyme activity was considered as 100%. Each value is mean ± SD of three determinations.
pH 8.5. The optimum temperature for the enzyme was 30 ◦ C. The relative activity at 20 ◦ C and 35 ◦ C were approximately 95% and 75% compared with the activity at 30 ◦ C, respectively (Fig. 5). The effect of pH on the enzyme stability was shown in Fig. 6. The endurance of enzyme in acidic buffer was stronger than in alkali buffer. Taken the relative initial activity as 100%, the stability dropped significantly at pH 8.5 and pH 9.0, the enzyme lost 85% and 94% of the activity when incubated for 3.5 h at 30 ◦ C. On the other hand, the enzyme lost 11% of its activity in pH 6.0 after 3.5 h at 30 ◦ C. After 96 h of incubation in 4 ◦ C, 78% of the enzyme activity was retained in pH 6.0, but only 30% of enzyme activity was retained in pH 8.5 (data not shown). The result suggested that the enzyme had better storage stability in pH 6.0 than in pH 8.5. In the thermal
Fig. 6. Effects of pH and temperature on stability of the purified enzyme. The purified enzyme was incubated at 30 ◦ C in various buffer of pH 6.0 (), pH 7.0 ( ), pH 8.0 ( ), pH 8.5 (), pH 9.0 ( ) for 3.5 h, and at 4 ◦ C in pH 6.0 (䊏), pH 8.5 ( ). The initial enzyme prior to incubate was used to determine the 100% activity value.
stability analysis, the enzyme activity decreased markedly at the optimal temperature 30 ◦ C and the optimal pH 8.5, only 31% and 20% of its activity were retained after 0.5 h and 1 h of incubation. Among the substrate specificity test, the RMDH displayed catalytic activity for (R)-mandelate, but no catalytic activity was observed for (S)-mandelate, suggesting that the RMDH is highly specific for (R)mandelate. The RMDH showed catalytic activity for (R)-mandelate both in the presence of NAD+ and NADP+ . The Km and kcat for (R)mandelic acid were 2.0 × 10−2 mM and 0.9 s−1 , respectively. The Km was 1.8 × 10−2 mM for NAD+ , and 1.5 × 10−2 mM for NADP+ . The kcat was 0.9 s−1 for NAD+ , and 0.3 s−1 for NADP+ . The kcat /Km was 5.0 × 104 M−1 /s for NAD+ and 2.0 × 104 M−1 /s for NADP+ , respectively. Compared with NADP+ , NAD+ displayed higher kcat /Km . The result suggested that the RMDH has higher catalytic efficiency in the presence of NAD+ . 3.5. Effects of metal ions and chemicals on the enzyme activity
Fig. 5. Effect of temperature on activity of the purified enzyme. The maximum residue enzyme activity was considered as 100%. Each value is mean ± SD of three determinations.
The effects of metal ions and chemical reagents on enzyme activity were summarized in Table 2. The enzyme activity was notablely enhanced by K+ up to 144% and 169% with the concentration of 1 mM and 10 mM, respectively. The enzyme activity was slightly increased when 1 mM of Mn2+ was used. 1 mM of Na+ , Ca2+ and Li+ did not obviously affect the enzyme activity. However, 10 mM of Mn2+ and Ni2+ demonstrated high inhibition of the enzyme activity. Hg2+ and Zn2+ strongly inhibited the enzyme activity both in concentration of 1 mM and 10 mM. In addition, EDTA inhibited the enzyme activity about 68% and 89% in the concentration of 10 mM and 50 mM, respectively. DTT had a positive effect on enzyme activity, showing an enhancement of 26% and 33% in the concentration of 10 mM and 50 mM. 0.25% of TritonX-100 had slight improvement on enzyme activity, whereas 0.75% of TritonX-100 slightly inhibited the enzyme activity. Moreover, SDS strongly inhibited the enzyme activity in both the concentration of 0.25% and 0.75%.
J. Wang et al. / Journal of Molecular Catalysis B: Enzymatic 120 (2015) 23–27 Table 2 Effects of metal ions and chemicals on the RMDH activity. The residual activities were expressed as percent relative activity by comparison with reaction mixture in the absence of metal ions and chemicals. Metal ion
Na+ K+ Mg2+ Ni2+ Li+ Mn2+ Ca2+ Hg2+ Zn2+ Chemical
Residual activity (%) 1 mM final concentration (±SD)
10 mM final concentration (±SD)
100 ± 0.4 144 ± 13.4 81 ± 2.3 71 ± 2.4 100 ± 1.8 121 ± 9.0 96 ± 3.4 1.3 ± 1.2 3 ± 1.3
112 ± 0.8 169 ± 4.7 79 ± 5.7 37 ± 1.7 83 ± 6.1 39 ± 3.1 77 ± 1.4 0 0
Residual activity (%) 10 mM final concentration (±SD)
50 mM final concentration (±SD)
EDTA DTT
32 ± 2.7 126 ± 16.9 0.25% Final concentration (±SD)
11 ± 1.8 133 ± 12.0 0.75% Final concentration (±SD)
SDS (w/v) TritonX-100 (v/v)
27 ± 1.0 115 ± 3.3
23 ± 2.1 89 ± 4.0
4. Conclusion In the study, a novel RMDH was purified and characterized from a screened strain P. putida NUST506. The purified enzyme preparation was homogeneous with a specific activity of 0.33 U/mg. The molecular weight of the enzyme was calculated to be 61 kDa by SDS–PAGE. The enzyme has optimum activity at pH 8.5 and 30 ◦ C, and showed higher stability in acidic buffer than in alkali buffer. The enzyme activity was increased by K+ and DTT, but strongly inhibited by Zn2+ , Hg2+ , SDS and EDTA. The enzyme exhibited activity for (R)mandelate, but no activity was detected for (S)-mandelate. Both NAD+ and NADP+ are the coenzyme of the RMDH, but the enzyme
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has higher catalytic efficiency in the presence of NAD+ . Future studies will focus on analyzing the reductive reaction producing (R)-mandelic acid catalyzed by the RMDH, as well as determining the primary structure of the RMDH by Edman degradation or peptide mass fingerprinting and cloning encoded gene of enzyme in order to understand the mechanism of its activity. References [1] C. Mateo, A. Chmura, S. Rustler, F. van Rantwijk, A. Stolz, R.A. Sheldon, Tetrahedron: Asymmetry 17 (2006) 320–323. [2] P.T. Grover, N.N. Bhongle, S.A. Wald, C.H. Senanayake, J. Org. Chem. 65 (2000) 6283–6287. [3] R.N. Patel, Curr. Opin. Biotechnol. 12 (2001) 587–604. [4] F. Effenberger, B. Horsch, S. Forster, T. Ziegler, Tetrahedron. Lett. 31 (1990) 1249–1252. [5] S. Rustler, H. Motejadded, J. Altenbuchner, A. Stolz, Appl. Microbiol. Biotechnol. 80 (2008) 87–97. [6] S.H. Lee, J.H. Choi, S.H. Park, J.I. Choi, S.Y. Lee, Enzyme. Microb. Technol. 35 (2004) 429–436. [7] P. Wang, D. Li, J. Yang, L. Jiang, J. Feng, C. Yang, R. Shi, J. Taiwan Inst. Chem. Eng. 45 (2014) 744–748. [8] B.Y. Kim, K.C. Hwang, H.S. Song, N. Chung, W.G. Bang, Biotechnol. Lett. 22 (2000) 1871–1875. [9] H. Huang, J. Xu, Biochem. Eng. J. 30 (2006) 11–15. [10] K. Miyamoto, H. Ohta, Biotechnol. Lett. 14 (1992) 363–366. [11] E. Takahashi, K. Nakamichi, M. Furui, J. Ferment. Bioeng. 80 (1995) 247–250. [12] T. Nagasawa, T. Yoshida, K. Ishida, The Novel Gibberella Nitrilase for Stereo Selective Syntheses of Carboxylic Acids, Mitsubishi Rayon Co., Ltd, Japan, 2009, pp. 16. [13] A. Banerjee, S. Dubey, P. Kaul, B. Barse, M. Piotrowski, U.C. Banerjee, Mol. Biotechnol. 41 (2009) 35–41. [14] W. Wu, L. Xing, B. Zhou, Z. Lin, Microb. Cell Fact 10 (2011) 1–8. [15] D.J. Holme, H. Peck, Analytical Biochemistry, third edition, Addison Weskey Longman, New York, 1998, pp. 264–265. [16] J.R. Lorsch, in: L. Jon (Ed.), Methods in Enzymology, Academic Press, California, 2014, pp. 3–15. [17] H.J. Park, C.O. Reiser, S. Kondruweit, H. Erdmann, R.D. Schmid, M. Sprinzl, Eur. J. Biochem. 205 (1992) 881–885. [18] M.M. Bradford, Anal. Biochem. 72 (1976) 248–254. [19] U.K. Laemmli, Nature 227 (1970) 680–685. [20] V.S. Skosyrev, G.V. Vasil’Eva, M.G. Lomaeva, L.V. Malachova, V.N. Antipova, V.G. Bezlepkin, Russ. J. Genet. 49 (2013) 464–469. [21] N.J. Palleroni, Environ. Microbiol. 12 (2010) 1377–1383. [22] T.B. May, A.M. Chakrabarty, Trends Microbiol. 2 (1994) 151–157. [23] P.I. Nikel, E. Martínez-García, V. de Lorenzo, Nat. Rev. Microbiol. 12 (2014) 368–379.
Contents lists available at ScienceDirect
Journal of Molecular Catalysis B: Enzymatic journal homepage: www.elsevier.com/locate/molcatb
Characterization of a novel (R)-mandelate dehydrogenase from Pseudomonas putida NUST506 Jizhong Wang, Jing Feng, Weile Li, Chengli Yang, Xing Chen, Bingxin Bao, Junfang Yang, Peng Wang, Dali Li ∗ , Ruofu Shi Department of Bioengineering, Nanjing University of Science & Technology, Nanjing 210094, Jiangsu, People’s Republic of China
a r t i c l e
i n f o
Article history: Received 23 January 2015 Received in revised form 29 April 2015 Accepted 30 April 2015 Available online 11 May 2015 Keywords: Characterization (R)-Mandelate dehydrogenase Biocatalysis Pseudomonas putida Purification
a b s t r a c t (R)-Mandelate dehydrogenase (RMDH) has the potential to produce chiral mandelic acid. The present work reports isolation and identification of a Pseudomonas putida NUST506 contained RMDH. The strain was rod shaped with polar flagella, approximately 0.6–0.9 m wide and 1.7–2.3 m long. The RMDH was purified from the strain. The molecular weight of the enzyme was calculated to be 61 kDa. The optimal conditions for RMDH were pH 8.5 and 30 ◦ C. At the optimum condition, the Km and kcat of the RMDH were 2.0 × 10−2 mM and 0.9 s−1 for (R)-mandelic acid, 1.8 × 10−2 mM and 0.9 s−1 for NAD+ , as well as 1.5 × 10−2 mM and 0.3 s−1 for NADP+ , respectively. The enzyme activity was increased by K+ and dithiothreitol (DTT), but obviously inhibited by Zn2+ , Hg2+ , sodium dodecyl sulfate (SDS) and ethylenediamine tetra-acetic acid (EDTA). © 2015 Published by Elsevier B.V.
1. Introduction
2. Materials and methods
During the past few years, considerable attention has been focused on the field of chiral mandelic acid, because of its important role as building blocks widely used in the synthesis of chiral pharmaceutical and biochemical materials [1–3]. Many methods have been reported for the production of chiral mandelic acid. The chiral mandelic acid can be obtained from racemic cyanohydrins by enzyme-catalyzed enantioselective degradation [4,5], from methyl mandelate by lipase-catalyzed hydrolysis [6], or through selectively consuming (S)- or (R)-mandelic acid from the racemic mandelic acid by biotransformation and leaving (R)- or (S)-mandelic acid in reaction system [7–9]. The microbial cells enantioselective degradation of (R)-mandelic acid have been reported with Alcaligenes bronchisepticus [10], Pseudomonas polycolor [11], Gibberella fujikuroi [12], and Pseudomonas putida [8,9,13]. However, to the best of our knowledge, little is known about the characterization of RMDH. In this study, we isolated a strain P. putida NUST506 containing NAD+ -dependent RMDH and presented the purification and characterization the enzyme from the strain.
2.1. Chemicals
∗ Corresponding author. Tel.: +86 025 84315512; fax: +86 025 84315512. E-mail address: [email protected] (D. Li). http://dx.doi.org/10.1016/j.molcatb.2015.04.017 1381-1177/© 2015 Published by Elsevier B.V.
The chemicals used were of highest grade and purchased from Sinopharm Chemical Reagent Corporation (Nanjing, China). TOYOPEARL Butyl-650M was obtained from Tosoh Corporation (Osaka, Japan). The blue plus protein marker was purchased from Beijing TransGen Biotech Co. Ltd (Beijing, China). 2.2. Microorganism The microorganism used in the study was isolated from soil around the schoolyard of Nanjing University of Science and Technology (Nanjing, China). The isolates have been screened on agar plate containing (w/v) 0.4% mandelic acid, 0.4% peptone, 0.1% KH2PO4 , 0.06% MgSO4 , 0.02% FeSO4 , 0.02% MnSO4 , 0.04% CaC12 , 2% agar, and the pH was adjusted to 7.0 using 1 M NaOH. The samples were cultivated on plates and incubated at 30 ◦ C for 48 h. The bacterial colonies observed on the plates were considered as the microorganism we wanted and were used for further studies. The strain was identified as P. putida by 16S rDNA sequence. The sequence of the 16S rDNA was amplified using a polymerase chain reaction (PCR) with the forward primer (5 -AGAGTTTGATCCTGGCTCAG-3 ) and the reverse primer (5 TACGGCTACCTTGTTACGACTT-3 ). The PCR conditions used were an initial denaturation at 95 ◦ C for 3 min, followed by 35 cycles
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J. Wang et al. / Journal of Molecular Catalysis B: Enzymatic 120 (2015) 23–27
of denaturation at 94 ◦ C for 45 s, annealing at 55 ◦ C for 30 s, and extension at 72 ◦ C for 90 s, and a final extension at 72 ◦ C for 5 min. The DNA nucleotide sequence was determined by GenScript Corporation (Nanjing, China). The sequence obtained was analyzed for similarities to other known sequences found in the GenBank database using BLAST program of the NCBI database. The cells were cultivated on LB agar plates with 0.4% mandelic acid addition at 30 ◦ C for 12 h. Then the cells were collected and washed in 10 mM potassium phosphate buffer (pH 7.4). According to the similar method described previously [14], the cell morphology was examined with a transmission electron microscope (TEM, H-7650; Hitachi, Tokyo, Japan). 2.3. Partial optimization of culture conditions To determine the effect of culture time on the cell growth and on the RMDH product yield, two experiments were performed. One is recording the cell growth curve by detecting the changes of biomass concentration in fermentation broth, the other is analyzing the enzyme product yield by assaying total enzyme activity of cultivation samples. The stock culture was inoculated into 1 L Erlenmeyer flask containing 250 mL of medium (w/v, 1.0% NaCl, 1.0% peptone, 0.5% yeast extract, 0.4% mandelic acid, pH 7.0). After inoculation, the culture was incubated with shaking cultivation at 30 ◦ C and 120 rpm for up to 22 h. Two milliliters of medium were sampled at 1 h intervals and analyzed for biomass concentration by measuring the optical density value at a wavelength of 600 nm (OD600 ) using a spectrophotometer (U-1800; Hitachi, Tokyo, Japan). Fifty milliliters of fermentation broth were sampled at 2 h intervals from the fourteenth hour to the twenty-second hour of the culture broth, and rapidly cooled in an ice bath. Then the cells were collected by refrigerated centrifugation at 4 ◦ C (3000 × g, 5 min), and washed thrice with deionized water. Subsequently the crude enzyme was extracted from cell pellet with ultrasonic processor (VC 130PB, Sonics and Materials Inc., USA). The enzyme activity was determined at standard assay conditions. The activity was expressed as percent relative activity with respect to maximum activity, which was considered as 100%. 2.4. Culture conditions for enzyme production The bacterial strain was cultivated in medium containing (w/v) 1.0% NaCl, 1.0% peptone, 0.5% yeast extract, 0.4% mandelic acid (pH 7.0). After pre-cultivated for 12 h, the strain was inoculated into 500 mL of Erlenmeyer flasks containing 100 mL of medium, and shaken at 30 ◦ C with 120 rpm for 18 h. The cells were harvested by refrigerated centrifugation at 4 ◦ C (3000 × g, 5 min), washed with deionized water three times. 2.5. Purification of the enzyme All purification steps were carried out at 4 ◦ C, and deionized water was used unless otherwise stated. The cell pellet was resuspended in a minimum volume of deionized water and then broken by ultrasonic processor. Cell debris was removed by centrifuging for 20 min at 10,000 × g, and the supernatant was used as the crude enzyme. Ammonium sulfate was added by a slow addition to 70% saturation with stirring. The precipitate formed and was collected by centrifugation (10,000 × g, 20 min), then dissolved in 20% saturation of ammonium sulfate solution preliminarily prepared in an ice bath. After centrifugation (10,000 × g, 20 min), the supernatant was applied to a TOYOPEARL Butyl-650 M column (1.6 cm × 25 cm) equilibrated with 50 mM phosphate buffer (pH 7.0) containing 20% saturation of ammonium sulfate. After removing the unbound proteins through washing step, the column was eluted with the same
buffer containing 20% to 0% saturation of ammonium sulfate at a flow rate of 1 mL/min and the active fractions were collected. 2.6. Biochemical properties of the enzyme The effect of temperature on enzyme activity was studied from 20 ◦ C to 60 ◦ C in 50 mM sodium phosphate buffer (pH 8.5). The effect of pH on enzyme activity was studied in the pH range of 5.0–10.0 at 30 ◦ C. The maximum residue enzyme activity was considered as 100%. The pH stability of the enzyme was examined by incubating the enzyme at 30 ◦ C in various buffers from pH 6.0 to 9.0. The buffers were used as follows: 50 mM citric acid-disodium hydrogen phosphate buffer, pH 5.0–8.0; 50 mM Tris–HCl buffer, pH 8.0–9.0; 50 mM glycine-NaOH buffer, pH 9.0–10.0. For measuring the thermal stability, the enzyme was incubated at 30 ◦ C and 4 ◦ C in 50 mM Tris–HCl buffer (pH 8.5). The activity of the enzyme in the beginning of the test was considered as 100% activity. Kinetic parameters (Km and kcat ) of the enzyme for (R)-mandelic acid, NAD+ and NADP+ were determined using Lineweaver-Burk method at pH 8.5 and 30 ◦ C [15,16]. For (R)-mandelic acid, the activity assay was performed in a mixture containing various concentrations of (R)-mandelic acid ranging from 0.02 mM to 2.75 mM and 125.0 M of NAD+ . For NAD+ and NADP+ , 2.75 mM of (R)-mandelic acid and NAD+ in a range of 1.0 M to 125.0 M, as well as NADP+ in a range of 0.9 M to 112.5 M were used for assaying activity. The influence of metal ions on the enzyme activity was investigated using chloride salts of Li+ , Na+ , K+ , Mg2+ , Ca2+ , Mn2+ , Ni2+ , Zn2+ and Hg2+ in a final concentration of 1 mM and 10 mM. The effect of EDTA, DTT in a final concentration of 10 mM and 50 mM, and SDS, Triton X-100 in a final concentration of 0.25% and 0.75% on enzyme activity were separately studied. The activity of enzyme without any additive was taken as 100%. 2.7. Assay of the enzyme activity The enzyme activity was assayed spectrophotometrically at 30 ◦ C by monitoring the increase in the absorbance of NADH at 340 nm. The reaction mixture containing 100 L of 66 mM substrate, 40 L of 3 mM NAD+ , 2 mL of 50 mM sodium phosphate buffer (pH 8.0) and 100 L of enzyme solution. A molar absorption coefficient of NADH 340 = 6.22 × 103 /M cm was used [17]. One unit of enzyme activity was defined as the amount of enzyme reducing 1 mol of NADH generation per minute under the above reaction conditions [7]. An equal concentration of (R)-mandelate and (S)mandelate, instead of the substrate, were used to determine the substrate specificity. 2.8. Protein quantification and determination of molecular mass Protein concentrations were determined by the Bradford method using bovine serum albumin as a standard protein [18]. The molecular mass of the purified enzyme was determined by SDS–polyacrylamide gel electrophoresis (PAGE). SDS–PAGE was performed as described by Laemmli using a 5% stacking gel and a 10% separating gel [19]. The molecular mass of the enzyme was calculated with GelAnalyzer (software, GelAnalyzer.com) [20]. 2.9. High performance liquid chromatography (HPLC) High performance liquid chromatography (HPLC) (LC-10AT; Shimadzu, Kyoto, Japan) was used for detecting the concentration of mandelic acid, benzoylformic acid and mandelic acid enantiomers. All reaction liquids were centrifuged at 10,000 × g for 10 min prior to analyze. Mandelic acid and benzoylformic acid were detected by a Shimadzu VP-ODS C18 column (150 mm × 4.6 mm) with a mobile phase of methanol-phosphate buffer (6.6 g/L
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Na2 HPO4 , 6.8 g/L KH2 PO4 ) (1:9, v/v) at a flow rate of 1 mL/min, detected with a Shimadzu SPD-10A detector at 230 nm in room temperature. The mandelic acid enantiomers were separated on a chiral HPLC column ␥-CD (150 mm × 4.6 mm, YMC, Kyoto, Japan) with a mobile phase of phosphate buffer (6.6 g/L Na2 HPO4 , 6.8 g/L KH2 PO4 )-ethanol-acetonitrile (65:20:15, v/v) at a flow rate of 1 mL/min, detected at 230 nm in room temperature.
3. Results and discussion 3.1. Molecular and morphological identification The 16S rDNA sequence of the strain isolated has been deposited in NCBI GenBank with accession number KP204477. According to 16S rDNA sequence analysis, the strain we obtained was identified as P. putida. We named the strain as P. putida NUST506. The colony of the strain was convex with a rough undulate edge on nutrient agar plate. From the TEM image (Fig. 1), the cell was rod shaped (approximately 0.6–0.9 m wide and 1.7–2.3 m long) with polar flagella. The morphological identification of the strain isolated was in agreement with the description of Palleroni [21]. As shown in Fig. 1, there was some viscous mixture around the cell. The viscous mixture may be viscous exopolysaccharide produced by the P. putida [22,23].
3.2. The bacteria growth and enzyme production Fig. 2 showed the effect of culture time on bacteria growth and on enzyme product yield. From the zero to the eighth hour, no significant bacterial growth was observed. From the eighth to twentieth hour, the bacteria grew into a logarithmic phase. During the phase, the bacteria had an exponential growth last for 12 h with a final cell concentration value of 1.96. After 20 h of cultivation, the growth curves of the cell entered the stationary phase, with the cell density maintained at a level of 1.98 in OD600 value. The enzyme product yield was gradually increased from the fourteenth hour to the eighteenth hour with relative yield of 79% to 100%. However, the enzyme production yield began getting down after 18 h of cultivation. Finally, the product yield decreased to 61% at the twenty-second hour.
Fig. 1. TEM image of the identified strain.
Fig. 2. Effect of culture time on enzyme production.
3.3. Purification of the enzyme As summarized in Table 1, the crude enzyme with 87.5 mg protein showed a specific activity of 0.03 U/mg. After ammonium sulfate precipitation, a specific activity of 0.03 U/mg with 92.0% recovery was obtained. After the final purification step, the enzyme was purified 11.0 folds with a specific activity of 0.33 U/mg. The purified enzyme was demonstrated as apparent single protein band on SDS–PAGE and its molecular weight was estimated to be 61 kDa (Fig. 3). 3.4. Effects of pH and temperature on the enzyme activity and stability The effect of pH on the activity of enzyme was analyzed at 30 ◦ C in pH buffer range of 5.0 to 10.0. Maximum activity was measured at pH 8.5 (Fig. 4). The influence of temperature on the enzyme activity was investigated over a temperature range from 20 ◦ C to 70 ◦ C at
Fig. 3. SDS–PAGE following the various purification steps of the enzyme. Lane 1, purified enzyme from TOYOPEAEL butyl hydrophobic chromatography; Lane 2, enzyme after ammonium sulfate treatment; Lane 3, crude enzyme extract; Lane M, molecular standard marker.
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J. Wang et al. / Journal of Molecular Catalysis B: Enzymatic 120 (2015) 23–27
Table 1 Purification of the RMDH. Purification step
Volume (mL)
Total protein (mg)
Total activity (U)
Specific activity (U/mg)
Yield (%)
Purification fold
Crude extract Ammonium sulfate treatment Hydrophobic interaction chromatography
5.5 11.5 7.5
87.5 70.9 6.1
2.5 2.3 2.0
0.03 0.03 0.33
100.0 92.0 80.0
1.0 1.0 11.0
Fig. 4. Effect of pH on activity of the purified enzyme. The maximum residue enzyme activity was considered as 100%. Each value is mean ± SD of three determinations.
pH 8.5. The optimum temperature for the enzyme was 30 ◦ C. The relative activity at 20 ◦ C and 35 ◦ C were approximately 95% and 75% compared with the activity at 30 ◦ C, respectively (Fig. 5). The effect of pH on the enzyme stability was shown in Fig. 6. The endurance of enzyme in acidic buffer was stronger than in alkali buffer. Taken the relative initial activity as 100%, the stability dropped significantly at pH 8.5 and pH 9.0, the enzyme lost 85% and 94% of the activity when incubated for 3.5 h at 30 ◦ C. On the other hand, the enzyme lost 11% of its activity in pH 6.0 after 3.5 h at 30 ◦ C. After 96 h of incubation in 4 ◦ C, 78% of the enzyme activity was retained in pH 6.0, but only 30% of enzyme activity was retained in pH 8.5 (data not shown). The result suggested that the enzyme had better storage stability in pH 6.0 than in pH 8.5. In the thermal
Fig. 6. Effects of pH and temperature on stability of the purified enzyme. The purified enzyme was incubated at 30 ◦ C in various buffer of pH 6.0 (), pH 7.0 ( ), pH 8.0 ( ), pH 8.5 (), pH 9.0 ( ) for 3.5 h, and at 4 ◦ C in pH 6.0 (䊏), pH 8.5 ( ). The initial enzyme prior to incubate was used to determine the 100% activity value.
stability analysis, the enzyme activity decreased markedly at the optimal temperature 30 ◦ C and the optimal pH 8.5, only 31% and 20% of its activity were retained after 0.5 h and 1 h of incubation. Among the substrate specificity test, the RMDH displayed catalytic activity for (R)-mandelate, but no catalytic activity was observed for (S)-mandelate, suggesting that the RMDH is highly specific for (R)mandelate. The RMDH showed catalytic activity for (R)-mandelate both in the presence of NAD+ and NADP+ . The Km and kcat for (R)mandelic acid were 2.0 × 10−2 mM and 0.9 s−1 , respectively. The Km was 1.8 × 10−2 mM for NAD+ , and 1.5 × 10−2 mM for NADP+ . The kcat was 0.9 s−1 for NAD+ , and 0.3 s−1 for NADP+ . The kcat /Km was 5.0 × 104 M−1 /s for NAD+ and 2.0 × 104 M−1 /s for NADP+ , respectively. Compared with NADP+ , NAD+ displayed higher kcat /Km . The result suggested that the RMDH has higher catalytic efficiency in the presence of NAD+ . 3.5. Effects of metal ions and chemicals on the enzyme activity
Fig. 5. Effect of temperature on activity of the purified enzyme. The maximum residue enzyme activity was considered as 100%. Each value is mean ± SD of three determinations.
The effects of metal ions and chemical reagents on enzyme activity were summarized in Table 2. The enzyme activity was notablely enhanced by K+ up to 144% and 169% with the concentration of 1 mM and 10 mM, respectively. The enzyme activity was slightly increased when 1 mM of Mn2+ was used. 1 mM of Na+ , Ca2+ and Li+ did not obviously affect the enzyme activity. However, 10 mM of Mn2+ and Ni2+ demonstrated high inhibition of the enzyme activity. Hg2+ and Zn2+ strongly inhibited the enzyme activity both in concentration of 1 mM and 10 mM. In addition, EDTA inhibited the enzyme activity about 68% and 89% in the concentration of 10 mM and 50 mM, respectively. DTT had a positive effect on enzyme activity, showing an enhancement of 26% and 33% in the concentration of 10 mM and 50 mM. 0.25% of TritonX-100 had slight improvement on enzyme activity, whereas 0.75% of TritonX-100 slightly inhibited the enzyme activity. Moreover, SDS strongly inhibited the enzyme activity in both the concentration of 0.25% and 0.75%.
J. Wang et al. / Journal of Molecular Catalysis B: Enzymatic 120 (2015) 23–27 Table 2 Effects of metal ions and chemicals on the RMDH activity. The residual activities were expressed as percent relative activity by comparison with reaction mixture in the absence of metal ions and chemicals. Metal ion
Na+ K+ Mg2+ Ni2+ Li+ Mn2+ Ca2+ Hg2+ Zn2+ Chemical
Residual activity (%) 1 mM final concentration (±SD)
10 mM final concentration (±SD)
100 ± 0.4 144 ± 13.4 81 ± 2.3 71 ± 2.4 100 ± 1.8 121 ± 9.0 96 ± 3.4 1.3 ± 1.2 3 ± 1.3
112 ± 0.8 169 ± 4.7 79 ± 5.7 37 ± 1.7 83 ± 6.1 39 ± 3.1 77 ± 1.4 0 0
Residual activity (%) 10 mM final concentration (±SD)
50 mM final concentration (±SD)
EDTA DTT
32 ± 2.7 126 ± 16.9 0.25% Final concentration (±SD)
11 ± 1.8 133 ± 12.0 0.75% Final concentration (±SD)
SDS (w/v) TritonX-100 (v/v)
27 ± 1.0 115 ± 3.3
23 ± 2.1 89 ± 4.0
4. Conclusion In the study, a novel RMDH was purified and characterized from a screened strain P. putida NUST506. The purified enzyme preparation was homogeneous with a specific activity of 0.33 U/mg. The molecular weight of the enzyme was calculated to be 61 kDa by SDS–PAGE. The enzyme has optimum activity at pH 8.5 and 30 ◦ C, and showed higher stability in acidic buffer than in alkali buffer. The enzyme activity was increased by K+ and DTT, but strongly inhibited by Zn2+ , Hg2+ , SDS and EDTA. The enzyme exhibited activity for (R)mandelate, but no activity was detected for (S)-mandelate. Both NAD+ and NADP+ are the coenzyme of the RMDH, but the enzyme
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has higher catalytic efficiency in the presence of NAD+ . Future studies will focus on analyzing the reductive reaction producing (R)-mandelic acid catalyzed by the RMDH, as well as determining the primary structure of the RMDH by Edman degradation or peptide mass fingerprinting and cloning encoded gene of enzyme in order to understand the mechanism of its activity. References [1] C. Mateo, A. Chmura, S. Rustler, F. van Rantwijk, A. Stolz, R.A. Sheldon, Tetrahedron: Asymmetry 17 (2006) 320–323. [2] P.T. Grover, N.N. Bhongle, S.A. Wald, C.H. Senanayake, J. Org. Chem. 65 (2000) 6283–6287. [3] R.N. Patel, Curr. Opin. Biotechnol. 12 (2001) 587–604. [4] F. Effenberger, B. Horsch, S. Forster, T. Ziegler, Tetrahedron. Lett. 31 (1990) 1249–1252. [5] S. Rustler, H. Motejadded, J. Altenbuchner, A. Stolz, Appl. Microbiol. Biotechnol. 80 (2008) 87–97. [6] S.H. Lee, J.H. Choi, S.H. Park, J.I. Choi, S.Y. Lee, Enzyme. Microb. Technol. 35 (2004) 429–436. [7] P. Wang, D. Li, J. Yang, L. Jiang, J. Feng, C. Yang, R. Shi, J. Taiwan Inst. Chem. Eng. 45 (2014) 744–748. [8] B.Y. Kim, K.C. Hwang, H.S. Song, N. Chung, W.G. Bang, Biotechnol. Lett. 22 (2000) 1871–1875. [9] H. Huang, J. Xu, Biochem. Eng. J. 30 (2006) 11–15. [10] K. Miyamoto, H. Ohta, Biotechnol. Lett. 14 (1992) 363–366. [11] E. Takahashi, K. Nakamichi, M. Furui, J. Ferment. Bioeng. 80 (1995) 247–250. [12] T. Nagasawa, T. Yoshida, K. Ishida, The Novel Gibberella Nitrilase for Stereo Selective Syntheses of Carboxylic Acids, Mitsubishi Rayon Co., Ltd, Japan, 2009, pp. 16. [13] A. Banerjee, S. Dubey, P. Kaul, B. Barse, M. Piotrowski, U.C. Banerjee, Mol. Biotechnol. 41 (2009) 35–41. [14] W. Wu, L. Xing, B. Zhou, Z. Lin, Microb. Cell Fact 10 (2011) 1–8. [15] D.J. Holme, H. Peck, Analytical Biochemistry, third edition, Addison Weskey Longman, New York, 1998, pp. 264–265. [16] J.R. Lorsch, in: L. Jon (Ed.), Methods in Enzymology, Academic Press, California, 2014, pp. 3–15. [17] H.J. Park, C.O. Reiser, S. Kondruweit, H. Erdmann, R.D. Schmid, M. Sprinzl, Eur. J. Biochem. 205 (1992) 881–885. [18] M.M. Bradford, Anal. Biochem. 72 (1976) 248–254. [19] U.K. Laemmli, Nature 227 (1970) 680–685. [20] V.S. Skosyrev, G.V. Vasil’Eva, M.G. Lomaeva, L.V. Malachova, V.N. Antipova, V.G. Bezlepkin, Russ. J. Genet. 49 (2013) 464–469. [21] N.J. Palleroni, Environ. Microbiol. 12 (2010) 1377–1383. [22] T.B. May, A.M. Chakrabarty, Trends Microbiol. 2 (1994) 151–157. [23] P.I. Nikel, E. Martínez-García, V. de Lorenzo, Nat. Rev. Microbiol. 12 (2014) 368–379.