Enzyme Kinetics of Different Types of Flavin-Dependent

Copyright © 2015 American Chemical Society .... The new drugs of 2018. Pharmaceutical industry productivity was robust in 2018, a year in which th...
0 downloads 0 Views 267KB Size
Subscriber access provided by CMU Libraries - http://library.cmich.edu

Letter

Enzyme Kinetics of Different Types of Flavin-Dependent Monooxygenases Determine the Observable Contaminant Stable Isotope Fractionation Reto S. Wijker, Sarah Pati, Josef Zeyer, and Thomas B. Hofstetter Environ. Sci. Technol. Lett., Just Accepted Manuscript • DOI: 10.1021/acs.estlett.5b00254 • Publication Date (Web): 16 Oct 2015 Downloaded from http://pubs.acs.org on October 20, 2015

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a free service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are accessible to all readers and citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

Environmental Science & Technology Letters is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 21

Environmental Science & Technology Letters

Enzyme Kinetics of Different Types of Flavin-Dependent Monooxygenases Determine the Observable Contaminant Stable Isotope Fractionation Reto S. Wijker,†, ¶ Sarah G. Pati,‡,† Josef Zeyer,† and Thomas B. Hofstetter∗,‡,† Institute of Biogeochemistry and Pollutant Dynamics (IBP), ETH Zürich, CH-8092 Zürich, Switzerland, and Eawag, Swiss Federal Institute of Aquatic Science and Technology, CH-8600 Dübendorf, Switzerland E-mail: thomas.hofstetter@eawag.ch

Fax: +41 58 765 50 28, Phone: +41 58 765 50 76

∗ To

whom correspondence should be addressed Zürich ‡ Eawag ¶ Present address: Division of Geological and Planetary Sciences, California Institute of Technology, Pasadena, CA 91125, USA † ETH

1 ACS Paragon Plus Environment

Environmental Science & Technology Letters

1

Abstract

2

The assessment of oxidative pollutant biotransformation by compound-specific isotope

3

analysis (CSIA) is often complicated by the variability of kinetic isotope effects associated

4

with carbon oxygenation in enzymatic reactions. Here, we illustrate how information on the

5

kinetics of oxidative biocatalysis by flavin-dependent monooxygenases (FMOs) enables one to

6

assess if CSIA could be applied for tracking contaminant biodegradation. In “cautious” FMOs,

7

which form reactive flavin (hydro)peroxide species after substrate binding, the monooxygena-

8

tion of organic compounds is not rate-determining and consequently does not lead to substrate

9

isotope fractionation. Conversely, “bold” FMOs generate hydroperoxides regardless of sub-

10

strate availability and substrate disappearance is thus subject to isotope fractionation trends,

11

which are typical for hydroxylation reactions. Because monooxygenations of aromatic moieties

12

are often initial steps of organic pollutant transformation, knowledge of the kinetics of FMOs

13

and other oxidative enzymes can support decisions regarding the use of CSIA.

2 ACS Paragon Plus Environment

Page 2 of 21

Page 3 of 21

Environmental Science & Technology Letters

14

Introduction

15

Compound specific isotope analysis (CSIA) is an increasingly used approach for assessing (bio)de-

16

gradation of organic pollutants in the environment. 1–5 Bonding changes during initial steps of

17

transformation, especially bond cleavage reactions, give rise to kinetic isotope effects (KIEs), which

18

lead to characteristic changes of stable isotope ratios in the remaining contaminant. This so-called

19

stable isotope fractionation enables one to characterize contaminant transformation, especially

20

when isotope ratios of several elements are considered simultaneously. Based on a quantitative

21

understanding of KIEs, it is possible to derive the extent and rates of (bio)degradation from pollutant

22

stable isotope fractionation. 6–13 Such insights are invaluable for assessing degradation processes

23

that occur over timescales of years to decades.

24

During oxidative pollutant transformations, which are often enzyme-catalyzed reactions, 14 the

25

interpretation of contaminant isotope fractionation is challenging because of the complex kinetics

26

of enzymatic dioxygen activation. 15–17 Isotope fractionation can be masked if formation of reactive

27

oxygen species rather than bond cleavage reactions of the contaminant become rate-limiting. Such

28

phenomena may lead to erroneous interpretations of small isotope fractionation, 18–21 especially

29

when CSIA is restricted to one isotopic element (e.g., due to analytical constraints of continuous

30

flow isotope ratio mass spectrometry 22 ). Even though variations of isotope fractionation behaviour

31

are frequently addressed in CSIA studies, 23–29 there have been no attempts to interpret the latter

32

systematically based on the extensively-studied kinetics of oxidative biocatalysis.

33

Here, we show how the two major catalytic strategies of flavin-dependent monooxygenases

34

with regard to O2 activation can be used to rationalize the isotope fractionation associated with the

35

monooxygenation of organic micropollutants. Flavin-dependent monooxygenases (FMO) play a key

36

role in many biological processes 30–32 and are known to catalyze the initial steps of degradation of

37

many contaminants including chloro- and nitrophenol derivatives, 33–41 toluenes, 42 and xylenes. 43

38

FMOs have evolved two catalytic strategies to prevent wasteful NAD(P)H oxidation and therefore

39

differ regarding the sequence of substrate binding and O2 activation. 44 “Bold” FMOs generate

40

peroxides regardless of substrate availability and protect these peroxides from reacting until sub3 ACS Paragon Plus Environment

Environmental Science & Technology Letters

41

strate molecules are encountered. In contrast, the formation of the reactive flavin (hydro)peroxide

42

species responsible for hydroxylation “cautious” FMOs is only triggered in the presence of the sub-

43

strate. Therefore, we hypothesize that the isotope fractionation of organic substrates hydroxylated

44

by “bold” and “cautious” FMOs is fundamentally different. Rate of substrate disappearance would

45

be determined by its chemical reaction with “bold ” FMOs thus leading to measurable substrate

46

isotope fractionation. The transformation in “cautious” FMOs, in contrast, would be limited, for

47

example, by substrate binding, whose isotope effects are much smaller 45 and difficult to detect. A

48

priori knowledge of kinetic mechanisms of FMOs with organic pollutants as substrates could thus

49

enable one to delineate cases where CSIA may be suited to track biodegradation from cases where

50

it is not.

51

We tested the above hypothesis that monooxygenations by “bold” FMOs leads to measurable

52

substrate isotope fractionation whereas no isotope fractionation can be observed if the reaction is

53

catalyzed by “cautious” FMOs. To this end, we investigated the two known prototypes for “bold” and

54

“cautious” FMOs, namely the transformation of cyclohexanone by cyclohexanone monooxygenase

55

(EC 1.14.13.22, a Baeyer-Villiger monooxygenase) 46,47 and of 4-hydroxybenzoate (4-HBA) by 4-

56

hydroxybenzoate hydroxylase (EC 1.14.13.2). 48 Crude cell extracts containing 4-hydroxybenzoate

57

hydroxylase were used to rule out the masking of isotope fractionation by transport processes

58

across cell membranes. Insights from the work with the prototype-FMOs was applied to interpret

59

the catalytic strategy associated with the monooxygenation of 4-hydroxyphenylacetic acid (4-HPA)

60

by 4-hydroxyphenylacetate hydroxylase. While the mechanism of 4-HPA monooxygenation is

61

identical to the one for 4-HBA, 49 the 4-hydroxyphenylacetate hydroxylase belongs to the group of

62

two component flavin-dependent monooxygenases, 50 which are more widely related to contaminant

63

biodegradation 39,41 and supposedly react like “bold” Baeyer-Villiger monooxygenases. 49

4 ACS Paragon Plus Environment

Page 4 of 21

Page 5 of 21

Environmental Science & Technology Letters

64

Materials and Methods

65

Biodegradation and biotransformation experiments

66

We carried out three different types of experiments with (a) enzyme assays, (b) resting cells, and

67

(c) crude cell extracts to study the monooxygenation of cyclohexanone, 4-hydroxybenzoic acid (4-

68

HBA), and 4-hydroxyphenylacetate (4-HPA) as described in detail in the Supporting Information.

69

All experiments were performed in duplicates and aqueous samples, taken at predefined time-points

70

and acidified with 2 vol.% of HCl (7.5%) to stop substrate conversion, were stored at 4 ℃ until

71

concentration and isotopic analyses.

72

(a) Experiments in enzyme assays of a commercially available cyclohexanone monooxygenase

73

purified from Acinetobacter sp. were set up according to a modified procedure form Sheng et al. 46

74

to study the transformation of cyclohexanone . Assay solutions contained 5 ml 0.1 M glycine/NaOH

75

buffer (pH 9), 25 µM FAD, 3 mM NADPH and 50 µl cyclohexanone monooxygenase (≥12 U/mL).

76

The reaction was initiated with 1.5 mM cyclohexanone and subsequently incubated in a flask on a

77

rotary shaker at 25 ℃. NADPH oxidation was monitored at 340 nm with a spectrophotometer. Con-

78

trol experiments were carried out identically except for addition of cyclohexanone monooxygenase

79

or NADPH, respectively.

80

(b) Experiments with resting cells of Pseudomonas fluorescens Pf-5 were carried out in phos-

81

phate buffer (0.025 M, pH 7) and incubated on a rotary shaker at 28 ℃ with the substrate used in

82

the pre-grown culture, that is either 5 mM 4-HBA or 2 mM 4-HPA (see Supporting Information,

83

SI). Control experiments did not contain cells.

84

(c) Experiments with crude cell extract of Pseudomonas fluorescens Pf-5 were prepared to

85

study 4-HBA transformation. 25 Cells were suspended in phosphate buffer (0.02 M, pH 7.0) on

86

ice, followed by sonication (Sonics Vibra Cell VCX130) and centrifugation at 18000 rpm for 20

87

minutes. The supernatant was designated as crude cell extract. Reactions were performed on a

88

rotary shaker at 25 ℃ in 10 ml flasks containing 25 mM phosphate buffer (pH 7.0), 3 ml crude cell

89

extract, 40 µM FAD, and 1 mM NADPH. The reaction was initiated by adding 350 µM 4-HBA.

5 ACS Paragon Plus Environment

Environmental Science & Technology Letters

90

Control experiments were set up identically without NADPH.

91

Chemical, biological, and isotopic analyses

92

Protein concentrations were determined according to Lowry et al. 51 . Instrumental settings for

93

both chemical and isotopic analysis followed previously established procedures, 23,52,53 which are

94

described in the SI. Concentrations of 4-HBA, 4-HPA were quantified without further treatment

95

by reverse-phase HPLC with a UV-Vis detection. Ethyl acetate extracts of cyclohexanone were

96

measured by GC/MS. The natural abundance 13 C/12 C ratios of 4-HBA, 4-HPA, and cyclohexanone

97

were measured by high-temperature liquid chromatography coupled to isotope ratio mass spectrom-

98

etry. All C isotope ratios are expressed as δ13 C relative to Vienna PeeDee Belemnite (δ13 CVPDB )

99

using calibrated reference materials.

100

C isotope enrichment factors, ǫ C , were obtained from linear regression analysis of logarithms of

101

normalized δ13 C-values vs. fraction of the remaining reactant (c/c0 , eq. S1). Apparent 13 C-kinetic

102

isotope effects (13 C-AKIEs) for the reacting C bond(s) were calculated with eq. S2 by accounting

103

for the total number of C atoms and neglecting secondary isotope effects.

104

Results and Discussion

105

Cyclohexanone monooxygenase

106

The catalytic cycle of cyclohexanone monooxygenase is displayed in Scheme 1a and the δ13 C-values

107

of cyclohexanone (compound 1) during its transformation to ǫ-caprolactone (3) are shown in Figure

108

1a. We observed substantial C isotope fractionation in the remaining substrate corresponding to a

109

C isotope enrichment factor, ǫ C , of −8.7 ± 0.1h. The Baeyer-Villiger oxidation of cyclohexanone

110

to ǫ-caprolactone is well-studied and proceeds through a Criegee two-step mechanism, which

111

involves formation of a hemiperacetal intermediate (2). 54,55 We assumed that only the carbonyl-C

112

experiences a primary C isotope effect to calculate a large

13 C-AKIE

6 ACS Paragon Plus Environment

of 1.055 ± 0.007, which is

Page 6 of 21

Page 7 of 21

Environmental Science & Technology Letters

13 C-KIE

113

of similar magnitude as the

for the carbonyl position of cyclohexanone (1.051 ± 0.004)

114

during Baeyer-Villiger oxidation in dichloromethane by 13 C-NMR. 56 The good agreement of isotope

115

effects for the enzymatic and chemical Baeyer-Villiger oxidation of cyclohexanone suggests that

116

the oxidation of the substrate was the rate-limiting step of cyclohexanone disappearance in our

117

experiments.

118

As shown in Scheme 1a, the overall kinetic mechanism of cyclohexanone monooxygenase

119

involves the reduction of the oxidized enzyme, E-Flox , by NADPH followed by the rapid oxygenation

120

of the reduced flavin NADP+ -E-Flred to a (hydro)peroxide. While the hydroperoxide species is

121

unreactive, the peroxide, NADP+ -E-Fl–OO− , rapidly transforms 1 to 3. 46 Dehydration of the flavin

122

C4 -hydroxide (NADP+ -E-Fl–OH) as well as the release of 3 and NADP+ lead to the recovery of

123

the oxidized enzyme, E-Flox . Spectroscopic analyses by Sheng et al. 46 revealed that the reaction

124

of the Criegee adduct (2 → 3) and dehydration of NADP+ -E-Fl–OH are faster than the first step

125

of substrate transformation (1 → 2). The authors also ruled out that the release 3 and NADP+

126

limited the rate of catalytic turnover for cyclohexanone as substrate. The current knowledge of the

127

catalytic cycle of cyclohexanone monooxagenase thus strongly supports the above interpretation of

128

the 13 C-kinetic isotope effects. As hypothesized for a “bold” FMO, our data indicate that the initial

129

oxidation of cyclohexanone is rate-determining and, consequently, responsible for the observed

130

substrate isotope fractionation.

131

4-Hydroxybenzoate hydroxylase

132

Resting cells of Pseudomonas fluorescens Pf-5 transformed 4-hydroxybenzoate (4-HBA) within 7.5

133

h following zero-order kinetics and we did not observe significant substrate C isotope fractionation

134

(Figure 1b). The ǫ C -value was 0.03 ± 0.05h. The corresponding 13 C-AKIE was 0.9997 ± 0.0004,

135

that is not different from unity. 4-HBA transformation in experiments with crude cell extracts of

136

Pseudomonas fluorescens Pf-5 was approximately three times faster (Table S1) but the ǫ C -value

137

in this experiment increased only slightly to −0.2 ± 0.1h. The fact that C isotope enrichment

138

in experiments with crude cell extracts is almost negligible suggests that the uptake of 4-HBA 7 ACS Paragon Plus Environment

Environmental Science & Technology Letters

139

did not mask the isotope effect of monooxygenation, in agreement with previously findings for

140

nitrophenols. 25 Changes of C hybridization upon (enzymatic) hydroxylation of aromatic moieties

141

such as those hypothesized for 4-HBA (Scheme 1b) 44,48,49 typically lead to larger C isotope

142

fractionation.

143

toluene, and nitrophenols. 25,28,29 These numbers imply that the C isotope fractionation during

144

monooxygenation by 4-hydroxybenzoate hydroxylase is at least tenfold smaller than expected from

145

bonding changes of 4-HBA and that the oxidation of the substrate was not the rate-limiting step of

146

4-HBA disappearance.

13 C-AKIE between 1.008 and 1.012 were reported for monooxygenation of benzene,

147

The kinetic mechanism of 4-hydroxybenzoate hydroxylase shown in Scheme 1b indeed features

148

important differences to that of cyclohexane monooxygenase. The oxidized enzyme E–Flox ex-

149

hibits a high affinity for 4-HBA (4 in Scheme 1b) and rapidly binds the substrate. Deprotonation of

150

4-HBA in the active site triggers a first conformational rearrangement of the enzyme, which limits

151

the overall rate of the enzymatic reaction. This step is followed by rapid reduction of the flavin

152

by NADPH. 44,48,49 Because the association rate constant of E-Flox for 4-HBA is several orders of

153

magnitude larger than for E-Flred , 44 the substrate is kinetically trapped while E-Flred reduces O2

154

to form the oxidant, flavin hydroperoxide (E-Fl–OOH). 4-HBA reacts in an electrophilic aromatic

155

substitution to compound 5, which tautomerises rapidly to 3,4-dihydroxybenzoate (6). The return

156

to the initial state (E-Flox ) requires further conformational change, in which E-Fl–OH is protonated

157

to eliminate H2 O and release 6. High substrate levels trap E-Fl–OH in the 4-hydroxybenzoate

158

hydroxylase and may limit the rate of enzymatic activity in addition to conformational rearrange-

159

ments of 4-hydroxybenzoate hydroxylase upon 4-HBA binding. The absence of substrate isotope

160

fractionation, as shown for 4-HBA in Figure 1b, is thus a consequence of the kinetic mechanisms

161

of a “cautious” FMO.

162

4-Hydroxyphenylacetate hydroxylase

163

Hydroxylation of 4-hydroxyphenylacetate (4-HPA, 7, Scheme 1c) by resting cells of Pseudomonas

164

fluorescens Pf-5 was associated with measurable C isotope fractionation (ǫ C = −1.1±0.1h, Figure 8 ACS Paragon Plus Environment

Page 8 of 21

Page 9 of 21

Environmental Science & Technology Letters

165

1c). The 13 C-AKIE of 1.0088 ± 0.0007 agrees well with the range of isotope effects reported for

166

aromatic hydroxylations obtained with resting cells and crude cell extracts as discussed above. The

167

magnitude of

168

ylase, where the rate of 4-HPA removal is determined by its reaction with a flavin hydroperoxide

169

(EO -Fl–OOH, Scheme 1c). Such two-component FMOs differ from the one-component FMOs in

170

that reduced flavins are formed in a separate, NADH-dependent reductase (ER ) as co-substrates and

171

need to be transferred to the monooxygenase (EO ) as opposed to being bound as prosthetic group to

172

the oxygenase. 44,49 This catalytic strategy implies that the activity of the reductase is not coupled

173

to substrate binding like in the 4-hydroxybenzoate hydroxylase. Instead, the flavin hydroperoxide

174

is formed regardless of the presence of 4-HPA after binding of the flavin to ER (Scheme 1c). This

175

behaviour resembles the kinetic mechanism of the “bold” cyclohexane monooxygenase (Scheme

176

1a).

177

Implications for the assessment of pollutant biotransformation

178

Both metabolic and co-metabolic monooxygenations are frequently the initial step of organic

179

pollutant transformation 14 and therefore typical degradation processes to be assessed by CSIA. 1–5,22

180

FMOs catalyze such oxidations at aromatic and (cyclic) ketone moieties, for example in chloro-

181

and nitrophenols as well as alkylated benzenes. A selection of bacterial strains that carry out

182

these reactions is listed in Table 1. The kinetic mechanisms of FMOs discussed above may

183

determine the observable isotope fractionation of many organic contaminants. We have recently

184

observed that monooxygenation of 4-nitrophenol by three microorganisms, in which FMOs have

185

been identified, 39,40,58 is not necessarily associated with substrate isotope fractionation. 25 Only

186

in two of three cases did the

187

atom hybridization upon hydroxylation. Evidence presented here suggests that isotope fractionation

188

during 4-nitrophenol monooxygenation was observable if the FMOs followed the kinetic mechanism

189

of “bold” FMOs. Conversely, the absence of isotope fractionation in 4-nitrophenol, as observed in

190

experiments with Pseudomonas sp. 1A, may have been due to oxidative catalysis by a “cautious”

13 C-AKIE

is consistent with the catalytic cycle of 4-hydroxyphenylacetate hydrox-

13 C-AKIEs

correspond to values that are typical for changes of C

9 ACS Paragon Plus Environment

Environmental Science & Technology Letters

191

FMO. These examples show that knowledge of the kinetic mechanisms of oxidative enzymes,

192

especially with regard to the sequence of oxygen activation and substrate binding, is key for assessing

193

contaminant transformation by CSIA. While our study illustrates these points for FMOs, similar

194

kinetic mechanisms may also be encountered in oxidative reactions of copper- or iron-dependent

195

enzymes. 15 Increasing our understanding of the enzymes involved in contaminant hydroxylation

196

may be crucial for future applications of CSIA.

197

Acknowledgement

198

We thank Dr. Andreas Essig for providing the two Pseudomonas fluorescens strains.

199

Supporting Information Available

200

Description of growth conditions, analytical methods, and data evaluation procedures. Compilation

201

of rates, ǫ C -, and

202

http://pubs.acs.org/.

203

Notes

204

The authors declare no competing financial interest.

205

References

13 C-AKIE-values.

This material is available free of charge via the Internet at

206

(1) Elsner, M.; Jochman, M. A.; Hofstetter, T. B.; Hunkeler, D.; Bernstein, A.; Schmidt, T. C.;

207

Schmimmelmann, A. Current challenges in compound-specific stable isotope analysis of

208

environmental organic contaminants. Anal. Bioanal. Chem. 2012, 403, 2471–2491.

209

(2) Thullner, M.; Centler, F.; Richnow, H.-H.; Fischer, A. Quantification of organic pollutant

210

degradation in contaminated aquifers using compound specific stable isotope analysis - Review

211

of recent developments. Org. Geochem. 2012, 42, 1440–1460. 10 ACS Paragon Plus Environment

Page 10 of 21

Page 11 of 21

Environmental Science & Technology Letters

212

(3) Schmidt, T. C.; Jochmann, M. A. Origin and fate of organic compounds in water: Char-

213

acterization by compound-specific stable isotope analysis. Annu. Rev. Anal. Chem. 2012, 5,

214

133–155.

215

216

(4) Hofstetter, T. B.; Berg, M. Assessing transformation processes of organic contaminants by compound-specific stable isotope analyses. TrAC-Trends Anal. Chem. 2011, 30, 618–627.

217

(5) Hofstetter, T. B.; Bolotin, J.; Pati, S. G.; Skarpeli-Liati, M.; Spahr, S.; Wijker, R. S. Isotope

218

effects as new proxies for organic pollutant transformation. CHIMIA 2014, 68, 788–792.

219

(6) Aeppli, C.; Hofstetter, T. B.; Amaral, H. I. F.; Kipfer, R.; Schwarzenbach, R. P.; Berg, M. Quan-

220

tifying in-situ transformation rates of chlorinated ethenes by combining compound-specific

221

stable isotope analysis, groundwater dating, and carbon isotope mass balances. Environ. Sci.

222

Technol. 2010, 44, 3705–3711.

223

(7) Wijker, R. S.; Bolotin, J.; Nishino, S. F.; Spain, J. C.; Hofstetter, T. B. Using compound-

224

specific isotope analysis to assess biodegradation of nitroaromatic explosives in the subsurface.

225

Environ. Sci. Technol. 2013, 47, 6872–6883.

226

(8) Revesz, K. M.; Sherwood Lollar, B.; Kirshtein, J. D.; Tiedeman, C. R.; Imbrigiotta, T. E.;

227

Goode, D. J.; Shapiro, A. M.; Voytek, M. A.; Lacombe, P. J.; Busenberg, E. Integration of

228

stable carbon isotope, microbial community, dissolved hydrogen gas, and 2 HH2 O tracer data

229

to assess bioaugmentation for chlorinated ethene degradation in fractured rocks. J. Contam.

230

Hydrol. 2014, 156, 62–77.

231

(9) McKelvie, J. R.; MacKay, D. M.; de Sieyes, N. R.; Lacrampe-Couloume, G.; Sherwood Lol-

232

lar, B. Quantifying MTBE biodegradation in the Vandenberg Air Force Base ethanol release

233

study using stable carbon isotopes. J. Contam. Hydrol. 2007, 94, 157–165.

234

(10) Ponsin, V.; Maier, J.; Guelorget, Y.; Hunkeler, D.; Bouchard, D.; Villavicencio, H.; Höhener, P.

235

Documentation of time-scales for onset of natural attenuation in an aquifer treated by a crude-

236

oil recovery system. Sci. Total Environ. 2015, 512, 62–73. 11 ACS Paragon Plus Environment

Environmental Science & Technology Letters

237

(11) Damgaard, I.; Bjerg, P. L.; Bælum, J.; Scheutz, C.; Hunkeler, D.; Jacobsen, C. S.; Tuxen, N.;

238

Broholm, M. M. Identification of chlorinated solvents degradation zones in clay till by high

239

resolution chemical, microbial and compound specific isotope analysis. J. Contam. Hydrol.

240

2013, 146, 37–50.

241

(12) Zwank, L.; Berg, M.; Elsner, M.; Schmidt, T. C.; Schwarzenbach, R. S.; Haderlein, S. B. New

242

evaluation scheme for two-dimensional isotope analysis to decipher biodegradation processes:

243

Application to groundwater contamination by MTBE. Environ. Sci. Technol. 2005, 39, 1018–

244

1029.

245

(13) Nijenhuis, I.; Schmidt, M.; Pellegatti, E.; Paramatti, E.; Richnow, H.-H.; Gargini, A. A

246

stable isotope approach for source apportionment of chlorinated ethene plumes at a complex

247

multi-contamination events urban site. J. Contam. Hydrol. 2013, 153, 92–105.

248

249

250

251

252

253

(14) Schwarzenbach, R. P.; Gschwend, P. M.; Imboden, D. M. Environmental Organic Chemistry, 2nd ed.; 1311 pp; John Wiley & Sons: Hoboken, NJ, 2003; p 1311. (15) Bertini, I., Gray, H. B., Stiefel, E. I., Valentine, J. S., Eds. Biological Inorganic Chemistry; University Science Books: Sausalito, CA, 2007. (16) Bugg, T. Introduction to Enzyme and Coenzyme Chemistry, 3rd ed.; Blackwell Publishing Ltd.: Chichester, UK, 2012.

254

(17) Selverstone Valentine, J., Foote, C. S., Greenberg, A., Liebman, J. F., Eds. Active Oxygen in

255

Biochemistry; Springer-Science+Business Media: Dordrecht, 1995; Vol. SEARCH Series,

256

Volume 3.

257

258

259

260

(18) Cleland, W. W. The use of isotope effects to determine enzyme mechanisms. Arch. Biochem. Biophys. 2005, 433, 2–12. (19) Northrop, D. B. The expression of isotope effects on enzyme-catalyzed reactions. Annu. Rev. Biochem. 1981, 50, 103–131. 12 ACS Paragon Plus Environment

Page 12 of 21

Page 13 of 21

261

262

263

264

265

266

Environmental Science & Technology Letters

(20) Cleland, W. W. Use of isotope effects to determine enzyme mechanisms. J. Label. Compd. Radiopharm. 2007, 50, 1006–1015. (21) Duggleby, R. G.; Northrop, D. B. The expression of kinetic isotope effects during the time course of enzyme-catalyzed reactions. Bioorganic Chemistry 1989, 17, 177–193. (22) Jochmann, M. A.; Schmidt, T. C. Compound-specific Stable Isotope Analysis; The Royal Society of Chemistry, 2012.

267

(23) Pati, S. G.; Kohler, H. P. E.; Bolotin, J.; Parales, R. E.; Hofstetter, T. B. Isotope effects of

268

enzymatic dioxygenation of nitrobenzene and 2-nitrotoluene by nitrobenzene dioxygenase.

269

Environ. Sci. Technol. 2014, 48, 10750–10759.

270

(24) Wijker, R. S.; Adamczyk, P.; Bolotin, J.; Paneth, P.; Hofstetter, T. B. Isotopic analysis of

271

oxidative pollutant degradation pathways exhibiting large H isotope fractionation. Environ.

272

Sci. Technol. 2013, 47, 13459–13468.

273

(25) Wijker, R. S.; Kurt, Z.; Spain, J. C.; Bolotin, J.; Zeyer, J.; Hofstetter, T. B. Isotope fractionation

274

associated with the biodegradation of 2- and 4-nitrophenols via monooxygenation pathways.

275

Environ. Sci. Technol. 2013, 47, 14185–14193.

276

(26) Dorer, C.; Vogt, C.; Kleinsteuber, S.; Stams, A. J. M.; Richnow, H.-H. Compound-specific

277

isotope analysis as a tool to characterize biodegradation of ethylbenzene. Environ. Sci. Technol.

278

2014, 48, 9122–9132.

279

(27) Pati, S. G.; Shin, K.; Skarpeli-Liati, M.; Bolotin, J.; Eustis, S. N.; Spain, J. C.; Hofstet-

280

ter, T. B. Carbon and nitrogen isotope effects associated with the dioxygenation of aniline and

281

diphenylamine. Environ. Sci. Technol. 2012, 46, 11844–11853.

282

(28) Vogt, G.; Cyrus, E.; Herklotz, I.; Schlosser, D.; Bahr, A.; Herrmann, S.; Richnow, H.-H.;

283

Fischer, A. Evaluation of toluene degradation pathways by two-dimensional stable isotope

284

fractionation. Environ. Sci. Technol. 2008, 42, 7793–7800. 13 ACS Paragon Plus Environment

Environmental Science & Technology Letters

285

(29) Fischer, A.; Herklotz, I.; Herrmann, S.; Thullner, M.; Weelink, S. A. B.; Stams, A. J. M.;

286

Schlomann, M.; Richnow, H. H.; Vogt, C. Combined carbon and hydrogen isotope fraction-

287

ation investigations for elucidating benzene biodegradation pathways. Environ. Sci. Technol.

288

2008, 42, 4356–4363.

289

290

291

292

(30) van Berkel, W. J. H.; Kamerbeek, N. M.; Fraaije, M. W. Flavoprotein monooxygenases, a diverse class of oxidative biocatalysts. J. Biotechnol. 2006, 124, 670–689. (31) Huijbers, M. M. E.; Montersino, S.; Westphal, A. H.; Tischler, D.; van Berkel, W. J. H. Flavin dependent monooxygenases. Arch. Biochem. Biophys. 2014, 544, 2–17.

293

(32) Joosten, V.; van Berkel, W. J. H. Flavoenzymes. Curr. Opin. Chem. Biol. 2007, 11, 195–202.

294

(33) Gisi, M. R.; Xun, L. Y. Characterization of chlorophenol 4-monooxygenase (TftD) and NADH:

295

Flavin adenine dinucleotide oxidoreductase (TftC) of Burkholderia cepacia AC1100. J. Bac-

296

teriol. 2003, 185, 2786–2792.

297

(34) Wieser, M.; Wagner, B.; Eberspacher, J.; Lingens, F. Purification and characterization of 2,4,6-

298

trichlorophenol-4-monooxygenase, a dehalogenating enzyme from Azotobacter sp strain GP1.

299

J. Bacteriol. 1997, 179, 202–208.

300

(35) Belchik, S. M.; Xun, L. Y. Functions of flavin reductase and quinone reductase in 2,4,6-

301

trichlorophenol degradation by Cupriavidus necator JMP134. J. Bacteriol. 2008, 190, 1615–

302

1619.

303

(36) Louie, T. M.; Webster, C. M.; Xun, L. Y. Genetic and biochemical characterization of a 2,4,6-

304

trichlorophenol degradation pathway in Ralstonia eutropha JMP134. J. Bacteriol. 2002, 184,

305

3492–3500.

306

(37) Xun, L. Y.; Orser, C. S. Purification and Properties of Pentachlorophenol Hydroxylase, A

307

Flavoprotein From Flavobacterium sp. Strain Atcc 39723. J. Bacteriol. 1991, 173, 4447–

308

4453. 14 ACS Paragon Plus Environment

Page 14 of 21

Page 15 of 21

Environmental Science & Technology Letters

309

(38) Wang, H.; Tiirola, M. A.; Puhakka, J. A.; Kulomaa, M. S. Production and characterization

310

of the recombinant Sphingomonas chlorophenolica pentachlorophenol 4-monooxygenase.

311

Biochem. Biophys. Res. Commun. 2001, 289, 161–166.

312

(39) Kadiyala, V.; Spain, J. C. A two-component monooxygenase catalyzes both the hydroxylation

313

of p-nitrophenol and the oxidative release of nitrite from 4-nitrocatechol in Bacillus sphaericus

314

JS905. Appl. Environ. Microbiol. 1998, 64, 2479–2484.

315

316

(40) Spain, J. C.; Gibson, D. Pathway for biodegradation of para-nitrophenol in a Moraxella sp. Appl. Environ. Microbiol. 1991, 57, 812–819.

317

(41) Takeo, M.; Yasukawa, T.; Abe, Y.; Niihara, S.; Maeda, Y.; Negoro, S. Cloning and charac-

318

terization of a 4-nitrophenol hydroxylase gene cluster from Rhodococcus sp PN1. J. Biosci.

319

Bioeng. 2003, 95, 139–145.

320

321

(42) Newman, L. M.; Wackett, L. P. Purification and characterization of toluene 2-monooxygenase from Burkholderia cepacia G4. Biochemistry 1995, 34, 14066–14076.

322

(43) Cafaro, V.; Scognamiglio, R.; Viggiani, A.; Izzo, V.; Passaro, I.; Notomista, E.; Dal Piaz, F.;

323

Amoresano, A.; Casbarra, A.; Pucci, P.; Di Donato, A. Expression and purification of the

324

recombinant subunits of toluene/o-xylene monooxygenase and reconstitution of the active

325

complex. Eur. J. Biochem. 2002, 269, 5689–5699.

326

327

(44) Palfey, B. A.; McDonald, C. A. Control of catalysis in flavin-dependent monooxygenases. Arch. Biochem. Biophys. 2010, 493, 26–36.

328

(45) Swiderek, K.; Paneth, P. Binding isotope effects. Chem. Rev. 2013, 113, 7851–7879.

329

(46) Sheng, D. W.; Ballou, D. P.; Massey, V. Mechanistic studies of cyclohexanone monooxygenase:

330

Chemical properties of intermediates involved in catalysis. Biochemistry 2001, 40, 11156–

331

11167.

15 ACS Paragon Plus Environment

Environmental Science & Technology Letters

332

333

334

335

(47) Pazmino, D. E. T.; Dudek, H. M.; Fraaije, M. W. Baeyer-Villiger monooxygenases: recent advances and future challenges. Curr. Opin. Chem. Biol. 2010, 14, 138–144. (48) Entsch, B.; Cole, L. J.; Ballou, D. P. Protein dynamics and electrostatics in the function of p-hydroxybenzoate hydroxylase. Arch. Biochem. Biophys. 2005, 433, 297–311.

336

(49) Ballou, D. P.; Entsch, B.; Cole, L. J. Dynamics involved in catalysis by single-component and

337

two-component flavin-dependent aromatic hydroxylases. Biochem. Biophys. Res. Commun.

338

2005, 338, 590–598.

339

(50) Jimenez, J. I.; Minambres, B.; Garcia, J. L.; Diaz, E. In Pseudomonas; Ramos, J. L., Ed.;

340

Genomic insights in the metabolism of aromatic compounds in Pseudomonas; Springer: New

341

York, 2004; Vol. 3 Biosynthesis of Macromolecules and Molecular Metabolism; pp 425–462.

342

(51) Lowry, O. H.; Rosebrough, N. J.; Farr, A. L.; Randall, R. J. Protein measurement with the

343

folin phenol reagent. J. Biol. Chem. 1951, 193, 265–275.

344

(52) Berg, M.; Bolotin, J.; Hofstetter, T. B. Compound-specific nitrogen and carbon isotope analysis

345

of nitroaromatic compounds in aqueous samples using solid-phase microextraction coupled

346

to GC/IRMS. Anal. Chem. 2007, 79, 2386–2393.

347

(53) Skarpeli-Liati, M.; Arnold, W. A.; Turgeon, A.; Cramer, C. J.; Hofstetter, T. B. pH-Dependent

348

equilibrium isotope fractionation associated with compound-specific nitrogen and carbon

349

isotope analysis by SPME-GC/IRMS. Anal. Chem. 2011, 83, 1641–1648.

350

351

352

353

(54) Smith, M. B.; March, J. March’s Advanced Organic Chemistry, 6th ed.; John Wiley & Sons Inc.: New York, NY, 2007. (55) Clayden, J.; Greeves, N.; Warren, S.; Wothers, P. Organic Chemistry; Oxford University Press: Oxford, 2001.

16 ACS Paragon Plus Environment

Page 16 of 21

Page 17 of 21

Environmental Science & Technology Letters

354

(56) Singleton, D. A.; Szymanski, M. J. Simultaneous determination of intermolecular and in-

355

tramolecular 13 C and 2 H kinetic isotope effects at natural abundance. J. Am. Chem. Soc. 1999,

356

121, 9455–9456.

357

(57) Chakraborty, S.; Ortiz-Maldonado, M.; Entsch, B.; Ballou, D. P. Studies on the mechanism of

358

p-hydroxyphenylacetate 3-hydroxylase from Pseudomonas aeruginosa: A system composed

359

of a small flavin reductase and a large flavin-dependent oxygenase. Biochemistry 2010, 49,

360

372–385.

361

(58) Perry, L. L.; Zylstra, G. J. Cloning of a gene cluster involved in the catabolism of p-nitrophenol

362

by Arthrobacter sp strain JS443 and characterization of the p-nitrophenol monooxygenase. J.

363

Bacteriol. 2007, 189, 7563–7572.

364

365

(59) Kotani, T.; Yurimoto, H.; Kato, N.; Sakai, Y. Novel acetone metabolism in a propane-utilizing bacterium, Gordonia sp strain TY-5. J. Bacteriol. 2007, 189, 886–893.

366

(60) Rehdorf, J.; Kirschner, A.; Bornscheuer, U. T. Cloning, expression and characterization of a

367

Baeyer-Villiger monooxygenase from Pseudomonas putida KT2440. Biotechnol. Lett. 2007,

368

29, 1393–1398.

369

(61) Iwaki, H.; Wang, S. Z.; Grosse, S.; Bergeron, H.; Nagahashi, A.; Lertvorachon, J.; Yang, J. Z.;

370

Konishi, Y.; Hasegawa, Y.; Lau, P. C. K. Pseudomonad cyclopentadecanone monooxygenase

371

displaying an uncommon spectrum of Baeyer-Villiger oxidations of cyclic ketones. Appl.

372

Environ. Microbiol. 2006, 72, 2707–2720.

17 ACS Paragon Plus Environment

Environmental Science & Technology Letters

Page 18 of 21

373

Table 1. Examples of oxygenations and substrates degraded by microorganisms that exhibit flavindependent monooxygenases.

Substrates

Strain

Reference

Hydroxylation 4-Nitrophenol

Trichlorophenol (TCP)

Pseudomonas sp. 1A Bacillus spharericus JS905 Arthrobacter sp. JS443 Burkholderia cepacia AC1100 Azetobacter sp strain GP1 Cupriavidus necator JMP134 Ralstonia eutropha JMP134

40 39,41 58 33 34 35 36

Flavobacterium sp. Strain ATCC 39723 Sphingomonas chlorophenolica

37

Burkholderia cepacia G4 Pseudomonas stutzeri OX1

42

Aceton

Gordonia sp. TY-5

59

(Cyclic) ketones

Pseudomonas putida KT2440 Pseudomonas sp. HI-70

60

Pentachlorophenol (PCP) Toluene and o-xylene

38

43

Baeyer-Villiger oxidation

18 ACS Paragon Plus Environment

61

Page 19 of 21

Environmental Science & Technology Letters

(a) Cyclohexanone monooxygenase ("bold" FMO) NADPH E-Flox

NADPH

NADP

E-Flox

E-Flred

O2

NADP E-Fl-OO –

3 H 2O NADP

1

O

NADP

NADP

NADP

E-Fl

E-Fl

E-Fl

O

HO

H+

O O

O

O

O

O

3

2

1

(b) 4-Hydroxybenzoate hydroxylase ("cautious" FMO) 4

NADPH

E-Flox

NADP NADPH

NADP

E-Flox

E-Flox

E-Flred

E-Flred

4

4

4

4

6 H 2O

O2 E-Fl

E-Fl

E-Fl

O

O

HO OH

OH

H

O

O OH

O

O

OH O

O

OH O

HB

6

B 4

5

(c) 4-Hydroxyphenylacetate hydroxylase NAD ER-Flred NAD Fl red

NADH ER

ER-Flox

O2

EO

E O-Fl-OOH

E O-Flred

NADH + Fl ox 7

9

E O-Fl

OH O O

9

OH

H OH

O

O

O

OH

O

E O-Fl

E O-Fl

OH

O

HB 8

OH O

B O

7

Scheme 1. Catalytic cycles of FMOs studied in this work (modified from Palfey and McDonald 44 , Sheng et al. 46 , Ballou et al. 49 ). (a) Cyclohexanone monooxygenase, the prototype for “bold” FMOs. E stands for the enzyme, Flox and Flred are the oxidized and reduced flavin prosthetic groups, respectively. E-Fl–OO− is the flavin peroxide. Cyclohexanone (1) is oxidized via a Criege intermediate (2) to ǫ -caprolactone (3). (b) 4-Hydroxybenzoate hydroxylase, the prototype for “cautious” monooxygenases. 44,48 The oxidized flavin prosthetic group (Flox ) binds the substrate 4-HBA (4) prior to reduction of the 4-HBA-enzyme complex by NADPH. Compound 6 is 3,4-dihydroxybenzoate, B and BH+ stand for a neutral and protonated base. (c) The 4-hydroxyphenylacetate hydroxylase is a two component FMO 49,57 where ER and EO are the reductase and oxidase, respectively. Compound 7 is 4-HPA, 9 is 3,4-dihydroxyphenylacetate.

19 ACS Paragon Plus Environment

Environmental Science & Technology Letters

0 (a) Cyclohexanone

-5

-1

k0 = 240±11 µM min (g protein)

13

! C (‰)

-10 -15

13

-1

C-AKIE = 1.055±0.007 "C = -8.7±1.0

-20 -25 -30 1.0 0 -5

13

! C (‰)

-10

0.8

0.6

0.4

0.2

0.0

(b) 4-hydroxybenzoate -1

k0 = 31±1 µM min (g protein) "C = 0.07±0.05

-1

-15 -20 -25 -30 1.0 0 -5

0.8

0.6

0.4

0.2

(c) 4-hydroxyphenylacetate -1

k0 = 11±1 µM min (g protein)

13

! C (‰)

-10 -15

0.0

13

-1

C-AKIE = 1.0088±0.0007 "C = -1.1±0.1

-20 -25 -30 1.0

0.8

0.6

0.4

0.2

0.0

c/c0

Figure 1. Substrate C isotope fractionation measured in the three experimental systems vs. fraction of remaining substrate c/c0 . k0 stands for the zero-order reaction rate constant, 13 C-AKIE is the apparent 13 C-kinetic isotope effect, and ǫ C is the substrate C isotope enrichment factor. (a) Cyclohexanone transformation by cyclohexanone monooxygenase in enzyme assays, (b) Transformation of 4-HBA by 4-hydroxybenzoate hydroxylase in resting cells of Pseudomonas fluorescens Pf5, (c) Transformation of 4-HPA by 4-hydroxyphenylacetate hydroxylase in resting cells of Pseudomonas fluorescens Pf5. Lines were calculated with eq. S1. Light gray symbols and dashed lines refer to data from control experiments that did not contain enzymes or cells. Note that error bars are smaller than marker size.

20 ACS Paragon Plus Environment

Page 20 of 21

Page 21 of 21

Environmental Science & Technology Letters

ACS Paragon Plus Environment