Aerobic Utilization of Hydrocarbons, Oils and Lipids 9783319397825

Citation preview

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization: An Introduction F. Widdel and F. Musat

Abstract

Hydrocarbons represent “energy-rich” growth substrates for aerobic microorganisms and in principle allow high growth yields. In contrast, the energy gain with hydrocarbons in many anaerobic microorganisms is very low. The maximum gain of energy per mol of hydrocarbon degraded in the catabolism is predicted from calculated ΔG values. Some anaerobic degradation reactions of hydrocarbons with very low-energy gain as well as anaerobic activation reactions of hydrocarbons deserve particular attention from a bioenergetic point of view.

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Some Basic Thermodynamic Aspects of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Energetics of Hydrocarbon Utilization by Microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Catabolic Net Reactions of Hydrocarbons from the Energetic Perspective . . . . . . . . . . . 3.2 Hydrocarbon Activation from the Energetic Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Quantitative Aspects of Cell Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 ATP and Growth Yields . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Requirement for Minerals (N, P, Fe) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Appendix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 2 8 8 13 18 18 25 27 28 39

F. Widdel (*) Max Planck Institute for Marine Microbiology, Bremen, Germany e-mail: [email protected] F. Musat UFZ-Helmholtz Centre for Environmental Research, D-04318 Leipzing, Germany e-mail: [email protected] # Springer International Publishing AG 2016 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-319-39782-5_2-1

1

2

1

F. Widdel and F. Musat

Introduction

The study of microbial growth with hydrocarbons and their degradation often gets into energetic aspects, even though at a glance the metabolism of hydrocarbons is not basically different from that of other organic compounds. The overall metabolism in a chemotrophic organism follows the universal bifurcate carbon flow: One part of the carbon substrate together with sources of other elements (N, P, S, Fe, etc.) is used for synthesis of cell components, a process referred to as anabolism (synthetic metabolism, assimilation). The anabolic “upgrading” of the substrate requires and dissipates much energy, which is usually provided in the form of ATP and derived from another part of the carbon substrate. This part of the substrate necessarily undergoes degradation; the degradative substrate flow is referred to as catabolism (energy metabolism, dissimilation). Still, there are some energetic peculiarities in the metabolism of hydrocarbons which deserve attention. (1) First, even though flammability of hydrocarbons at the air implies “energy richness,” they are not energy rich under all circumstances. In the absence of oxygen, hydrocarbons are less energy rich than for instance the less flammable glucose. Whereas the latter provides energy for various modes of fermentative growth, fermentation of saturated, aromatic, and many other unsaturated nonaromatic hydrocarbons is energetically not feasible1; this is one reason why they tend to be preserved in deep reservoirs. (2) Second, hydrocarbons are chemically unreactive at room temperature. Their use in the metabolism has to begin with an activation reaction, the introduction of a functional group, which may require and “waste” energy from the overall energy budget of the microorganism. Also energies of transition states in the activation reactions have been of interest for a mechanistic understanding. (3) Third, for the theoretical treatment of energy conservation with hydrocarbons as well as for the estimation of microbial cell mass involved in hydrocarbon (petroleum) bioremediation, growth yields (cell mass produced per amount of hydrocarbon utilized) are of interest. This chapter briefly addresses some of these energetic peculiarities and quantitative aspects of hydrocarbon metabolism (Fig. 1).

2

Some Basic Thermodynamic Aspects of Hydrocarbons

Hydrocarbons, the main constituents of oil and gas, are the major source of energy in our industrialized society. A prominent property of hydrocarbons is thus their “energy richness.” More precisely, this term expresses that energy is released if they are oxidized with oxygen and that the amount of energy released per unit mass (the gravimetric energy density) of a liquid or solid hydrocarbon is higher than that from the oxidation of many other chemical compounds or elements (Appendix Table 5). In the case of gaseous hydrocarbons, a high volumetric energy density is 1

A fermentable hydrocarbon is, for instance, the unsaturated acetylene. Also some other unsaturated hydrocarbons are, at least theoretically, fermentable.

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

3

lism

Cell synthesis

abo

NH4+

An

HPO2– 4

Nonproductive

FeII, FeIII Hydrocarbon

Activated product

Energy

Energy

m

olis

tab

Ca

Aox

Aox

Oxidized form of electron acceptor

Ared Reduced form of electron acceptor

Ared

CO2

Energy dissipation

Fig. 1 The metabolism of hydrocarbons in chemotrophic microorganisms follows the universal bifurcate substrate flow into cell synthesis and degradation. A peculiarity in comparison to the metabolism of most non-hydrocarbon substrates is the activation which may require and dissipate energy

obvious if compared to that of other gases (Appendix Table 5). This “energy richness” is due to the high affinity of the two constituents, hydrogen and carbon, for oxygen and to the absence of oxidized carbon groups (such as C
OH or C ¼ O groups). The low atomic masses of hydrogen and carbon2 is another factor that contributes to the high gravimetric energy density. It is the high gravimetric energy density which, together with the abundance of hydrocarbons in the form of petroleum, has made them ideal fuels for vehicles and aircrafts. Another technical advantage is the formation of volatile products (CO2, H2O vapor). Feasibility and maximum energy gains of formulated stoichiometric reactions are expressed by their free energy changes, the ΔG -values. If a reaction is feasible under the given conditions (exergonic reaction), the ΔG-value is negative by convention. A positive value necessarily indicates that the reaction can in principle not occur under the given conditions (endergonic reaction), and a value of zero indicates that reactants and products are in equilibrium. Most reactions in chemistry and biology are associated with liberation of heat to the surroundings (exothermic reactions),

2

H, 1.008; C, 12.011.

4

F. Widdel and F. Musat

which is expressed by their heat or enthalpy3 changes (ΔH-values). Some reactions consume heat from the surroundings (endothermic reactions), and a few of such type also occur in microorganisms. Free energy or enthalpy changes are calculated from   free energies of formation (ΔGf , sometimes also termed Gf ) or enthalpies of   formation (ΔHf , sometimes also termed Hf ), respectively, which are given for standard conditions4 and for which there is a broad data basis. Appendix Table 6 compiles the values under standard conditions for several hydrocarbons and a number of other compounds which often appear in catabolic reactions. For a reaction a Aþb B!c Cþd D

(1)

(with a, b, c, d being the stoichiometric factors), the standard free energy change (viz., for all compounds at standard conditions) is the difference

      ΔG ¼ c ΔGf C þ d ΔGf D
a ΔGf A þ b ΔGf B

(2)

Calculation of the free energy change ΔG for nonstandard activities (a, in case of gases termed fugacity; a must not be confused with the stoichiometric factor a) considers the “nonchemical” energy change associated with dilution or concentration (“volume work”) of each component. These are logarithmic functions involving the gas constant and absolute temperature, the sum of which modifies the free energy change for standard activities, ΔGStandard, according to ΔG ¼ ΔGStandard þ R T ln

acC  adD aaA  abB

(3)

T in this equation must be the temperature for which the underlying ΔGStandard value has been given (viz., usually 298.15 K), and ΔG values at other temperatures cannot be calculated by this equation.5 The activities (effective concentrations) of solutes, a, can be usually substituted with acceptable precision by the actual concentrations in mol l
1 ; similarly, the fugacities (effective pressures) of gases can be substituted by

3

Heat change of reaction under constant pressure. T ¼ 298:15 K ð25 CÞ; standard activity of solutes, a ¼ 1; standard (partial) pressure of gases ¼ 101 kPa (standard fugacity ¼ 1). 5 ΔGStandard values at temperatures other than can be calculated via the integrated “Delta @ 298.15  KΔH ΔG version” of the Gibbs-Helmholtz equation @T T p ¼ T 2 : Assuming that temperature dependence of ΔH within the range of physiologically relevant temperatures is negligible, the free energy change at temperatures other than 298.15 K (but at standard activities) is 4

ΔGStandard ¼ T

T  ΔG 298:15

 þ

1


 T  ΔH 298:15

The same result is obtained from ΔG ¼ ΔH
TΔS S by assuming that ΔH and ΔS are essentially constant within the range of physiologically relevant temperatures.

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

5

their pressures in atm, an otherwise obsolete unit.6 With such simplification, as well as with R ¼ 8:315  10
3 kJ K
1 mol
1, T ¼ 298:15 K ð25 CÞ, the common use of kJ as energy unit, and ln x ¼ 2:303 lg x, (3) converts to 

ΔG ¼ ΔG þ 5:71 lg

½Cc ½Dd ½Aa ½Bb

ðat 298:15 KÞ

(4)

Hydrocarbons in the aqueous surroundings of microorganisms can be often considered with good approximation to have the activities of their gaseous, liquid, or solid standard states, viz., aHydrocarbon ¼ 1, or ½Hydrocarbon ¼ 1. For instance, if a gaseous hydrocarbon at standard pressure dissolves in water and reaches the dissolution equilibrium (ΔG of transfer ¼ 0), it is thermodynamically treated like the gas, even though the dissolved concentration is in the range of 10
3 M (Appendix Fig. 6). The same holds true for liquid hydrocarbons: Despite the extremely low saturation concentration of long-chain alkanes in water, the hydrocarbon dissolved in water has the activity (strictly speaking the chemical potential) of the pure liquid hydrocarbon phase. If inorganic (fully oxidized) Carbon is involved, also acid-base dissociation has to be Considered (Appendix Fig. 7). The free energy data (Appendix Table 6) reveal some basic and sometimes “counterintuitive” thermodynamic properties of hydrocarbons. Many hydrocarbons  are metastable (thermodynamically unstable; ΔGf positive) with respect to the elements, even though decay into the elements is usually “kinetically inhibited.” In the case of acetylene (ethyne), however, compression at room temperature can trigger the release of the energy in a violent decay into the elements. For this reason, compressed welding acetylene in steel bottles must be stabilized by adsorption to a carrier such as acetone. But also hydrocarbons that are stable with respect to the elements (even the rather stable ethane) are metastable with respect to decay into native carbon and methane, the most stable hydrocarbon: 2 C2 H6 ! CGraphite þ 3 CH4  ΔG ¼
43:3 kJðmol C2 H6 Þ
1

(5)

In the presence of CO2 or bicarbonate, even methane is metastable:

6

CH4 þ CO2 ! 2 CGraphite þ 2 H2 O  ΔG ¼
29:2 kJðmol CH4 Þ
1

(6)

CH4 þ CO2 ! 2 CGraphite þ 2 H2 O  ΔG ¼
29:2 kJðmol CH4 Þ
1

(7)

The apparent correctness of the old unit atm is due to the fact that it is numerically equivalent with standard fugacity ¼ 1. Activities and fugacities are by definition without units, and the formally ½A correct approximated substitution would aA ¼ ½A Actual , etc. Here, the use of the modern unit Pa or Standard kPa for [A], etc. is coherent.

6

F. Widdel and F. Musat

Nevertheless, formation of elemental carbon by reactions (5, 6, and 7) is kinetically strongly inhibited and has not been observed in abiotic or biotic systems at room temperature. But once the element has been formed by geothermal metamorphism of buried biomass or petroleum (Tissot and Welte 1984), it is the thermodynamically stable species of carbon as long as additional reducing or oxidizing components are absent. In the presence of a mild oxidant, not the element but rather CO2 and its ionic
forms, HCO3
and CO3 2 , are the thermodynamically stable forms of carbon. Other forms or intermediate oxidation states ( H2 C ¼ O, CO, HCOOH, C2 -compounds, etc.), like all natural organic compounds, are metastable7 with respect to a conversion to CH4, CO2 and H2O, even without involvement of an oxidant or reductant. If on the other hand a reductant with negative enough redox potential is present, the only stable form of carbon is CH4. Again, intermediate oxidation states (CH3OH, reduced C2-compounds, etc.) are metastable. The stability “regions” of the mentioned carbon species are elegantly illustrated in a plot of the redox potential versus the pH (E-pH-diagram, Pourbaix diagram; Fig. 2). Another thermodynamically interesting principle is revealed in the homologous series of n-alkanes. n-Alkanes become increasingly unstable with increasing chain length, whereas the heat of formation shows an opposite trend (Fig. 3).8 The heat is the energy released during C
H bond formation from the elements (even though such alkane formation is not observed in reality). Hence, the thermodynamically feasible disintegration of a long-chain alkane into its elements would consume heat: C16 H34 ! 16 CGraphite þ 17 H2  ΔG ¼
49:8 kJðmol C16 H34 Þ
1  ΔH ¼ þ454:4 kJðmol C16 H34 Þ
1

(8)

This thermodynamically “allowed” cooling of the surroundings (and the system), which is a decrease in the entropy of the surroundings, is explained by the numerically higher entropy increase of the reacting system; the molecules of the gaseous H2 that are formed in high number carry a high amount of “hidden heat.”9 Furthermore, the homologous n-alkane series reveals the transition from gaseous to liquid  hydrocarbons (n-butane/n-pentane), which is mirrored by a discontinuity of the ΔHf values. This is because liquid pentane has “given off” the heat of condensation to the  surroundings (liquid n-pentane, the real standard state: ΔH f ¼
173 kJ mol
1 ;

7

The extremely low hypothetical equilibrium concentrations of these species can be calculated. Linearity in the series of the higher alkanes may be a “pre-assumption” and basis for calculation of   ΔGf or ΔHf values of compounds in homologous series via incremental additions. In the numerous sources of thermodynamic data, the original basis underlying such data is often difficult to trace back. 9 Also, the highly ordered (“improbable”) structure of the long-chain alkane contributes to thermodynamic instability. 8

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

7

0.4

Co2 (g) 0.2 HCO3– 0.0 CO32–

C

E –0.2 [V]

H

–0.4

H

2O

2

CH4 (g)

–0.6

–0.8 0

2

4

6

10

8

12

14

pH

Fig. 2 Stability diagram (Pourbaix diagram) of carbon species. The equilibrium (borderline) redox 

potential E (in V) as a function of pH was calculated from ΔGf values (in J) according to E ¼ X X ΔGof, red ΔGof, ox
∏aox þ 0:0592 lg with n ¼ number of electrons; F ¼ 96, 485 C n F n ∏ared
1 þ mol ; a ¼ activity . Πaox includes the H -activity, the negative logarithm of which is the pH. Activities or fugacities: CO2 , CH4 , 1:0; HCO3
and CO3 2
, 10
2 (black) or 1.0 (gray). The borderlines between CO2 ðgÞ, HCO3
and CO3 2
in their standard states represent the pKa values. Note that the pKa1 value for CO2(g) is 7.8 (vertical gray line), whereas that of CO2 (aq) is 6.35 (not shown here), the more commonly known one. The system H2O/H2 (electrochemically the same as 2Hþ =H2 ) is indicated for comparison



hypothetical gaseous standard state: ΔHf ¼
146 kJ mol
1 ). The discontinuity  of ΔGf is less pronounced. Liquid pentane as a highly volatile compound (boiling point, 36.2  C) is almost in equilibrium with the gaseous state (liquid n-pentane:   ΔGf ¼
9:21 kJ mol
1; hypothetical gaseous standard state: ΔGf ¼
8:11 kJ mol
1).

8

F. Widdel and F. Musat

Fig. 3 Free energy and enthalpy (heat) of formation of alkanes from C1 to C18. The free energy of formation of C16 includes a literature value and the value extrapolated in this graph

+100

2

4

6

8

C-Atoms 10 12

14

16

18

ΔGf°

0

ΔGf° or ΔHf° [kJ mol–1]

–100 Gaesous

Liquid

Solid

–200 ΔHf°

–300

–400

–500

–600

3

Energetics of Hydrocarbon Utilization by Microorganisms

The biological utilization of a hydrocarbon can be examined bioenergetically (1) on the level of the net reaction performed by a microorganism and (2) on the level of individual enzymatic reactions. Among the latter, those of hydrocarbon activation are usually of highest interest and therefore briefly addressed in this overview.

3.1

Catabolic Net Reactions of Hydrocarbons from the Energetic Perspective

The Δ G of a reaction (the “system”) is the maximum amount of energy that a second system can theoretically conserve via coupling to this reaction under full reversibility. However, coupling can only proceed outside of the equilibrium, viz., if the overall reaction of the two systems is more or less irreversible and dissipates free energy. The actual amount of useful energy provided by the catabolic reactions is therefore always less than the calculated ΔG. The subsequent anabolism with many highly irreversible reactions then dissipates most of the free energy. Table 1 lists generalized equations for the degradation of hydrocarbons and Table 2 several particular reactions with naturally important electron acceptors and the associated energy changes.

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

9

Table 1 Generalized equations for the catabolism (dissimilation, degradation) of hydrocarbons with various electron acceptors and for the anabolism (assimilation) of hydrocarbons into cell mass Catabolism 4 Cc Hh þ ð4c þ hÞ O2 ! 4c CO2 þ 2h H2 O 10 Cc Hh þ ð8c þ 2hÞ NO3
þ ð8c þ 2hÞ Hþ ! 10c CO2 þ ð4c þ hÞ N2 þ ð4c þ 6hÞ H2 O Cc Hh þ ð4c þ hÞ FeðOHÞ3 þ ð3c þ hÞ CO2 ! ð4c þ hÞ FeCO3 þ ð6c þ 2hÞ H2 Oa


8 Cc Hh þ ð4c þ hÞ SO4 2 þ ð8c þ 2hÞ Hþ ! 8c CO2 þ ð4c þ hÞ H2 S þ 4h H2 O 8 Cc Hh þ ð8c
2hÞ H2 O ! ð4c
hÞ CO2 þ ð4c þ hÞ CH4 Anabolism 4 Cc Hh þ h CO2 þ ð4c
hÞ H2 O ! ð4c þ hÞhCH2 Oi dass ¼ 0:133=ð4c þ hÞ mol g
1 Y ass ¼ ð4c þ hÞ=0:133 g mol
1 17 Cc Hh þ ð4h
cÞ CO2 þ ð14c
5hÞ H2 O ! ð4c þ hÞ C4 H7 O3 dass ¼ 0:165=ð4c þ hÞ mol g
1 Y ass ¼ ð4c þ hÞ=0:165 g mol
1 17 Cc Hh þ ð4h
cÞ CO2 þ ð4c þ hÞ NH3 þ ð10c
6hÞ H2 O ! ð4c þ hÞ C4 H8 O2 N dass ¼ 0:166=ð4c þ hÞ mol g
1 Y ass ¼ ð4c þ hÞ=0:166 g mol
1 a

Because many reactions take place in environments containing inorganic carbon, reactions with Fe(OH)3 are for convenience written with the relatively insoluble FeCO3 (siderite) as a product

The aerobic oxidation with O2 as electron acceptor provides biochemically the highest amount of energy, methanogenic degradation the lowest. Reactions with NO2
or N2O are even more exergonic than those with O2 (Table 2 includes methane oxidation with nitrite as an investigated example; Ettwig et al. 2008). However, there is no evidence that the higher energy available with NO2
or N2O in comparison to O2 as electron acceptor is conserved; biochemically, O2 allows conservation of even more energy from the same amount of organic substrate. The theoretically higher energy gain is due to the “extra energy content” of NO2
and N2O with respect to O2 : 4 NO2
þ 4 Hþ ! 2 N2 þ 3 O2 þ 2 H2 O, ΔG0 ¼
116 kJðmol NO2
Þ
1 ; 2 N2 O ! 2 N2 þ O2 , ΔG ¼
104 kJðmol N2 OÞ
1 . One of the least exergonic catabolic reactions is the anaerobic oxidation of methane (Table 2). Under certain environmental conditions, the net free energy change under in situ concentrations of the reactants may be only around ΔG ¼
20 kJ molðCH4 Þ
1 (Nauhaus et al. 2002). The fact that this minute amount is further shared between two organisms with different metabolism challenges the energetic understanding of energy conservation under “low-energy” conditions (viz., life at low chemical potential), a topic developed in the study of other syntrophic associations (Jackson and McInerney 2002; Schink 1997, 2002; chapter ▶ Introduction to Microbial Hydrocarbon Production: Bioenergetics by McInerney et al. in this volume). Another anaerobic reaction of a hydrocarbon of thermodynamic interest is the


813.3

! 3 CO2 ðgÞ þ 4 H2 O

! 6 CO2 ðgÞ þ 5 H2 SðaqÞ þ 8 H2 O

! 5 CH4 ðgÞ þ CO2 ðgÞ

2 C3 H8 ðgÞ þ 5 SO4 2
þ 10 Hþ

2 C3 H8 ðgÞ þ 2 H2 O

n -Hexane

! 7 CH4 ðgÞ þ CO2 ðgÞ

Propane C3 H8 ðgÞ þ 5 O2 ðgÞ

þ 14 H

4 C2 H6 ðgÞ þ 2 H2 O

4 C2 H6 ðgÞ þ 7 SO4

2


! 8 CO2 ðgÞ þ 7 H2 SðaqÞ þ 12 H2 O


63


117


2,108


36


73


1,467

! 4 CO2 ðgÞ þ 6 H2 O

Ethane 2 C2 H6 ðgÞ þ 7 O2 ðgÞ

CH4 ðgÞ þ SO4

þH

þ

2


CH4 ðgÞ þ SO4

þ


16.6d,e

! HCO3
þ HS
þ H2 O

CH4 ðgÞ þ SO4 2



344d
16.5d,e

þ2H

2


þ

! HCO3
þ H2 S ðaqÞ þ H2 O

! 8 FeCO3 þ 14 H2 O

CH4 ðgÞ þ 8 FeðOHÞ3 þ 7 CO2 ðgÞ


928.3
21.2d

! 3 CO2 ðgÞ þ 4 N2 ðgÞ þ 10 H2 O

3 CH4 ðgÞ þ 8 NO2
þ 8 Hþ


766.2

! CO2 ðgÞ þ H2 SðaqÞ þ 2 H2 O

! HCO3 þ H þ H2 O

! 5 CO2 ðgÞ þ 4 N2 ðgÞ þ 14 H2 O

5 CH4 ðgÞ þ 8 NO3
þ 8 Hþ

d


818.0d




þ

! CO2 ðgÞ þ 2 H2 O

Productsa

CH4 ðgÞ þ 2 O2 ðgÞ

Reactantsa Methane CH4 ðgÞ þ 2 O2 ðgÞ

Free energy change of reaction per mol hydrocarbonb, ΔG or ΔG0 (kJ mol
1 )

6


46


2,220


2


38


1,560


11.3


33.5


20.9


559


993


788


903.0


890.4

Enthalpy change of reaction per mol hydrocarbon, ΔH  (kJ mol
1 )

232

236


375

115

117


310

18


57

1


723


215


73


301


243

Entropy change of reaction per mol hydrocarbonb,c, ΔS or ΔS0 (J K
1 mol
1 )

Table 2 Thermodynamic characteristics of observed and some hypothetical reactions of selected hydrocarbons. The degradation of hydrocarbons to methane and carbon dioxide is often endothermic

10 F. Widdel and F. Musat


238
142

! 15 CH4 ðgÞ þ 9 CO2 ðgÞ

! 7 CO2 ðgÞ þ 4 H2 O

! 35 CO2 ðgÞ þ 18 N2 ðgÞ þ 38 H2 O

! 36 FeCO3 þ 58 H2 O

! 14 CO2 ðgÞ þ 9 H2 S ðaqÞ þ 8 H2 O

! 9 CH4 ðgÞ þ 5 CO2 ðgÞ

! 10 CO2 ðgÞ þ 4 H2 O

! 50 CO2 ðgÞ þ 24 N2 ðgÞ þ 44 H2 O

4 C6 H6 ðlÞ þ 18 H2 O

Toluene C7 H8 ðlÞ þ 9 O2 ðgÞ

5 C7 H8 ðlÞ þ 36 NO3
þ 36 Hþ

C7 H8 ðlÞ þ 36 FeðOHÞ3 þ 29 CO2 ðgÞ

2 C7 H8 ðlÞ þ 9 SO4 2
þ 18 Hþ

2 C7 H8 ðlÞ þ 10 H2 O

Naphthalene C10 H8 ðcÞ þ 12 O2 ðgÞ

5 C10 H8 ðcÞ þ 48 NO3
þ 48 Hþ


4,782


5,093


1,689


3,590


3,823


135


214


1,423

! 30 FeCO3 þ 48 H2 O


3,008


3,202

! 24 CO2 ðgÞ þ 15 H2 SðaqÞ þ 12 H2 O

! 30 CO2 ðgÞ þ 15 N2 ðgÞ þ 30 H2 O

5 C6 H6 ðlÞ þ 30 NO3
þ 30 Hþ

C6 H6 ðlÞ þ 30 FeðOHÞ3 þ 24 CO2 ðgÞ

! 12 CO2 ðgÞ þ 6 H2 O

Benzene 2 C6 H6 ðlÞ þ 15 O2 ðgÞ


372


632


9,757

4 C6 H6 ðlÞ þ 15 SO4 2
þ 30 Hþ

! 49 CH4 ðgÞ þ 15 CO2 ðgÞ

! 64 CO2 ðgÞ þ 49 H2 S ðaqÞ þ 68 H2 O

4 C16 H34 ðlÞ þ 30 H2 O

4 C16 H34 ðlÞ þ 49 SO4

2


þ 98 H

! 80 CO2 ðgÞ þ 49 N2 ðgÞ þ 134 H2 O

þ

! 32 CO2 ðgÞ þ 34 H2 O

5 C16 H34 ðlÞ þ 98 NO3
þ 98 Hþ


10,392


137

4 C6 H14 ðlÞ þ 19 SO4

! 19 CH4 ðgÞ þ 5 CO2 ðgÞ


4,023

4 C6 H14 ðlÞ þ 10 H2 O

! 12 CO2 ðgÞ þ 14 H2 O

n -Hexadecane 2 C16 H34 ðlÞ þ 49 O2 ðgÞ

þ 38 H

þ


238

2


! 24 CO2 ðgÞ þ 19 H2 SðaqÞ þ 28 H2 O

2 C6 H14 ðlÞ þ 19 O2 ðgÞ


4,541


5,156

96

2


2,421


3,449


3,910

71


7


2,026


2,883


3,268

206


50


9,445


10,701

66


33


4,163

808


212

801

806


2,456

473


293

691

696


2,024

419


220

1,937

1,952

1,047


1,036

682

688


471

(continued)

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . 11

! 10 CO2 ðgÞ þ 6 H2 S ðaqÞ þ 4 H2 O

! 6 CH4 ðgÞ þ 4 CO2 ðgÞ

C10 H8 ðcÞ þ 6 SO4 2
þ 12 Hþ

C10 H8 ðcÞ þ 8 H2 O


186


313

Free energy change of reaction per mol hydrocarbonb, ΔG or ΔG0 (kJ mol
1 )
2,247 186

61

Enthalpy change of reaction per mol hydrocarbon, ΔH  (kJ mol
1 )
3,171

1,245

1,253

Entropy change of reaction per mol hydrocarbonb,c, ΔS or ΔS0 (J K
1 mol
1 )
3,098

b

Indicated standard states: g, gaseous; l, liquid; aq, aqueous, dissolved in water; c, crystalline If protons are involved, ΔG 0 (viz., for ½Hþ  ¼ 10
7 M, pH ¼ 7) is given    c Here calculated via ΔS ¼ ΔH
ΔG T d Standard free energy changes of reactions formulated with CO2 differ from those formulated with HCO3
because the reaction HCO3
þ Hþ ! CO2 ðgÞ þH2 O is exergonic under standard conditions at pH ¼ 7, with ΔG0 ¼
4:7 kJ mol
1 e Reactions with H2S and HS
as products are energetically equivalent because both sulfide species are essentially in equilibrium under standard conditions

a

Productsa ! 48 FeCO3 þ 76 H2 O

Reactantsa C10 H8 ðcÞ þ 48 FeðOHÞ3 þ 38 CO2 ðgÞ

Table 2 (continued)

12 F. Widdel and F. Musat

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

13

conversion of alkanes to methane (Anderson and Lovley 2000; Jones et al. 2008; Zengler et al. 1999), an endothermic reaction (for explanation, see remark on (8)): 4 C16 H34 þ 30 H2 O ! 49 CH4 þ 15 CO2 ðgÞ  ΔG ¼
372 kJðmol C16 H34 Þ
1  ΔH ¼ þ206 kJðmol C16 H34 Þ
1

(9)

The Gibbs-Helmholtz equation predicts that the reaction becomes increasingly exergonic with increasing temperature (Dolfing et al. 2008). The process involves three organisms, (1) the hexadecane-degrading syntrophs ðC16 H34 þ 16 H2 O ! 8CH3 COO
þ 8 Hþ þ 17 H2 Þ , (2) acetate-cleaving microorganisms which are either methanogens ðCH3 COO
þ Hþ ! CH4 þ CO2 Þ or additional syntrophs ðCH3 COO
þ Hþ þ 2 H2 O ! 2 CO2 þ 4 H2 Þ, and (3) H2-utilizing methanogens ðCO2 þ 4 H2 ! CH4 þ 2 H2 OÞ. The available energy per transferred acetate, the “metabolic unit” in this syntrophism, is only around
47 kJ mol
1; this amount is shared between three organisms. The thermodynamic constraints of this reaction with respect to petroleum hydrocarbon conversion to methane have been examined (Dolfing et al. 2008).

3.2

Hydrocarbon Activation from the Energetic Perspective

As any chemical or biochemical reaction, the activation reaction of hydrocarbons involves two energetic aspects. These are the net ΔG of the reaction (and its share in the overall catabolic ΔG), and the energy level which during the activation reaction is attained by the energy-rich short-lived transition state in the active site of the hydrocarbon-activating enzyme; an apparent transition state may further resolve into elementary reactions upon closer examination (Fig. 4). Net free energy changes of several activation reactions of hydrocarbons are listed in Table 3. The activation of a hydrocarbon by introduction of a functional group to allow further metabolic processing is usually not a problem from a merely thermodynamic point of view. For instance, an O2-independent hydroxylation of methane by dehydrogenation at a hypothetical “methane dehydrogenase” employing a mildly oxidizing biological agent such as cytochrome cðCyt cox =Cyt cred , E ¼ E0 ¼ þ0:245 VÞ would be thermodynamically allowed: CH4 þ 2 Cyt c½Fe3þ  þ H2 O ! CH3 OH þ 2 Cyt c½Fe2þ  þ 2 Hþ ΔG 0 ¼
16 kJðmol CH4 Þ
1 

(10)

The problem lies in the high energy barrier, mainly due to the apolar and very stable C
H bond that must be attenuated by an appropriate biocatalyst. Despite the astounding capabilities of enzymes to decrease energy barriers of chemically difficult reactions, there is not always the ideal biochemical solution to any activation

14

F. Widdel and F. Musat ΔG of activation

Without catalyst S

Enzymatic, simple model

Enzymatic, elementary reactions

Net ΔG of activation reaction

1 2

ΔG

ΔG of catabolic reaction

3

4 5

7

8

9 P Proceeding reactions

Fig. 4 Free energy changes during a fictive reaction of a hydrocarbon (S; in principle any other substrate) converted to an end product (P), free energy change of the activation reaction, and free energies of transition states at the activating enzyme. The scheme is a simplification because it does not display any electron acceptor that allows oxidation and the indicated free energy changes

problem. Not every thermodynamically possible but kinetically inhibited reaction can be catalyzed to take place at any rate.10 To overcome the activation barrier and to reach high rates in such cases, activating enzymes invest an extra input of energy that is not conserved and makes the activation highly irreversible. Oxygenases which involve a strong oxidant (O2/2 H2O, E0 ¼ þ 0:818 V) and produce water besides the organic activation product are such energy-“wasting” catalysts that achieve high rates. The reaction with methane is CH4 þ O2 þ NADH þ Hþ ! CH3 OH þ NADþ þ H2 O  ΔG 0 ¼
344 kJ mol-1

(11)

The sacrifice of energy to achieve activation via oxygenases is also reflected by the consumption of reducing equivalents detained from the energy-conserving A prominent example is nitrogenase: Despite the long evolution of nitrogen fixation, an enzyme type has not evolved that catalyzes the thermodynamically feasible N2 reduction with H2 or energetically equivalent electron donors without an investment of energy.

10


35 to
39
31 to
35

R
CH2
CH3 þ
OOC
CH ¼ CH
COO
!
OOC
½ðRÞ
CH
ðCH3 ÞCH
HCH
COO


C6 H5 CH 3 þ
OOC
CH ¼ CH
COO
!
OOC
½C6 H5 CH 2 CH
HCH
COO


n-Alkane

Toluene


27 to
31

Addition to fumarate CH4 þ
OOC
CH ¼ CH
COO
!
OOC
½CH3 CH
HCH
COO
Methanee


335

C6 H6 þ O2 ! ½Intermediate; NADH
recycling ! o-C6 H4 ðOHÞ2

þ30


368

C15 H31
CH3 þ O2 þ NADH þ Hþ ! C15 H31
CH2 OH þ H2 O þ NADþ


344c

Free energy b  or change  ΔG
1 0 ΔG kJ mol

Methyl-coenzyme M reductase Methane CH4 þ CoM
S
S
CoB ! CoM
S
CH3 þ HS
CoB

nHexadecaned Benzene

Type of activation; compound Reactiona Mono- and dioxygenation Methane CH4 þ O2 þ NADH þ Hþ ! CH3 OH þ H2 O þ NADþ

(continued)

Rabus et al. (2001) Rabus et al. (2001) Rabus et al. (2001)

Shima and Thauer (2005)

Widdel et al. (2007) This article Widdel et al. (2007)

Reference

Table 3 Free energies of activation reactions of saturated, monounsaturated, and aromatic hydrocarbons. Also purely hypothetical reactions have been included. Values are given for standard conditions (pH ¼ 7, if protons are involved)

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . 15


31

Widdel et al. (2007) Widdel et al. (2007)

Widdel et al. (2007)

Reference

b

Fate of one reactant is visualized in bold ΔG 0 is indicated if protons are involved c Would be less exergonic with an electron donor of less negative redox potential than that of NADþ =NADH ð
0:320 VÞ d Sum of the following formal reactions: C15 H31
CH3 þ 0:5O2 ! C15 H31
CH2 OH , ΔG ¼
148:5 kJ mol
1 ; 0:5O2 þ NADH þ Hþ ! H2 O þ NADþ , ΔG0 ¼
219:6 kJ mol
1 (calculated from ΔE0 ¼ 1:138 V; Thauer et al. 1977) e Hypothetical reaction f Carboxylations have been suggested on the basis of chemical analyses g A carboxyl carrier and donor have not been suggested or identified. The present free energy change is based on a calculation with oxaloacetate as a purely hypothetical carboxyl donor that may be energetically comparable to potent carboxyl donors such as carboxy-biotin

a

C6 H6 þ Carrier
COO
! C6 H5
COO
þ Carrier
H


4

Carboxylationf Benzene CH3
CH2
CH ¼ CH2 þ H2 O ! CH3
CH2
HCH
CH2 OH

Benzeneg


7

Free energy b  or change  ΔG
1 0 ΔG kJ mol

Type of activation; compound Reactiona Addition of water to isolated double bond CH3
CH2
CH ¼ CH2 þ H2 O ! CH3
CH2
HCH
CH2 OH Butenee

Table 3 (continued)

16 F. Widdel and F. Musat

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

17

respiratory chain: The oxygenase reaction consumes two reducing equivalents from the metabolism, and the insertion of the oxygen atom to yield the alcohol “cancels” two additional reducing equivalents; hence, four reducing equivalents are consumed. Despite the significant amount of free energy dissipated and reducing power consumed by oxygenase reactions, this drain is not critical. The total free energy from the aerobic oxidation in this example is CH4 þ 2 O2 ! CO2 þ 2 H2 O  ΔG ¼
818 kJðmol CH4 Þ
1

(12)

From the totally available 8 [H] per methane, 4 [H] are still available for the respiratory chain. With higher hydrocarbons, the drain of energy and reducing equivalents are even less relevant. An activation of hydrocarbons under anoxic conditions excludes oxygen11 and in the case of many catabolic net reactions with low-energy gain strongly restricts the energy that can be dissipated to achieve activation. A reaction with particularly low net energy gain is the anaerobic oxidation of methane with sulfate (Table 2). The activation reaction is most likely a reversal of the methyl-coenzyme M reductase (Mcr) reaction, the final step in methanogenesis which is exergonic under standard conditions ( CoM
S
CH3 þ HS
CoB ! CoM
S
S
CoB þ CH4 , ΔG ¼
30 ½10 kJ mol
1 ; Shima and Thauer 2005; Thauer and Shima 2008). For methane activation, the standard free energy of the reverse Mcr (rMcr) reaction in methane oxidizing archaea would thus be þ30 ½10 kJ mol
1 . Methane activation with the disulfide CoM
S
S
CoB can therefore only take place if the products CoM
S
CH3 and HS
CoB are kept at very low concentration by effective scavenge in subsequent reactions. With respect to energy conservation in the total process, such a highly “concentration-controlled” reaction would be advantageous because it would be always very close to the equilibrium and not dissipate much energy. For the activation of the strong C
H-bond of methane (absolute value,
440 kJ mol
1; McMillen and Golden 1982) by a thiyl radical or a NiIII center, which may be the most critical step, a decrease of the activation energy by a “dual-stroke engine” mechanism was proposed (Thauer and Shima 2008). rMcr has presumably two active sites, like Mcr. The release of the products from one site may transfer conformational energy to the other site where the substrates enter the reaction. However, this does not influence the equilibrium of the net activation reaction. Methane activation via a reversal of the Mcr reaction is not only of mechanistic but also of kinetic interest. The positive standard free energy change of the rMcr reaction sets severe limits to the rate of the formation of the initial intermediates.

11

The utilization of chlorate by facultatively anaerobic bacteria for hydrocarbon metabolism (Chakraborty and Coates 2004; Tan et al. 2006) involves O2 that is generated from an intermediate ðClO2
! Cl
þ O2 Þ.

18

F. Widdel and F. Musat

Using the Haldane equation,12 which connects the catalytic efficiencies of the forward and back reactions through an enzyme with the thermodynamic equilibrium constant of the reaction, the first step in AOM was estimated to be slower by a factor between 10
3 and 10
7 than the final step in methanogenesis (Shima and Thauer 2005; Thauer and Shima 2008). Also, the rate of the subsequent enzymatic step may be drastically limited by the low near-equilibrium concentrations of methylcoenzyme M and coenzyme B. The high content of the apparent rMcr in naturally enriched anaerobic methane oxidizers (Krüger et al. 2003) may be a means to compensate for the slowness of the enzyme. The carbon–carbon addition of non-methane hydrocarbons at their methyl or methylene group to fumarate is slightly exergonic (Table 3) and to our present knowledge not associated with energy conservation. However, in view of the net energy gain with non-methane hydrocarbons under anoxic conditions, such a loss is “affordable.” Only methane activation in an analogous way to yield methylsuccinate would be critical in an oxidation of methane with sulfate. The suggested mechanistic steps are an abstraction of a specific glycyl hydrogen in the polypeptide chain by a protein-activating enzyme (yielding
Gly▪
), subsequent hydrogen abstraction from a cysteyl group by the glycyl radical (yielding
CysS▪
), abstraction of a methyl hydrogen from toluene (yielding C6 H5
▪ CH2 ), addition of the benzyl radical to fumarate (yielding the benzylsuccinyl radical), and quenching of the radical to yield free benzylsuccinate and regenerate the cysteyl radical for the next catalytic round (chapter ▶ Anaerobic Degradation of Hydrocarbons: Mechanisms of C–H-Bond Activation in the Absence of Oxygen by Boll and Heider, this volume). Quantum chemical modeling of this reaction, for which a crystal structure of the enzyme was not available, supported the feasibility of the suggested steps (Himo 2002, 2005). The ratelimiting step was calculated to be the addition of the benzyl radical to fumarate.

4

Quantitative Aspects of Cell Synthesis

4.1

ATP and Growth Yields

The more exergonic a catabolic reaction and the higher the efficiency of ATP synthesis (proportion of total free energy conserved in ATP), the more cell mass can be synthesized from a given substrate. The quantitative treatment of the efficiency of free energy conservation in the form of ATP and the amounts of cell mass formed with various substrates are subjects of an own area of research in microbiology. In this research, the measurable molar growth yield is of central interest, 12

The Haldane equation describes the connection between the equilibrium concentrations of the reactants and products and their kinetic constants kcat and Km. The equilibrium constant is also thermodynamically
 given by the concentrations at ΔG ¼ 0. In case of the reaction S ! P, the connection is

½P ½S eq

kS =K S



m ¼ kcat ¼ e
ΔG P =K P cat

m

=ðRT Þ

:

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

19

besides calculated free energy changes and ATP yields known from well-established pathways such as glycolysis. The molar growth yield, Y, is defined as the amount of cell dry mass, X (in g) per amount of totally consumed substrate, Stot (in mol). On the other hand, for indication of the energy gain from the catabolism, a growth yield with respect to the dissimilated (viz., the energy yielding) proportion of the substrate, Sdiss, would be a more meaningful definition: Y¼

 X  g mol-1 Stot

Y diss ¼

X  Sdiss

g mol-1



(13a; b)

However, the latter definition and distinctive subscripts are not very common. Sdiss can be determined experimentally by quantifying the biomass, X, and the consumed
electron acceptor (O2, NO3
, FeIII, or SO4 2 ) or at least one of the products (CO2, N2, FeII/III, or H2S). The chemically formulated stoichiometric relationship between substrate and product (Table 1) then reveals Sdiss, which leads to Ydiss (13b). The fraction of the dissimilated substrate as part of the totally consumed substrate in anaerobic bacteria is usually much higher than in aerobic bacteria:     Sdiss Sdiss > Stot anaerobic Stot aerobic

(14)

Some measured growth yields of aerobic and anaerobic hydrocarbon utilizing microorganisms are listed in Table 4. If consumption of the electron acceptor or formation of the catabolic product has not been quantified, or if only a Y value (13a) has been reported, Ydiss can be calculated. With Sass for the assimilated amount of substrate, the totally consumed substrate is Stot ¼ Sdiss þ Sass

(15)

Division by the obtained cell mass yields Stot Sdiss Sass ¼ þ X X X

(16)

1 1 Sass ¼ þ Y Y diss X

(17)

and with definitions (13a, b)

The expression Sass/X may be termed the assimilatory substrate demand, dass (in mol g
1 ). The reciprocal term X/Sass can be defined as another type of yield, the amount of cell mass (in g) obtained per assimilated amount of substrate (in mol), and designated Yass. The connection is thus d ass ¼ 1=Y ass . This leads to

n-Heptadecane, O2 Benzene, O2

Toluene, O2

Toluene, O2

Micrococcus cereficans Candida tropicalis Ideal aerobe

Pseudomonas nautica Pseudomonas putida

Pseudomonas putida

Pseudomonas putida 1.0*

1.28*

0.50 1.20*

127f to 280f

0.60*,e to 1.32* 0.9* 1.0* 1.77

n-Octane, O2

92.1

159

185

148 197

120*,g 94.7 100

310 366 1240

204 226 401

164f to 563f

214

125

1.1*

Hydrocarbon and electron acceptor

Long-chain n-alkane mixture, O2 n-Hexadecane, O2 n-Hexadecane, O2 n-Hexadecane, O2

Nocardia sp.

Microorganism Aerobic Pseudomonas sp.

Growth yieldb (cell dry mass per amount of hydrocarbon) c molar, by mass   molar  ,  ðg g
1 Þ Y g mol
1 Y diss g mol
1

0.58 (58%)

0.54 (54%)

0.81 (81%) 0.48 (48%)

0.66 (66%) 0.62 (62%) 0.32 (32%)

0.77 (77%) to 0.50 (50%)

0.58 (58%)

Fraction of substrate catabolizedd, Sdiss/Stot

Einsele (1983) Einsele (1983) This article (see text) Bonin et al. (1992) Reardon et al. (2000) Reardon et al. (2000) Bordel et al. (2007)

Wodzinski and Johnson (1968) Wagner et al. (1969)

Reference

Table 4 A selection of growth yieldsa of aerobic and anaerobic microorganisms on hydrocarbons and calculated fraction of the dissimilated substrate

20 F. Widdel and F. Musat

0.83 0.037 0.059 0.31

Methane, SO4 2


n-Hexadecane, SO4 2


Toluene, SO4 2


0.70* 0.50*

Ethylbenzene, NO3


Toluene, O2 Naphthalene, O2

33*

13.8

13.5*,h 29*

0.6*

129*

91.6 82

0.59

88*

64.5 64.1

0.88 (88%)

0.97 (97%)

0.99 (99%)

0.68 (68%)

0.70 (70%) 0.78 (78%)

Rabus and Widdel (1995) Nauhaus et al. (2007) So and Young (1999) Rabus et al. (1993)

Dinkla et al. (2001) Wodzinski and Johnson (1968)

Only directly measured (“real”) growth yields are listed and not Ymax values obtained from extrapolation b Original value from the reference is indicated by asterisk; other values were calculated for this chapter (see text). Ydiss was calculated via (19). The needed Yass was calculated according to Table 3, assuming the biomass bulk formula C4H7O3; the Yass values (g mol
1 ) are as follows: methane, 48.5; n-octane, 303; npentadecane, 558; n-hexadecane, 594; n-heptadecane, 630; benzene, 182; toluene, 218; ethylbenzene, 255 c Relative to dissimilated substrate d Calculated according to Sdiss =Stot ¼ Y=Y diss e Additional assimilation of added yeast extract is likely f For convenience calculated with pentadecane (which was part of the mixture) g With 2% O2 in gas phase; with more O2 the yield decreased h Estimated by assuming that 55% of cell dry mass is protein

a

Anaerobic Betaproteobacterium, strain EbN1 Archaea (ANME-2) Deltaproteobacteria Deltaproteobacterium, strain AK-01 Desulfobacula toluolica

Pseudomonas putida Pseudomonas sp.

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . 21

22

F. Widdel and F. Musat

1 1 1 ¼ þ Y Y diss Y ass

(18)

Y ass  Y Y ass
Y

(19)

Rearrangement leads to Y diss ¼

The values for dass or Yass are calculated from chemically formulated stoichiometries. This requires the assumption of bulk formulas for cell dry mass. The simplest bulk formula is that of carbohydrates, hCH2Oi. For aerobic methanotrophs, the formula hC4H8O2Ni was used (van Dijken and Harder 1975). A simpler N-free variant with the same bulk oxidation state of carbon is hC4H7O3i (Pfennig and Biebl 1976). A precise yet more complicated formula, hC4.36H8.24O1.87Ni, was determined for an aerobic bacterium grown with heptadecane (Bonin et al. 1992). Considering the oxidation state of carbon is more important than including nitrogen. Because in the case of hydrocarbons the substrate carbon is more reduced than cell mass carbon, CO2 is included in the assimilation equations (Table 1). Now, also the fraction of the dissimilated substrate as part of the totally consumed substrate can be calculated even if only a Y value is available from the literature: Sdiss Y ass
Y ¼ Y ass Stot

(20)

For instance, for aerobic growth with hexadecane (M ¼ 226:45), a growth yield by mass of 1 g ðg C16 H34 Þ
1 has been reported, which equals a molar growth yield of Y ¼ 226 g ðmol C16 H34 Þ
1 . The assimilation equation is 17 C16 H34 þ 120 CO2 þ 54 H2 O ! 98 C4 H7 O3 dass ¼ 1:68  10
3 mol g
1 ; Y ass ¼ 594 g mol
1

(21)

Equation (19) yields Y diss ¼ 366 g mol
1. The fraction of the dissimilated substrate is Sdiss ¼ 0:62 ðor 62%Þ Stot

(22)

Above all, Y values are expected to provide information about the ATP yield as a parameter of high relevance to understand the efficiency of or losses in the energy flow: Free energy of catabolic reaction # Free energy in formed ATP # Free energy ðor ATPÞ consumed for cell synthesis

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

23

The ATP yield or qATP is the amount of ATP (in mol) formed per amount of dissimilated substrate (in mol). At first glance, the concept appears straightforward. From anaerobic pathways with biochemically known qATP, as for instance the homolactic fermentation of glucose ðqATP ¼ 2Þ, the amount of cell mass obtained per mol ATP, the so-called YATP, can be calculated from the determined growth yield via Y ATP ¼ Y diss =qATP. If for another bacterium of interest, the qATP is unknown but Ydiss has been determined, this should in principle allow to calculate the desired qATP parameter via qATP ¼ Y diss =Y ATP .13 However, there is a serious drawback in that determined YATP values, viz., the energy expenses for biomass synthesis, vary enormously for different growth substrates and among various bacteria. This is not surprising because synthesis of an amount of biomass for instance from free acetate as the growth substrate needs more ATP than synthesis from carbohydrates and amino acids added to the medium. But even with the same substrate for biosynthesis, determined YATP values among bacteria vary significantly. These problems are treated by the calculation of theoretical ATP demands for the synthesis of biomass with its diverse fractions (polysaccharides, protein, etc.) from starting substrates and by consideration of the fractions of energy or ATP that do not lead to productive growth. This nonproductive consumption of energy or ATP is interpreted as maintenance energy (Pirt 1965; Tempest and Neijssel 1984), an uncoupling of the anabolism from the catabolism at varying extent, or an extra “spill” of energy (Russell 2007) in addition to the “regular” dissipation. In the concept of Pirt (1965), the proportion of the substrate consumed per time for maintenance rather than for productive growth is regarded as a constant that is independent of the growth rate, μ. Hence, the slower the growth of a bacterium and the lower the biomass production per time, the higher the proportion of the substrate consumed for maintenance. If therefore growth yields of an organism at different growth rates are extrapolated to a theoretical infinitely high growth rate (no time required for growth) in a plot of 1/Ydiss versus 1/μ, the proportion of the substrate consumed for maintenance should become zero. At 1=μ ¼ 0 ðμ ¼ 1Þ the theoretically highest growth yield, Ymax (more precisely Ydissmax) is obtained that is used to gain information about qATP and YATP. Such concepts have been applied to vast series of non-hydrocarbon substrates (Heijnen and van Dijken 1992; Stouthamer 1988). In the case of hydrocarbons, aerobic methanotrophs (Leak and Dalton 1985; van Dijken and Harder 1975) and degraders of long-chain alkanes (Erickson 1981; Ferrer and Erickson 1979) have been of interest for such mainly theoretical studies. If the catabolism of a substrate is likely to involve conventional biochemical reactions (β-oxidation, citric acid cycle, dehydrogenations with NADþ and flavoenzymes, etc.) and an aerobic respiratory chain, a qATP value can be also predicted from the ATP-yielding reactions. Via YATP values determined in other

The qATP is conceptually related to the P=2e
ratio in aerobic and anaerobic respiration which indicates the number of ATP molecules formed per electron pair transported in the respiratory chain (in aerobes also P/O ratio). However, the qATP also includes ATP from substrate level phosphorylation.

13

24

F. Widdel and F. Musat O2 H2O

ATP

C15H31–CH2OH

C15H31–CH3

C15H31–CHO

NAD–H

NAD–H

7 (FADH2) + 7 NAD–H

15 OH2 + 32 NAD–H 30 e–

C15H31–CO–SCoA

NAD–H

Consumption not depicted 31 H2O

C15H31–COOH

Reducing equivalents

β–Oxidation

8 CH3–CO–SCoA

64 e–

47 H2O

8 (FADH2) + 24 NAD–H

Respiratory chain

TCA

16 CO2

23.5 O2 ≈ 26 ATP

≈ 91 ATP

8 ATP

≈ 124 ATP Catabolism: C16H34 + 24.5 O2 → 16 CO2 + 17 H2O –1

ΔG° = –10 393 kJ (mol C16H34)

Fig. 5 Reducing equivalents and ATP synthesis in the aerobic catabolism of hexadecane ðC16 H32 , M ¼ 226:45Þ. Reducing equivalents from enzyme-bound FADH2 enter the respiratory chain at the quinone (Q) level. The assumed proton translocation in the respiratory chain underlying this scheme is 10 Hþ =NADH and 6 Hþ =QH2 . A phosphorylation yield of 1 ATP per 3:5 Hþ was arbitrarily assumed here (based on the commonly assumed range of 1 ATP per 3 to 4 Hþ ). The resulting net ATP yield is thus 124 mol ATP per mol C16H34, 0.55 mol ATP per g C16H34, or 5.3 mol ATP per mol O2. For comparison, glucose (C6 H12 O6 , M ¼ 180:16) would yield 10 NADH and 2 QH2 allowing formation of 32 ATP via respiration; with 4 ATP from glycolysis and the tricarboxylic acid cycle, the net yield is 36 mol ATP per mol C6H12O6, 0.20 mol ATP per g C6H12O6, or 6.0 mol ATP per mol O2

studies, a Ymax can be subsequently predicted and compared to an experimentally determined one. As an example, Fig. 5 presents the catabolic scheme for aerobic degradation of hexadecane with qATP ¼ 124 ðmol=molÞ. According to the free energy change of the reaction (
10 392 kJ mol
1 ; Fig. 5), the average energy need for ATP synthesis would be 100 kJ ðmol ATPÞ
1. If a YATP of 10 g cell dry mass ðmol ATPÞ
1 is assumed that is likely for cell synthesis from the hexadecane-derived acetate units (Erickson 1981; Stouthamer 1988), this would lead to Y diss ¼ 1240 g cell mass ðmol C16 H34 Þ
1 . The Y value is obtained via a transformation of (19): Y¼

Y ass  Y diss Y ass þ Y diss

(23)

This yields (with the above Y ass ¼ 594 g mol
1 ) a value of Y ¼ 401 g mol
1 , which would be a yield by mass of 1.77 g cell mass ðg C16 H34 Þ
1 . This may be regarded as an “ideal” yield with hexadecane. The fraction of dissimilated hexadecane would be only

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

CH4 (g) a = 719 (ΔGf = –34.4)

–16.3 kJ +16.3 kJ

25

CH4 (g) a = 1.00 ΔGf ° = –50.75 kJ

Gas phase 0 kJ

0 kJ

0 kJ

0 kJ

Water phase

CH4 (g) a = 1.00 ΔGf ° = –34.4 kJ

–16.3 kJ +16.3 kJ

CH4 (aq) a = 0.00139 (ΔGf = –50.75)

All free energie values per mol

Fig. 6 The two standard states (framed) of methane. Aqueous methane, CH4(aq), in its standard state which corresponds to a very high partial pressure has a higher energy content than gaseous methane, CH4(g), in its standard state. Hence, indication of the ΔG oder ΔG 0 of a formulated reaction involving methane must indicate the applied standard state. Calculation of the free energy change of a reaction for real (measured) pressures or concentrations (according to (4)) must yield the same result with each standard state. Application of the gaseous standard state for calculation is also justified if there is no gas phase. Most natural conditions will be closest to the gaseous standard  state. The ΔGf of CH4 (aq) was calculated via the solubility of 0.00139 mol l
1 atm
1 (Wilhelm et al. 1977), assuming that this concentration is numerically equivalent with the activity of CH4 (aq) that is in equilibrium with CH4 (g) of standard pressure. In seawater, the dissolved methane concentration in equilibrium with gaseous methane of standard pressure is lower (Yamamoto et al. 1976), even though this has the same activity as methane in pure water

Sdiss ¼ 0:32 ðor 32%Þ Stot

(24)

Most of the substrate is therefore assimilated. The lower yields from experiments (Table 4) indicate significant energy consumption for maintenance or by uncoupling.

4.2

Requirement for Minerals (N, P, Fe)

Growth yields are not only of basic but also of practical interest because they can be used to estimate the amount of essential minerals required for oil-degrading bacteria. Since crude oil has an extremely low content of nitrogen, phosphorous, and iron, these important elements are often the limiting ones in oil biodegradation. Availability of sulfur is usually not a problem, because oil contains organic sulfur and many natural waters are rich in sulfate (seawater, 28 mM). In the environment and in cultures, microorganisms often obtain the limiting elements

0 kJ

–

H

+

0 kJ

H2O

+ 3.62 kJ

– 3.62 kJ

H

+

H2O – 3.62 kJ + + 3.62 kJ H

H2O

+ 3.62 kJ

–

H

+

0 kJ

H2O

–7

T

+ 4.72 kJ

– 4.72 kJ

H

+

H2O – 4.72 kJ + + 4.72 kJ H

H2O

–

H

+

0 kJ

H2O

HCO3 (aq) a = 1.249 (ΔGf = –591.57 kJ)

H

+

0 kJ

H2O

H2O (I), a = 1.00 , ΔGf° = –237.18 kJ T

0 kJ

CO2 (aq) a = 0.0346 (ΔGf = –394.36)

0 kJ

CO2 (g) a = 1.00 ΔG°f = –394.36 kJ

H (aq), a = 10 , ΔGf° = –39.97 kJ

+

HCO3 (aq) a = 1.00 ΔG °f = –586.85 kJ

H

+

0 kJ

H2O

+ 4.72 kJ

– 4.72 kJ

CO2 (aq) a = 0.232 (ΔGf = –389.64)

Gas phase

– 3.62 kJ

0 kJ

+ 4.72 kJ

– 4.72 kJ

Water phase pH = 7.0

0 kJ

CO2 (g) a = 6.7 (ΔG°f = –389.64 kJ)

Water phase pH = 7.0

Gas phase

+ 3.62 kJ

– 3.62 kJ

H2O +

+

H

CO2 (aq)

–

+

H

H2O

HCO3 (aq) a < 0.149

+

H

H2O

a < 0.0346

All free energie values per mol

H

H2O

Water phase pH = 7.0

Gas phase

Co2 (g) a < 1.00

Fig. 7 The three standard states (framed) of inorganic carbon (CO3 2
not included), the product of hydrocarbon oxidation. Most natural conditions will be closest to the gaseous standard state. See also remarks in legend of Appendix Fig. 6. Reactions are indicated for pH ¼ 7

HCO3 (aq) a = 4.31 (ΔGf = –583.23 kJ)

H

+

0 kJ

H2O

CO2 (aq) a = 1.00 ΔG°f = –386.02 kJ

0 kJ

CO2 (g) a = 28.9 (ΔGf = –386.02)

Natural systems

26 F. Widdel and F. Musat

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

27




as inorganic species (NH4 þ ,NO3
, H2 PO4
=HPO4 2 , Fe2þ , FeIII minerals, etc.). The above bulk formula for cell mass which considers nitrogen, hC4H8O2Ni, suggests a content of 14% N by mass; it does not consider phosphorus and iron. The extended Redfield ratio, ðCH2 OÞ106 ðNH3 Þ16 ðH3 PO4 Þ ¼ hC106 H263 O110 N16 Pi, which was derived from the originally determined molar C:N:P ratio of 106:16:1 of marine phytoplankton (Brewer et al. 1997), considers in addition phosphorus. Carbon in this formula has the bulk oxidation state as in carbohydrates, which may not very precisely reflect bacterial cell mass. “Redfield biomass” contains 6.3% N and 0.9% P by mass. With these ratios, 1 g biomass produced aerobically during complete con  sumption of 1 g (1.3 ml) hexadecane would need 0.24 g 4:5  10
3 mol NH4 Cl and   0:04 g 0:3  10
3 mol KH2 PO4 : In a marine environment with for instance 1 μM combined nitrogen and 0.06 μM phosphate, the microbial cell mass produced with 1 g hexadecane would consume the nitrogen and phosphorous from roughly 5 m3 water. However, such calculations should be applied reservedly in the study of natural hydrocarbon bioremediation. A lower in situ growth yield and N and P release from lysed cells may result in a lower than the calculated need for N and P. On the other hand, oil as a hydrophobic substrate is not distributed like soluble organic carbon in the water body but forms buoyant layers. Cells of hydrocarbon-degrading bacteria largely depend on physical contact with the oil, so that supply of biominerals by advective transport is a severely limiting factor (chapter ▶ Matrix-Hydrophobic Compound Interactions by Harms et al. in this volume). The controlled use of environmentally friendly immobilized N and P sources (as well as of iron sources that have not been considered here) that tend to stay in contact with oil may therefore be a justified method to stimulate oil degradation in eutrophic waters (chapter ▶ Bioremediation/ Biomitigation: Introduction by Ron and Rosenberg in this volume).

5

Research Needs

The application of thermodynamic data to microbial systems as a whole is a theoretical approach that is basic for the understanding of the overall catabolism of chemotrophic microorganisms (Thauer et al. 1977). Even though it is not regarded as an own field of microbiological research, the underlying formalism accompanies the study of numerous metabolic types of bacteria and may lead to the recognition of scientifically challenging questions that have not been encountered before. One of these is clearly the appropriate understanding of how microorganisms conserve energy at low chemical potential, viz., with low-energy substrates and combinations of electron donors and electron acceptors with marginal differences in their redox potential. Prominent processes of such type are anaerobic reactions involving hydrocarbons, such as the anaerobic oxidation of methane or conversion of non-methane hydrocarbons to methane by microbial consortia which even have to share the low net energy gain. Also individual enzymatic reactions in the anaerobic degradation of hydrocarbons, in particular the activating steps and intermediate energetic states (energy-rich transition states), need a deeper understanding from an energetic and kinetic point of view. There may be even open questions concerning growth yields

28

F. Widdel and F. Musat

and the efficiency of energy conservation during growth with hydrocarbons under various environmental conditions. Their examination could be relevant for the study of hydrocarbon bioremediation in oligotrophic aquatic environments.

Appendix Table 5 Hydrocarbons (methane, propane, n-hexane, and benzene as examples) and other substances as “energy carriers.” In a reaction with oxygen, liquid hydrocarbons reveal a high gravimetric energy density in comparison to many other compounds and elements (calculated for the highest oxides in their standard state). Gaseous hydrocarbons reveal a high volumetric energy density ΔG of oxidation with O2 Per mass of Per volume of substance substance   ðkJ m
3 Þ kJ kg
1

Substance Gases (101 kPa) H2
117:6  103

a


9:7  103

ΔH of oxidation with O2 Per mass of substance  kJ kg
1
141:8  103


11:7  103

CH4


51:0  10


33:5  10


55:5  10


36:4  103

C3H8


47:8  103


86:2  103 a


50:3  103


90:8  103 a

NH3b


19:9  10


13:8  10

3


22:5  10


15:6  103 a

3

3

3

3a

3


19:3  103 H2Sc Solids or liquids Li
40:4  103


26:8  103 a


23:3  103


32:4  103 a


21:6  106


43:0  103


23:0  106

B


55:2  103


135:7  106


58:8  103


144:7  106

CGraphite


32:8  10


74:4  10

3


32:8  10


74:4  106

C6H14


46:7  103


30:8  106


48:3  103


31:9  106

C6H6


41:0  10


36:0  10

3


41:8  10


37:5  106

CH3OH


21:9  103


17:5  106


22:7  103


18:1  106

CH3CH2OH


28:8  10


22:8  10

3


29:7  10


23:5  106

C6H12O6(α-DGlucose) Mg


16:0  103


25:0  106


15:6  103


24:3  106


23:4  103


40:8  106


24:8  103


43:1  106

Al


29:3  10


79:2  10

3


31:1  10


84:0  106

Si


30:5  103


71:1  106


32:4  103


75:5  106

Pwhite


21:8  10


39:7  10

3


24:1  10


43:9  106

S


11:6  103


22:7  106


12:3  103


24:1  106

Fe


6:6  10


52:3  10


7:4  10


58:3  106

3

3

3

3

3

3

6

6

6

6

6

6

3

For convenience, ideal behavior assumed. In reality, the volumetric energy density will be somewhat higher b If N2(g) is produced c If H2SO4(l) (l) is produced

Formula mass   g mol
1 16.043

16.043 30.069

44.096

58.123

58.123 72.150 72.150 86.177

86.177 100.203 114.23

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Alkanes Methane (g)

Methane (aq) Ethane (g)

Propane (g)

n-Butane (g)

2-Methylpropane (g)

n-Pentane (l)

2-Methylbutane (l) n-Hexane (l)

2-Methylpentane (l) n-Heptane

n-Octane (l)


8.11e 1.0c 1.28e 6.41e


50.72c
50.8d
50.75f
34.4g
32.82c
32.6e
32.89f
23.49c
23.6d
23.4e
17.03c
17.2d
15.7e
20.9d
18.0e
9.3d
9.21e
14.6e
3.8d
4.28e

Free energy offormationfrom the  elements, ΔGf kJ mol
1

310.23c 310.1d


126.15c
125.6d
125e
134.2d
132e
173.1c
173.5d
179e
198.7c
198.8d
199e
204e
224.4c
249.9c

269.91c 270.2d


103.85c
104.7d

(continued)

291e 328.6c 328e 361.1c

260e 296.1d 296e

294.6d 295e 262.7d

229.60c 229.1d

186.26c 186.2e

b, Entropy   S J K
1 mol
1


84.68c
83.8d


74.81c
74.9d

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Table 6 Thermodynamic properties of hydrocarbons and other compounds. Data are from other compilationsa

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . 29

Formula mass   g mol
1 114.23 114.23 114.23 142.28 170.34 184.36 198.39 226.44 240.47 254.50 70.134 84.161

28.054 42.080

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

2-Methylheptane (l) 3-Methylheptane (l) 4-Methylheptane (l) n-Decane (l)

n-Dodecane (l)

n-Tridecane (l) n-Tetradecane (l) n-Hexadecane (l)

n-Heptadecane (l) n-Octadecane (c) Cyclopentane (l)

Cyclohexane (l)

Unsaturated hydrocarbons, nonaromatic Ethene (g)

Propene (g)

Table 6 (continued)

68.15c 68.4d 62.78c 62.8d 74.8e

53.9 36.4d 36.5e 26.8e 26.7d

e

52.26c 52.5d 20.42c 20.0d 20.4e


480e
569e
105.1d
106e
156c
156.4d

219.56c 219.3d 267.05c 266.6d 227e

204.4d

497e 204.3d

523e 555e

352e 358e 350e 425.5d 426e 490.6d


255.1c
252c
252c
300.9d

3.85e 4.68e 7.8e 17.5d 17.4e 28.1d 28.4e 33.8e 38.8e 49.8h 52.2i
350.9d
352e
378e
403e
454.4h

b, Entropy   S J K
1 mol
1

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Free energy offormationfrom the  elements, ΔGf kJ mol
1

30 F. Widdel and F. Musat

56.107

56.107

56.107

26.038 78.113

92.140

106.17 106.17 106.17 106.17 120.19 128.17 142.20 142.20 154.21

1-Butene (g)

cis-2-Butene (g)

trans-2-Butene (g)

Ethyne (g) Aromatic hydrocarbons Benzene (l)

Toluene (l)

Ethylbenzene (l) o-Xylene (l)

m-Xylene (l) p-Xylene (l)

1,3,5-Trimethylbenzene (l)

Naphthalene (c)

1-Methylnaphthalene (l) 2-Methylnaphthalene (c) Biphenyl (c)

255e 246.5d 246e 252.2d 247.4d


12.5e
24.4e

189.4d 192.6d 254.2d

201d

103.9d

221d 219e

12.4d 8.08e


25.4e
24.4d
24.3e
63.4d
63.5e 78.53c 77.9d 56.3d 44.9d 99.4d

173.3c 173.4d

49.0c

124.3c 124.4d 124.5f 113.8d 110e 114.22f 120e 110.3d 111e 107.7d 110.1d

(continued)

254.8d 220d 205.9d

273.6d 273e 166.9d

296.59c 296.5d 296e 200.94c

305.71c 305.6d 307e 300.94c 300.8d


0.13c 0.1d 1.17e
6.99c
7.1d
5.70e
11.17c
11.4d
10.1e 226.73c

71.39c 71.3d 72.0e 65.95c 65.9d 67.3e 63.06c 63d 64.3e 209.2c

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . 31

Formula mass   g mol
1 178.23 178.23

32.042 32.042 46.069 46.069 60.096 60.096 60.096 60.096 74.122 74.122 74.122 242.44 108.14

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Anthracene (c)

Phenanthrene (c)

Alcohols, phenolic compounds Methanol (l)

Methanol (aq) Ethanol (l)

Ethanol (aq) 1-Propanol (l) 1-Propanol (aq) 2-Propanol (l)

2-Propanol (aq) 1-Butanol (l)

1-Butanol (aq) 2-Butanol (l) 1-Hexadecanol (c)

Benzyl alcohol (l)

Table 6 (continued)


171.84f
177.0d
98.7d
98.8e
27.5d
27.3e


166.27c
166.8d
175.39f
174.78c
174.2d
181.75f
170.6d
175.81f
180.3d
182e
185.94f
162.5d


342.6d
686.7d
684e
160.7d

225.1d 451.9d 452e 216.7d

226.4d 252e

180.6d 180e


318.1d
319e
327.3d

194.6d

160.7c 161.0d

126.8c 127.2d

207.6d 207e 211.7d 212e

b, Entropy   S J K
1 mol
1


302.6d


277.69c
277.0d


238.66c
239.1d

129.2d 128e 116.2d 113e

286.0d 268.3d

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Free energy offormationfrom the  elements, ΔGf kJ mol
1

32 F. Widdel and F. Musat

94.113

124.14 30.026

30.026 44.053 44.053 58.080 58.080 72.107 106.12 46.026

46.026 45.018

Phenol (s)

1,2-Dihydroxybenzene (s) Aldehydes, ketones Formaldehyde (g)

Formaldehyde (aq) Acetaldehyde (l)

Acetaldehyde (aq) Butyraldehyde (l) Acetone (l)

Acetone (aq) 2-Butanone (l)

Benzaldehyde (l) Carboxylic acids, carboxylates Formic acid (l)

Formic acid (aq) Formate
ðaqÞ


356.3j
351.04f
334.9j


361.35c
360e


102.53c
109.9d
111e
112.97f
130.54f
128.12c
128.3d
139.9f
127e
155.4c
155.8d
161.17f
151.4d
156e 9.4d


50.9c
50.4d
47.5e
47.6f
210.0d


424.72c
425.1d
423e
410.3j
410.3j


273.3d
279e
87.0d


247e
248.1c
242.1d


192.30c
191.8d

(continued)

163.7j 91.7j

128.95c

238.8d 241e

247e 200.4c 200.6d

160.2c 160.4d

218.77c

150.2d


361.1d
108.57c
116e

146c 144d 142e


165c
165.1d
163e

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . 33

73.071

Propionate
ðaqÞ

256.43 122.12 121.12 114.06 116.07 130.10 200.23 206.20

Palmitic acid (c) Benzoic acid (c)

Benzoate
ðaqÞ

Fumarate ðaqÞ

Succinate ðaqÞ

Methylsuccinate2
ðaqÞ

ð1
MethylpentylÞ succinate ðaqÞ

Benzylsuccinate2
ðaqÞ


521.1 to
525.4l


644.0 to
647.3l


681.6 to
685.5l


690.23f


604.21f


305.0f
245.3c
245.6f
229.3k


335.96f


23.0d
28e


882e
385.1c
385.2d

242.6d 242e

452e 167.6c

178.7c 86.6c


485.76c
486.01c


352.63f

159.8c 159.9d


484.5c
484.4d


389.9c
390.2d
392e
396.46c
369.31c
369.41f
361.08f

b Entropy 
1,
1   S J K mol

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Free energy offormationfrom the  elements, ΔGf kJ mol
1

Hydrocarbon-derived nitrogen, oxygen, sulfur, and halocompounds Methylamine (g) 31.057 32.3d 27.5e 32.065
40.0f Methylammoniumþ ðaqÞ

2


2


115.15

Hexanoate ðaqÞ




Butyrate ðaqÞ

2


60.052 59.045

Acetic acid (aq) Acetate
ðaqÞ

87.098

60.052

Acetic acid (l)




Formula mass   g mol
1

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Table 6 (continued)

34 F. Widdel and F. Musat

45.084 79.101 79.101 93.128 46.069 74.122 48.10 62.13 62.13 84.14 110.17 34.033 88.005 50.488 96.944 119.38 153.82 64.515

Ethylamine (g)

Pyridine (l) Pyridine (aq) Aniline (l)

Dimethyl ether (g)

Diethyl ether (l) Methanethiol (g)

Dimethyl sulfide (l) Ethanethiol (l) Thiophene (l)

Thiophenol (l)

Fluoromethane (g)

Tetrafluoromethane (g)

Chloromethane (g)

Dichloromethane (l) Trichloromethane (l) Tetrachlorromethane (l)

Chloroethane (g)


213.8d
222e
888.3d
862e
58.5d
58.1e
63.3e
71.2e
62.6d
68.4e
60.5d
53.0e

181.3d 177.1f 149.2d 148e
112.9d
114e
116.7d
9.9d 0.754e 5.72e
5.7e 121.2d 122e 134d

37.3d

31.3d 29.7e
184.1d
185e
279.3d
22.9d
12.4e
65.4e
73.7e 80.6d 81.7e 64.1d 62.8e
237.8d
247e
933.6d
908e
81.9d
82.0e
117e
132e
132.8d
139e
112.1d
105e


47.4d
48.5e 100.2d

(continued)

261.3d 262e 234.2d 233e 179e 203e 216.2d 214e 275.8d 275e

222.8d

222.8d

196e 207e 181.2d

253.1d 255.1d

191.4d 192e 267.1d

177.9d

289.9d

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . 35

Formula mass   g mol
1 96.104 112.56

2.0158 1.0074 1.0074 12.011 12.011 28.011 44.010 44.010 61.017 60.009 28.0134 18.038 44.013

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Fluorobenzene (l) Chlorobenzene (l)

Inorganic compounds H2(g) Hþ ðaqÞ, pH ¼ 0

Hþ ðaqÞ, pH ¼ 7

C, graphite (c) C, diamond (c)

CO (g)

CO2(g)

CO2(aq)

HCO3
ðaqÞ

CO3 2
ðaqÞ

N2(g) NH4 þ ðaqÞ

N2O (g)

Table 6 (continued)


39.97l


137.17c
137.15f
394.36c
394.39d
385.98c
386.02f
586.77c
586.85f
527.81c
527.90f 0
79.31c
79.37e 104.20c 104.18f

0 2.900c

0

ΔGf

191.61c 113.4c 219.85c

82.05c


56.9c


677.14c 0
132.51c

91.2c

213.74c 213.80d 117.6c


393.51c
393.52d
413.80c
691.99c

197.67c

5.740c 2.377c

(ΔS 0 ) 134.06

130.684c 0

0 1.895c 1.897d
110.53c

0

0 0

206e 209.2d 194e


145e 11.0d 10.6e


69.0e 89.2d 93.7e 0 0


b, Entropy   S J K
1 mol
1

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Free energy offormationfrom the  elements, ΔGf kJ mol
1

36 F. Widdel and F. Musat

62.005

31.999 18.015

32.06 34.08 34.08 33.072 96.06 18.999

NO3
ðaqÞ

O2(g)

H2O (l)

S, (α, rhombic; c)

H2S (g)

H2S (aq)

HS
ðaqÞ

SO4 2
ðaqÞ

F
ðaqÞ 35.453 54.937 114.95 86.937 55.846

Cl ðaqÞ

Mn2þ ðaqÞ

MnCO3 MnO2 Fe2þ ðaqÞ




46.006

NO2
ðaqÞ


131.23
131.3m
227.8j
228.0m
816.0m
465.1m
78.90c
78.87m c


33.56c
33.3d
27.83c
27.87d 12.08c 12.05m
744.53c
744.63f
278.79c


285.83c


237.13c
237.14d
237.178f
237.18m 0


223.3j
220.7m
889.3m
520.0m
89.10m


167.16

c

(continued)


84j
73.6m 100m 53m
137.7c

56.5c


13.8c

20.1c


909.27c
332.63c

62.08c

31.80c 32.056d 205.79c 205.7d 121c

205.138c 205.147d 69.91c 69.95d

125.2j 140m 146.4c 146.5j


20.63c
20.5d
39.7c
39.8d
17.6c

0


106.3j
104.6m
205.0c
206.7j
207.3m 0


37.2f,m
34.54j
108.74c
111.34f
110.7j 0

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . . 37

55.845 115.86 106.87 159.69 231.54

Fe3þ ðaqÞ

FeCO3 (siderite; c)

Fe(OH)3 (amorphous)

Fe2O3 (α, hematite; c)

Fe3O4 (magnetite; c)


315.9c 92.9j 105m 96j 87.40c


48.5c
748.2j
737.0m
824.8j
824.2c
824.6m
1118.4c
1115.7m


4.7c
4.60m
674.3j
666.7m
695j
699m
742.2c
742.7m
1015.4c
1012.6m

146.4c

b, Entropy   S J K
1 mol
1

Enthalpy of formation from  the  elements, ΔH f kJ mol
1

Free energy offormationfrom the  elements, ΔGf kJ mol
1

Original sources cited in the used compilations were not consulted. If a precise and rounded value is given in the compilations, the precise value is indicated here  b The absolute entropy values may be used to calculate entropy changes of reactions as well as entropies of formation, ΔSf ; the latter can be used to prove     consistency of literature data via ΔGf ¼ ΔH f
298:15ΔSf . Example of n-hexane (C6H14): ΔSf ¼ 291:6
ð6  5:74Þ
ð7  130:68Þ ¼
657:6 J K
1    mol
1 . ΔGf ¼
198:8
ð298:15 
0:6576Þ ¼
2:7 kJ mol
1 . The result is close to the value given in the literature source
3:8 kJ mol
1 c Atkins and de Paula (2006) d Dean (2004) e D’Ans and Lax (1983) f Thauer et al. (1977) g Calculated via solubility of 1.39 mol l
1 atm
1 at 25  C (from Wilhelm et al. 1977) h Via extrapolation or interpolation of the listed data i Zengler et al. (1999) j Garrels and Christ (1965) (data transformed by using 1 cal ¼ 4:1868 J) k Widdel et al. (2007) l Free energy associated with dilution of 1 mol Hþ from a ¼ 1 to a ¼ 10
7 , which is R T ln 10
7 m Stumm and Morgan (1981)

a

Formula mass   g mol
1

Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Table 6 (continued)

38 F. Widdel and F. Musat

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

39

References Anderson RT, Lovley DR (2000) Hexadecane decay by methanogenesis. Nature 404:722–723 Atkins PW, de Paula J (2006) Physical chemistry, 8th edn. Oxford University Press, Oxford Bonin P, Gilewicz M, Bertrand JC (1992) Effects of oxygen on Pseudomonas nautica grown on nalkane with or without nitrate. Arch Microbiol 157:538–545 Bordel S, Muñoz R, Díaz L, Villaverde S (2007) New insights on toluene biodegradation by Pseudomonas putida F1: influence of pollutant concentration and excreted metabolites. Appl Microbiol Biotechnol 74:857–866 Brewer PG, Goyet C, Friedrich G (1997) Direct observation of the oceanic CO2 increase revisited. Proc Natl Acad Sci U S A 94:8308–8313 Chakraborty R, Coates JD (2004) Anaerobic degradation of monoaromatic hydrocarbons. Appl Microbiol Biotechnol 64:437–446 D’Ans J, Lax E (1983) Taschenbuch für Chemiker und Physiker, Bd 2, 2. Aufl. Springer, Berlin Dean JA (2004) Lange’s handbook of chemistry, 16th edn. McGraw-Hill, New York Dinkla IJT, Gabor E, Janssen DB (2001) Effects of iron limitation on the degradation of toluene by Pseudomonas strains carrying the TOL (pWWO) plasmid. Appl Environ Microbiol 67:3406–3412 Dolfing J, Larter SR, Head IM (2008) Thermodynamic constraints on methanogenic crude oil biodegradation. ISME J 2:442–452 Einsele A (1983) Biomass from higher n-alkanes. In: Rehm H-J, Reed G (eds) Biotechnology, vol 3. Verlag Chemie, Weinheim, pp 43–81 Erickson LE (1981) Energetic yields associated with hydrocarbon fermentations. Biotechnol Bioeng 23:793–803 Ettwig KF, Shima S, van de Pas-Schoonen KT, Kahnt J, Medema MH, Op den Camp HJ, Jetten MS, Strous M (2008) Denitrifying bacteria anaerobically oxidize methane in the absence of Archaea. Environ Microbiol 10:3164–3173 Ferrer A, Erickson LE (1979) Evaluation of data consistency and estimation of yield parameters in hydrocarbon fermentations. Biotechnol Bioeng 21:2203–2233 Garrels RM, Christ CL (1965) Solutions, minerals and equilibria. Harper & Row, New York Heijnen JJ, Van Dijken JP (1992) In search of a thermodynamic description of biomass yields for the chemotrophic growth of microorganisms. Biotechnol Bioeng 39:833–858 Himo F (2002) Catalytic mechanism of benzylsuccinate synthase, a theoretical study. J Phys Chem B 106:7688–7692 Himo F (2005) C
C bond formation and cleavage in radical enzymes, a theoretical perspective. Biochim Biophys Acta 1707:24–33 Jackson BE, McInerney MJ (2002) Anaerobic microbial metabolism can proceed close to thermodynamic limits. Nature 415:454–456 Jones DM, Head IM, Gray ND, Adams JJ, Rowan AK, Aitken CM, Bennett B, Huang H, Brown A, Bowler BF, Oldenburg T, Erdmann M, Larter SR (2008) Crude-oil biodegradation via methanogenesis in subsurface petroleum reservoirs. Nature 451:176–180 Krüger M, Meyerdierks A, Glöckner FO, Amann R, Widdel F, Kube M, Reinhardt R, Kahnt J, Thauer RK, Shima S (2003) A conspicuous nickel protein in microbial mats that oxidise methane anaerobically. Nature 426:878–881 Leak DJ, Dalton H (1985) Growth yields of methanotrophs. Appl Microbiol Biotechnol 23:477–481 McMillen DF, Golden DM (1982) Hydrocarbon bond dissociation energies. Annu Rev Phys Chem 33:493–532 Nauhaus K, Boetius A, Krüger M, Widdel F (2002) In vitro demonstration of anaerobic oxidation of methane coupled to sulphate reduction in sediment from a marine gashydrate area. Environ Microbiol 4:296–305

40

F. Widdel and F. Musat

Nauhaus K, Albrecht M, Elvert M, Boetius A, Widdel F (2007) In vitro cell growth of marine archaeal-bacterial consortia during anaerobic oxidation of methane with sulphate. Environ Microbiol 9:187–196 Pfennig N, Biebl H (1976) Desulfuromonas acetoxidans gen. nov. and sp. nov., an new anaerobic, sulfur-reducing, acetate-oxidizing bacterium. Arch Microbiol 110:3–12 Pirt SJ (1965) The maintenance energy of bacteria in growing cultures. Proc R Soc Lond 163B:224–231 Rabus R, Widdel F (1995) Anaerobic degradation of ethylbenzene and other aromatic hydrocarbons by new denitrifying bacteria. Arch Microbiol 163:96–103 Rabus R, Nordhaus R, Ludwig W, Widdel F (1993) Complete oxidation of toluene under strictly anoxic conditions by a new sulfate-reducing bacterium. Appl Environ Microbiol 59:1444–1451 Rabus R, Wilkes H, Behrends A, Armstroff A, Fischer T, Pierik AJ, Widdel F (2001) Anaerobic initial reaction of n-alkanes: evidence for (1-methylpentyl)succinate as initial product and for involvement of an organic radical in the metabolism of n-hexane in a denitrifying bacterium. J Bacteriol 183:1707–1715 Reardon KF, Mosteller DC, Bull Rogers JD (2000) Biodegradation kinetics of benzene, toluene, and phenol as single and mixed substrates for Pseudomonas putida F1. Biotechnol Bioeng 69:385–400 Russell JB (2007) The energy spilling reactions of bacteria and other organisms. J Mol Microbiol Biotechnol 13:1–11 Schink B (1997) Energetics of syntrophic cooperation in methanogenic degradation. Microbiol Mol Biol Rev 61:262–280 Schink B (2002) Anaerobic digestion: concepts, limits and perspectives. Water Sci Technol 45:1–8 Shima S, Thauer RK (2005) Methyl-coenzyme M reductase and the anaerobic oxidation of methane in methanotrophic Archaea. Curr Opin Microbiol 8:643–648 So CM, Young LY (1999) Isolation and characterization of a sulfate-reducing bacterium that anaerobically degrades alkanes. Appl Environ Microbiol 65:2969–2976 Stouthamer AH (1988) Bioenergetics and yields with electron acceptors other than oxygen. In: Erickson LE, Fung DY-C (eds) Handbook of anaerobic fermentations. Marcel Dekker, New York, pp 345–437 Stumm W, Morgan JJ (1981) Aquatic chemistry, 2nd edn. John Wiley & Sons, New York Tan NC, van Doesburg W, Langenhoff AA, Stams AJ (2006) Benzene degradation coupled with chlorate reduction in a soil column study. Biodegradation 17:113–119 Tempest DW, Neijssel OM (1984) The Status of YATP and maintenance energy as biologically interpretable phenomena. Annu Rev Microbiol 38:459–486 Thauer RK, Shima S (2008) Methane as a fuel for anaerobic microorganisms. Annu NY Acad Sci 1125:158–170 Thauer RK, Jungermann K, Decker K (1977) Energy conservation in chemotrophic anaerobic bacteria. Bacteriol Rev 41:100–180 Tissot BP, Welte DH (1984) Petroleum formation and occurrence. Springer, Berlin van Dijken JP, Harder W (1975) Growth yields of microorganisms on methanol and methane. A theoretical study. Biotechnol Bioeng 17:15–30 Wagner F, Kleemann T, Zahn W (1969) Microbial transformations of hydrocarbons. II. Growth constants and cell composition of microbial cells derived from n-alkanes. Biotechnol Bioeng 11:393–408 Widdel F, Musat F, Knittel K, Galushko A (2007) Anaerobic degradation of hydrocarbons with sulphate as electron acceptor. In: Barton LL, Hamilton WA (eds) Sulphate-reducing bacteria. Cambridge University Press, Cambridge, pp 265–303 Wilhelm E, Battino R, Wilcock RJ (1977) Low-pressure solubility of gases in liquid water. Chem Rev 77:219–262 Wodzinski RS, Johnson MJ (1968) Yields of bacterial cells from hydrocarbons. Appl Microbiol 16:1886–1891

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon. . .

41

Yamamoto S, Alcauskas JB, Crozier TE (1976) Solubility of methane in distilled water and seawater. J Chem Eng Data 21:78–80 Zengler K, Richnow HH, Roselló-Mora R, Michaelis W, Widdel F (1999) Methane formation from long-chain alkanes by anaerobic microorganisms. Nature 401:266–269

Enzymes for Aerobic Degradation of Alkanes in Yeasts Ryouichi Fukuda and Akinori Ohta

Abstract

A wide variety of yeasts can utilize n-alkanes as sole carbon and energy sources. The degradation pathways of n-alkanes in yeasts and the enzymes associated with these pathways have been studied intensively in the ascomycetous yeasts, Candida tropicalis, Candida maltosa, and Yarrowia lipolytica, for biotechnological applications, such as conversion of n-alkanes to proteins or useful compounds, as well as for elucidating the metabolism of hydrophobic substrates by fungi. Here, we describe the aerobic degradation pathway of n-alkanes in yeasts and the enzymes that catalyze the reactions involved in the degradation. In n-alkaneassimilating yeasts, incorporated n-alkanes are hydroxylated to fatty alcohols by cytochromes P450 of the CYP52 family in the endoplasmic reticulum (ER). Fatty alcohols are oxidized in the ER or the peroxisome to fatty aldehydes and finally to fatty acids, which are then activated to acyl-CoAs and metabolized by β-oxidation or used for lipid synthesis.

Contents 1 2 3 4

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Uptake of n-Alkanes by Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Terminal Hydroxylation of n-Alkanes by Cytochrome P450 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Oxidation of Fatty Alcohols to Fatty Aldehydes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 4 4 8

R. Fukuda (*) Department of Biotechnology, The University of Tokyo, Tokyo, Japan e-mail: [email protected] A. Ohta Department of Biological Chemistry, College of Bioscience and Biotechnology, Chubu University, Kasugai, Aichi, Japan e-mail: [email protected] # Springer International Publishing AG 2017 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-319-39782-5_7-1

1

2

R. Fukuda and A. Ohta

5 Oxidation of Fatty Aldehydes to Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 6 Activation and Utilization of Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

1

Introduction

n-Alkanes are common compounds in nature, and a variety of microorganisms, including bacteria, yeasts, and filamentous fungi, can utilize these compounds as their sole carbon and energy sources. The assimilation of n-hexadecane has been assessed in ~700 yeast species among the 1,270 yeasts listed in The Yeasts: A Taxonomic Study, and ~180 of these 700 species have been shown to have the ability to assimilate n-hexadecane (Kurtzman et al. 2011). These ~180 yeasts belong to 28 genera of ascomycetous and basidiomycetous yeasts including Candida, Debaryomyces, Metschnikowia, Yarrowia, and Cryptococcus (Fig. 1). Neither the

Candida tropicalis Candida maltosa Candida parapsilosis 0.02 963 Lodderomyces elongisporus Candida dubliniensis 1000 Candida albicans Scheffersomyces stipitis Meyerozyma guilliermondii Debaryomyces hansenii Millerozyma farinosa Saccharomyces cerevisiae Yarrowia lipolytica Candida apicola Starmerella bombicola Metschnikowia pulcherrima Schizosaccharomyces pombe Cryptococcus_musci 987

971

1000 365 991 598 979 477 528 841 745

1000

Fig. 1 Phylogenetic tree of n-alkane-assimilating yeasts. Phylogenetic tree of D1/D2 regions of 26S ribosomal DNA of the yeasts that can assimilate n-alkanes and the model yeasts, Saccharomyces cerevisiae and Schizosaccharomyces pombe, was constructed using ClustalW (DDBJ, v2.1) and drawn using NJplot. The scale bar indicates 0.02 substitutions per site. The bootstrap values by 1000 repetitions are indicated. The accession numbers of sequences of D1/D2 regions from GenBank are as follows: Candida albicans (U45776), Candida apicola (U45703), Candida dubliniensis (U57685), C. maltosa (U45745), Candida parapsilosis (U45754), Candida tropicalis (U45749), Cryptococcus musci (KC585415), Debaryomyces hansenii (U45808), Lodderomyces elongisporus (U45763), Metschnikowia pulcherrima (U45736), Meyerozyma guilliermondii (U45709), Millerozyma farinosa (U45739), Saccharomyces cerevisiae (U44806), Scheffersomyces stipitis (U45741), Schizosaccharomyces pombe (U40085), Starmerella bombicola (U45705), and Yarrowia lipolytica (U40080)

Enzymes for Aerobic Degradation of Alkanes in Yeasts

3

n-Alkane

Plasma membrane

ER

CYP52-family P450

Peroxisome CH2OH

CH2OH

FADH

FAOD CHO

CHO

FALDH

FALDH

COOH

ACS Acyl-CoA

COOH

ACS Acyl-CoA β-Oxidation

Lipids Acetyl-CoA Fig. 2 Proposed pathway of n-alkane metabolism in yeasts. Incorporated n-alkanes are hydroxylated to fatty alcohols by cytochromes P450 of the CYP52 family. Fatty alcohols are oxidized to fatty aldehydes by fatty alcohol dehydrogenase (FADH) in the ER or fatty alcohol oxidase (FAOD) in the peroxisome. Fatty aldehydes are oxidized to fatty acids by fatty aldehyde dehydrogenase (FALDH) in the ER or the peroxisome. Fatty acids are activated to acyl-CoAs by acyl-CoA synthetase (ACS) and are metabolized through β-oxidation pathway in the peroxisome or utilized for membrane or storage lipid synthesis

model yeast Saccharomyces cerevisiae nor Schizosaccharomyces pombe, however, can assimilate n-alkanes. The metabolic pathway of n-alkanes has been studied intensively in Candida tropicalis (Tanaka and Fukui 1989), Candida maltosa (Mauersberger et al. 1996), and Yarrowia lipolytica (Barth and Gaillardin 1996; Barth and Gaillardin 1997; Fickers et al. 2005; Fukuda 2013; Fukuda and Ohta 2013; Nicaud 2012) and has attracted considerable attentions owing to its involvement in the production of singlecell protein (SCP) and other useful materials, including long-chain dicarboxylic acids and tricarboxylic acid (TCA) cycle intermediates, from n-alkanes (Fickers et al. 2005; Tanaka and Fukui 1989). In order to establish and improve systems for the production of useful materials from n-alkanes using yeasts, the n-alkane metabolic pathway and associated enzymes must be understood in detail. In C. tropicalis, C. maltosa, and Y. lipolytica, incorporated n-alkanes are sequentially oxidized to fatty acids, which are then metabolized via the β-oxidation pathway in the peroxisome or utilized for the synthesis of membrane or storage lipids (Fig. 2). This chapter discusses the pathway of n-alkane oxidation to fatty acids and the subsequent activation of fatty acids to acyl-CoAs, as well as the enzymes catalyzing these reactions in yeasts. The metabolism and utilization of fatty acids in yeasts will be described in another chapter.

4

2

R. Fukuda and A. Ohta

Uptake of n-Alkanes by Yeasts

n-Alkanes exhibit poor solubility in water, and it has been proposed that yeasts facilitate the uptake of n-alkanes by two mechanisms, which are not mutually exclusive. In the first mechanism, reported for some species of n-alkane-assimilating yeasts, bioemulsifiers or biosurfactants are secreted by yeasts to solubilize n-alkanes (Barth and Gaillardin 1996; Tanaka and Fukui 1989): Y. lipolytica secretes a 28-kDa emulsifier, named liposan, composed of 83% carbohydrate and 17% protein (Cirigliano and Carman 1984, 1985), while Starmerella bombicola and Candida apicola produce the biosurfactant, sophorolipid (Van Bogaert et al. 2007). The physiological roles of these bioemulsifiers and biosurfactants in the uptake of n-alkanes, however, remain to be examined. In the second mechanism, n-alkaneassimilating yeasts adhere to n-alkane droplets to enhance the uptake of n-alkanes. Protrusions or slime-like outgrowths have been observed on the cell surface of C. tropicalis, C. maltosa, and Y. lipolytica cultured in the presence of n-alkanes (Kim et al. 2000; Mauersberger et al. 1996; Osumi et al. 1975; Tanaka and Fukui 1989). The slime-like outgrowths have been shown to reach the cell membrane through electron-dense channels in C. tropicalis, while the endoplasmic reticulum (ER) was found close to the cell membrane beneath such channels (Osumi et al. 1975). From these observations, it was suggested that n-alkanes are attached to the protrusions or slime-like outgrowths and are thereby transported through the channels to the ER, where they are hydroxylated to fatty alcohols (Sect. 3). The molecular structures of these protrusions or slime-like outgrowths and their roles in n-alkane assimilation, however, are not well understood. Two models have been proposed for the uptake of n-alkanes by yeasts: one is a passive, diffusion-like mechanism facilitated by the hydrophobic properties of nalkanes, while the other is an active, energy-dependent mechanism mediated by transporter proteins. Although the molecular mechanism of n-alkane uptake by yeasts is still unclear, the uptake of 14C-labeled n-hexadecane by Y. lipolytica was shown to be upregulated by the incubation with n-decane and inhibited by KCN and 2,4-dinitrophenol (Bassel and Mortimer 1985). Mutations in the 16 loci were also shown to significantly reduce n-hexadecane uptake in Y. lipolytica (Bassel and Mortimer 1985). These results support the model of n-alkane incorporation across the plasma membrane by one or more energy-dependent transporters. A Y. lipolytica strain with an insertion mutation in ABC1 encoding an ATP-binding cassette (ABC) transporter was shown to exhibit defective growth on n-hexadecane, but not on n-decane; however, the involvement of Abc1p in the uptake of n-hexadecane remains to be investigated (Thevenieau et al. 2007).

3

Terminal Hydroxylation of n-Alkanes by Cytochrome P450

Incorporated n-alkanes are transported from the plasma membrane to the ER, where they are hydroxylated to fatty alcohols by cytochromes P450 (P450s) belonging to the CYP52 family using molecular oxygen (Fig. 2) (Nelson 2009). Genes encoding

Enzymes for Aerobic Degradation of Alkanes in Yeasts

5

CYP52-family P450s have been identified in various n-alkane-assimilating yeasts, including C. tropicalis (Sanglard et al. 1987; Seghezzi et al. 1992; Seghezzi et al. 1991), C. maltosa (Ohkuma et al. 1991, 1995), Candida albicans (Kim et al. 2007; Panwar et al. 2001), Candida dubliniensis, Candida parapsilosis, Debaryomyces hansenii (Yadav and Loper 1999), Lodderomyces elongisporus, Meyerozyma guilliermondii, Scheffersomyces stipitis, Starmerella bombicola (Van Bogaert et al. 2009), and Y. lipolytica (Fig. 3) (Fickers et al. 2005; Hirakawa et al. 2009; Iida et al. 1998, 2000). A striking feature of the CYP52-family P450s in these yeasts is that they exist as multiple paralogs of the CYP52-family P450 gene. Eight genes encoding CYP52-family P450s have been identified in C. tropicalis and C. maltosa, and Y. lipolytica has twelve CYP52-family P450 genes. Furthermore, C. albicans, C. dubliniensis, C. parapsilosis, D. hansenii, L. elongisporus, M. guilliermondii, and S. stipitis have more than four genes encoding CYP52family P450s. During the evolution of n-alkane-assimilating yeasts, the CYP52family P450 genes may have multiplicated and functionally diversified to efficiently metabolize n-alkanes and their metabolites or to detoxify them (see following paragraph). The CYP52-family P450s of C. maltosa are encoded by ALK1–ALK8, and quadruple deletion of ALK1, ALK2, ALK3, and ALK5 was shown to cause defects in the utilization of n-alkanes for growth (Ohkuma et al. 1998). In Y. lipolytica, the CYP52-family P450s are encoded by ALK1–ALK12, and a mutant in which all twelve ALK genes are deleted completely lost the ability to grow on n-alkanes (Takai et al. 2012). These results clearly indicate the essential roles of the CYP52family P450s in the assimilation of n-alkanes (Fig. 4). The Y. lipolytica deletion mutant of twelve ALK genes was furthermore shown not to be able to grow on ndecane even in the presence of glucose, suggesting that n-decane is toxic to the yeast cells and that the CYP52-family P450s are involved in the detoxification of n-decane (Takai et al. 2012). Substrate specificities have been studied in a subset of the CYP52-family P450s of C. tropicalis, C. maltosa, C. albicans, and Y. lipolytica, and it has been shown that they have distinct substrate preferences (Fig. 3) (Eschenfeldt et al. 2003; Iwama et al. 2016; Kim et al. 2007; Ohkuma et al. 1998; Zimmer et al. 1996). In accordance with their involvement in n-alkane metabolism, some CYP52-family P450s were shown to preferentially hydroxylate n-alkanes, while, in contrast, a subset of CYP52-family P450s preferred hydroxylation of the ω-terminal end (ω-hydroxylation) of fatty acids. Some CYP52-family P450s hydroxylated both n-alkanes and the ω-terminal end of fatty acids. The amino acid sequences of Alk4 (CYP52A7) and Alk5 (CYP52A8) of C. tropicalis (Seghezzi et al. 1992); Alk5 (CYP52A9), Alk7 (CYP52A10), and Alk8 (CYP52A11) of C. maltosa (Zimmer et al. 1996, 1998); and CYP52A21 of C. albicans (Kim et al. 2007) – all of which were shown to preferentially hydroxylate the ω-terminal ends of fatty acids – showed significant sequence similarities (Fig. 3). Interestingly, it has been reported that CYP52A3 of C. maltosa catalyzes the oxidation of fatty alcohol and fatty aldehyde in addition to hydroxylating n-alkane and the ω-terminus of fatty acids (Scheller et al. 1998).

6

R. Fukuda and A. Ohta

1000 0.1

1000

416

709

1000 1000 392 509 575 306 468

745

809

618 465 996 272 1000 564 473

538

1000 998 297

1000

1000 149 475 876 733 701 460 266 574 550

985

1000

1000

1000

1000

504

1000

1000

995

CYP52A1 (Ct Alk1) CYP52A4 (Cm Alk3) CYP52A22 (Ca) CYP52A2 (Ct Alk2) CYP52A23 (Ca) CYP52A5 (Cm Alk2) CYP52A6 (Ct Alk3) CYP52A24 (Ca) CYP52A3 (Cm Alk1) CYP52A44 (Dh) CYP52A53 (Ss) CYP52A54 (Ss) CYP52A56 (Ss) CYP52A43 (Dh) CYP52A57 (Ss) CYP52A55 (Ss) CYP52A45 (Dh) CYP52A47 (Dh) CYP52A46 (Dh) CYP52A7 (Ct Alk4) CYP52A8 (Ct Alk5) CYP52A9 (Cm Alk5) CYP52A21 (Ca) CYP52A11 (Cm Alk8) CYP52A10 (Cm Alk7) CYP52D2 (Ct) CYP52D1 (Cm Alk4) CYP52F1 (Yl Alk1) CYP52F10 (Yl Alk9) CYP52F2 (Yl Alk2) CYP52F11 (Yl Alk10) CYP52F3 (Yl Alk3) CYP52F9 (Yl Alk12) CYP52F4 (Yl Alk4) CYP52F5 (Yl Alk5) CYP52F7 (Yl Alk7) CYP52F6 (Yl Alk6) CYP52F8 (Yl Alk8) CYP52C1 (Ct Alk7) CYP52C3 (Ca) CYP52C2 (Cm Alk6) CYP52B1 (Ct Alk6) CYP52S1 (Yl Alk11) CYP52E3 (Sb) CYP52M1 (Sb) CYP52N1 (Sb) CYP51 (Yl)

AF A AF A A

F F F F F F AF AF AF A AF F F F AF F

F F

Fig. 3 Phylogenetic tree of the CYP52-family P450s in yeasts. Phylogenetic tree of the CYP52family P450s of n-alkane-assimilating yeasts was constructed using ClustalW (DDBJ, v2.1) and drawn using NJplot. CYP51 of Y. lipolytica was used as an out-group. The scale bar indicates 0.05 substitutions per site. The bootstrap values by 1000 repetitions are indicated. The accession numbers of sequences from UniProtKB are as follows: CYP52A21 (Q59K96), CYP52A22 (Q5AAH7), CYP52A23 (Q5AAH6), CYP52A24 (Q5A8M1), CYP52C3 (Q5AGW4), CYP52A3 (P16496), CYP52A5 (Q12581), CYP52A4 (P16141), CYP52A9 (Q12586), CYP52A10 (Q12588), CYP52A11 (Q12589), CYP52C2 (Q12587), CYP52D1 (Q12585), CYP52A1 (P10615), CYP52A2 (P30607), CYP52A6 (P30608), CYP52A7 (P30609), CYP52A8 (P30610), CYP52B1 (P30611), CYP52C1 (P30612), CYP52D2 (Q874J0), CYP52A43 (Q6BVP2), CYP52A44 (Q6BVH7), CYP52A45 (Q6BNW0), CYP52A46 (Q6BNV9), CYP52A47 (Q6BNV8), CYP52A53 (A3LRT5), CYP52A54 (A3LR60), CYP52A55 (A3LS01), CYP52A56 (A3LZV9), CYP52A57 (A3LSP0), CYP52E3 (B8QHP3), CYP52M1 (B8QHP1), CYP52N1 (B8QHP5), CYP52F1 (O74127), CYP52F2 (O74128), CYP52F3 (O74129), CYP52F4 (O74130), CYP52F5 (O74131),

Enzymes for Aerobic Degradation of Alkanes in Yeasts

7

In Y. lipolytica, eleven CYP52-family P450s belong to the CYP52F subfamily and one belongs to the CYP52S subfamily. The CYP52F-subfamily P450s, Alk1p–Alk10p and Alk12p, appear to constitute a monophyletic clade in the phylogenetic tree of the CYP52-family P450s (Fig. 3). The CYP52F-subfamily P450s of Y. lipolytica can also be classified into four groups: P450s with significant n-alkanehydroxylating activity, P450s with significant hydroxylating activity for the ω-terminus of dodecanoic acid, P450s with significant hydroxylating activity for both n-alkanes and dodecanoic acid, and P450s with faint or no oxidizing activity for these substrates. Alk1p, Alk9p, Alk2p, and Alk10p, which have been shown to exhibit substrate preferences for n-alkanes, share significant sequence similarities, while Alk5p and Alk7p, which have ω-hydroxylation activities to dodecanoic acid, are structurally similar (Iwama et al. 2016). Alk proteins that were shown to catalyze the oxidation of n-alkanes showed distinct preferences for different n-alkane chain lengths. Alk1p and Alk3p were shown to hydroxylate n-alkanes of various carbon numbers. Alk10p, too, oxidized n-alkanes of a wide range of lengths, but was shown to preferentially oxidize shorter-chain n-alkanes. Alk2p, Alk6p, and Alk9p, on the Fig. 4 n-Alkane metabolism in Y. lipolytica. See text for details

n-Alkane

ER Alk proteins

Plasma membrane Peroxisome CH2OH

CH2OH

Adh? Fadh? Alk proteins?

Fao1

CHO

CHO

Hfd1, Hfd2A, Hfd4?

Hfd1, Hfd2A, Hfd2B, Hfd3

Faa1

Fat1

COOH

Acyl-CoA

Lipids

COOH

Acyl-CoA β-Oxidation

Acetyl-CoA

ä Fig. 3 (continued) CYP52F6 (O74132), CYP52F7 (O74133), CYP52F8 (O74134), CYP52F10 (A0A0K2S2A7), CYP52F11 (Q6CDW4), CYP52S1 (Q6CCE5), CYP52F9 (Q6CGD9), and CYP51 of Y. lipolytica (Q6CFP4). Species are indicated in parentheses as follows: C. albicans (Ca), C. maltosa (Cm), C. tropicalis (Ct), D. hansenii (Dh), S. stipitis (Ss), S. bombicola (Sb), and Y. lipolytica (Yl). P450s that were shown to catalyze the oxidation of n-alkanes or ω-termini of fatty acids are indicated as A or F, respectively

8

R. Fukuda and A. Ohta

other hand, preferred longer-chain n-alkanes. In Y. lipolytica, the genes encoding CYP52-family P450s are likely to have multiplicated after diverging from ancestral n-alkane-assimilating yeasts carrying one or a small number of genes encoding CYP52-family P450s.

4

Oxidation of Fatty Alcohols to Fatty Aldehydes

In n-alkane-assimilating yeasts, fatty alcohols are thought to be oxidized to fatty aldehydes by NAD+- or NADP+-dependent fatty alcohol dehydrogenase (FADH) in the ER or by H2O2-producing fatty alcohol oxidase (FAOD) in the peroxisome (Fig. 2) (Barth and Gaillardin 1996; Fickers et al. 2005; Fukuda 2013; Fukuda and Ohta 2013; Mauersberger et al. 1996; Tanaka and Fukui 1989). Such FAODs have been detected and characterized in several n-alkane-assimilating yeasts including C. tropicalis, C. maltosa, C. parapsilosis, and Y. lipolytica (Dickinson and Wadforth 1992; Kemp et al. 1994; Mauersberger et al. 1992). The FAOD-coding genes FAOT, FAO1, and FAO2 have been identified in C. tropicalis (Cheng et al. 2005; Eirich et al. 2004; Vanhanen et al. 2000). A FAOT deletion mutant exhibited defective growth on n-octadecane but not on shorter-chain n-alkanes or fatty acids, suggesting that FAOD encoded by FAOT is involved in the oxidation of fatty alcohol that is produced in the metabolism of n-octadecane and that one or more other enzymes are involved in the oxidation of shorter-chain n-alkanes (Cheng et al. 2005). In the genome sequence of Y. lipolytica, eight alcohol dehydrogenase genes, ADH1–ADH7 and FADH, and a fatty alcohol oxidase gene, FAO1, were identified (Gatter et al. 2014). Among these genes, a triple deletion mutant of ADH1, ADH3, and FAO1 showed severely defective growth on 1-dodecanol and 1-tetradecanol, but not on dodecanoic acid or tetradecanoic acid, suggesting that Adh1p, Adh3p, and Fao1p are involved in the assimilation of exogenous fatty alcohols (Iwama et al. 2015). Microscopic observation suggested that Fao1p localizes in the peroxisome. Adh1p and Adh3p are cytosolic proteins, but substantial amounts of these proteins were recovered in the membrane fraction of cell extracts, raising the possibility that Adh1p and Adh3p transiently localize to the membranes, possibly to the ER. A deletion mutant of ADH1–ADH7, FADH, and FAO1 exhibited slightly defective growth on n-decane and n-dodecane, but not on longer-chain n-alkanes. These results imply that any one or more of the enzymes encoded by these genes catalyze the oxidation of fatty aldehydes produced in the metabolism of shorter-chain n-alkanes (Fig. 4), but that one or more other enzymes are also involved in the oxidation of fatty alcohols derived from n-alkanes.

5

Oxidation of Fatty Aldehydes to Fatty Acids

In n-alkane-assimilating yeasts, fatty aldehydes generated during n-alkane metabolism are oxidized to fatty acids by fatty aldehyde dehydrogenase (FALDH) in the ER or in the peroxisome (Fig. 2). The model yeast S. cerevisiae has a single FALDH-

Enzymes for Aerobic Degradation of Alkanes in Yeasts

9

coding gene, HFD1. S. cerevisiae Hfd1 and its mammalian ortholog ALDH3A2 are involved in the conversion of hexadecenal to hexadecenoic acid during the degradation of sphingosine-1-phosphate, a metabolite of sphingolipids and a second messenger involved in various cellular processes (Nakahara et al. 2012). ALDH3A2 also plays a role in the degradation of phytanic acid in the peroxisome (Ashibe et al. 2007; Verhoeven et al. 1998) as well as in protection from oxidative stress associated with lipid peroxidation (Demozay et al. 2004). Mutations in ALDH3A2 have been shown to cause Sjögren-Larsson syndrome (De Laurenzi et al. 1996). Hfd1 and ALDH3A2 belong to a superfamily of NAD(P)+-dependent aldehyde dehydrogenases (Sophos et al. 2001). In n-alkane-assimilating yeasts, FALDH activities have been reported in Candida intermedia, C. tropicalis, and Y. lipolytica (Liu and Johnson 1971; Ueda and Tanaka 1990; Yamada et al. 1980). Genes encoding FALDHs that are involved in the assimilation of n-alkanes were identified in Y. lipolytica (Iwama et al. 2014). The genome of Y. lipolytica contains four orthologs, HFD1–HFD4, of S. cerevisiae HFD1 and mammalian ALDH3A2. A Y. lipolytica mutant lacking all four HFD genes did not grow on n-alkanes of 12–18 carbons and showed severe growth defects on n-alkanes of 10 and 11 carbons. The expression of any one of these genes, however, restored the growth of the deletion mutant on n-alkanes. Furthermore, bacterially produced Hfd proteins exhibited dehydrogenase activities to dodecanal and tetradecanal in vitro. Fluorescence microscopic analysis suggested that Hfd1p localizes to the ER and the peroxisome and that Hfd3p localizes to the peroxisome. Two HFD2 transcript variants, which encode Hfd2Bp containing a peroxisomal targeting signal 1 (PTS1)-like sequence at its C-terminus and Hfd2Ap without PTS1, were generated from HFD2. Hfd2Ap has been suggested to localize in the ER and the peroxisome, while Hfd2Bp localizes to the peroxisome. These results imply that Hfd proteins are involved in the oxidation of fatty aldehydes produced during metabolism of n-alkanes in the ER and the peroxisome (Fig. 4) (Iwama et al. 2014); however, growth of the quadruple deletion mutant of HFD genes on dodecanal or tetradecanal indicated that one or more other enzymes are involved in the assimilation of exogenous fatty aldehydes. The n-alkane-assimilating yeasts, C. tropicalis, C. albicans, C. dubliniensis, C. parapsilosis, L. elongisporus, and M. guilliermondii, have multiple FALDH genes. Although involvement of these FALDH genes in the assimilation of n-alkanes remains to be examined, it is possible that they were multiplicated to efficiently degrade fatty aldehydes produced during the metabolism of n-alkanes.

6

Activation and Utilization of Fatty Acids

Fatty acids play a critical role as hydrophobic moieties in lipid molecules that constitute biological membranes or are used as energy and carbon sources via β-oxidation. Fatty acids are also involved in a variety of cellular processes as precursors of signaling molecules and hormones as well as by acylation of proteins. Fatty acids are used for these processes in the forms of acyl-CoAs, which are

10

R. Fukuda and A. Ohta

synthesized from fatty acids and coenzyme A by acyl-CoA synthetase (ACS). Multiple ACS isozymes are encoded by the genomes of eukaryotes including yeasts, and some ACS isozymes exhibit distinct substrate specificities and subcellular distributions (Black and DiRusso 2007; Soupene and Kuypers 2008; Watkins and Ellis 2012). Y. lipolytica has five ACSs, Faa1p and Fat1p–Fat4p, and ten ACS-like enzymes, Aa11p–Aal10p (Dulermo et al. 2014, 2016; Tenagy et al. 2015; Wang et al. 2011). A deletion mutant of FAT1 exhibited severely defective growth on n-decane of 10 and 12 carbons and partially defective growth on 14 and 16 carbons. The FAA1 deletion mutant exhibited retarded growth on n-alkane of 16 carbons, while deletion mutants of other ACS genes did not show any defects on n-alkanes. In addition, a double deletion mutant of FAT1 and FAA1 showed severe growth defects on nalkanes of 10–18 carbons (Tenagy et al. 2015). These results suggest that Faa1p and Fat1p play critical roles in the activation of fatty acids produced during n-alkane assimilation (Fig. 4). The wild-type strain of Y. lipolytica was shown to grow in the presence of cerulenin, an inhibitor of fatty acid synthesis, when n-octadecane was supplemented, suggesting that the stearic acid produced by the oxidation of noctadecane is activated to stearoyl-CoA and that stearoyl-CoA or its derivatives support the growth of Y. lipolytica. The FAA1 deletion mutant, however, did not grow in the presence of cerulenin and n-octadecane, suggesting that FAA1 is involved in the activation of fatty acids derived from n-alkanes for essential cellular processes including membrane lipid synthesis. Fluorescent microscopic observation and fractionation analysis of cell extracts suggested that Fat1p localizes in the peroxisome, in agreement with the presence of a PTS1-like sequence at its C-terminus, while Faa1p localizes in the cytosol and to membranes. Roles of ACS isozymes of other n-alkane-assimilating yeasts in the metabolism of n-alkanes remain to be elucidated.

7

Research Needs

The oxidation pathway of n-alkanes to fatty acids in yeasts has been studied primarily in C. tropicalis, C. maltosa, and Y. lipolytica. Apart from the enzymes that are involved in the oxidation of fatty alcohols produced through the hydroxylation of n-alkanes, the enzymes that catalyze the steps in the oxidation of n-alkanes to fatty acids have been identified. As mentioned already, CYP52A3 of C. maltosa reportedly catalyzes the cascade of the sequential oxidation of n-hexadecane to hexadecanoic acid (Scheller et al. 1998). Accordingly, the CYP52-family P450s may catalyze the sequential oxidation of fatty alcohols to fatty acids in other n-alkane-assimilating yeasts. The aerobic degradation of n-alkanes and the enzymes involved in this degradation have been studied in only a limited subset of ascomycetous yeasts (C. tropicalis, C. tropicalis, and Y. lipolytica), while this process in other yeasts, particularly basidiomycetous yeasts, remains to be characterized. The metabolism of n-alkanes is regulated at the transcriptional level in C. tropicalis, C. maltosa, and Y. lipolytica. Transcription of genes encoding a subset of enzymes involved in the n-alkane metabolism is upregulated in the presence of

Enzymes for Aerobic Degradation of Alkanes in Yeasts

11

n-alkanes in these yeasts (Endoh-Yamagami et al. 2007; Hirakawa et al. 2009; Iida et al. 1998, 2000; Kobayashi et al. 2013, 2015; Mori et al. 2013; Ohkuma et al. 1991, 1995; Sanglard et al. 1987; Yamagami et al. 2004). Details of the transcriptional regulation of n-alkane metabolic genes in yeasts are described and discussed in other chapters. Two organelles, the ER and the peroxisome, are involved in the metabolism of n-alkanes in yeasts. Fundamental and critical questions that remain to be answered about these organelles in the context of n-alkane metabolism in yeasts are (1) how n-alkanes are imported into cells and transported to the ER and (2) how hydrophobic metabolites, fatty alcohols, fatty aldehydes, and fatty acids are transported from the ER to the peroxisome during n-alkane metabolism. C. maltosa and C. tropicalis produce dicarboxylic acids from n-alkanes and excrete them into culture medium (Arie et al. 2000; Tanaka and Fukui 1989). In a dicarboxylic acid-hyperproducing mutant of C. maltosa, the transcription of a CmCDR1 gene encoding an ABC transporter is highly activated in the later phase of culture on n-dodecane (Sagehashi et al. 2013). It would be of interest to test whether overexpression of CmCDR1 improves the efficiency of dicarboxylic acid production from n-alkane. The uptake, intracellular transport, and excretion of lipophilic compounds in eukaryotic cells are important and fundamental issues of basic cell biology. The elucidation and subsequent optimization of the mechanisms of those processes will contribute to improvements in efficiency of the production of useful compounds from n-alkanes.

References Arie M, Matsuda H, Furuhashi K, Takagi M (2000) Phylogenetic identification of n-alkane assimilating Candida yeasts based on nucleotide divergence in the 5' end of LSU rDNA gene. J Gen Appl Microbiol 46:257–262 Ashibe B, Hirai T, Higashi K, Sekimizu K, Motojima K (2007) Dual subcellular localization in the endoplasmic reticulum and peroxisomes and a vital role in protecting against oxidative stress of fatty aldehyde dehydrogenase are achieved by alternative splicing. J Biol Chem 282:20763–20773 Barth G, Gaillardin C (1996) Yarrowia lipolytica. In: Wolf K (ed) Non-conventional yeast in biotechnology. a handbook. Springer, Berlin/Heidelberg/New York, pp 313–388 Barth G, Gaillardin C (1997) Physiology and genetics of the dimorphic fungus Yarrowia lipolytica. FEMS Microbiol Rev 19:219–237 Bassel JB, Mortimer RK (1985) Identification of mutations preventing n-hexadecane uptake among 26 n-alkane non-utilizing mutants of Yarrowia (Saccharomycopsis) lipolytica. Curr Genet 9:579–586 Black PN, DiRusso CC (2007) Yeast acyl-CoA synthetases at the crossroads of fatty acid metabolism and regulation. Biochim Biophys Acta 1771:286–298 Cheng Q, Sanglard D, Vanhanen S, Liu HT, Bombelli P et al (2005) Candida yeast long chain fatty alcohol oxidase is a c-type haemoprotein and plays an important role in long chain fatty acid metabolism. Biochim Biophys Acta 1735:192–203 Cirigliano MC, Carman GM (1984) Isolation of a bioemulsifier from Candida lipolytica. Appl Environ Microbiol 48:747–750 Cirigliano MC, Carman GM (1985) Purification and characterization of liposan, a bioemulsifier from Candida lipolytica. Appl Environ Microbiol 50:846–850

12

R. Fukuda and A. Ohta

De Laurenzi V, Rogers GR, Hamrock DJ, Marekov LN, Steinert PM et al (1996) Sjögren-Larsson syndrome is caused by mutations in the fatty aldehyde dehydrogenase gene. Nat Genet 12:52–57 Demozay D, Rocchi S, Mas JC, Grillo S, Pirola L et al (2004) Fatty aldehyde dehydrogenase: potential role in oxidative stress protection and regulation of its gene expression by insulin. J Biol Chem 279:6261–6270 Dickinson FM, Wadforth C (1992) Purification and some properties of alcohol oxidase from alkanegrown Candida tropicalis. Biochem J 282:325–331 Dulermo R, Gamboa-Meléndez H, Dulermo T, Thevenieau F, Nicaud JM (2014) The fatty acid transport protein Fat1p is involved in the export of fatty acids from lipid bodies in Yarrowia lipolytica. FEMS Yeast Res 14:883–896 Dulermo R, Gamboa-Meléndez H, Ledesma-Amaro R, Thevenieau F, Nicaud JM (2016) Yarrowia lipolytica AAL genes are involved in peroxisomal fatty acid activation. Biochim Biophys Acta 1861:555–565 Eirich LD, Craft DL, Steinberg L, Asif A, Eschenfeldt WH et al (2004) Cloning and characterization of three fatty alcohol oxidase genes from Candida tropicalis strain ATCC 20336. Appl Environ Microbiol 70:4872–4879 Endoh-Yamagami S, Hirakawa K, Morioka D, Fukuda R, Ohta A (2007) Basic helix-loop-helix transcription factor heterocomplex of Yas1p and Yas2p regulates cytochrome P450 expression in response to alkanes in the yeast Yarrowia lipolytica. Eukaryot Cell 6:734–743 Eschenfeldt WH, Zhang Y, Samaha H, Stols L, Eirich LD et al (2003) Transformation of fatty acids catalyzed by cytochrome P450 monooxygenase enzymes of Candida tropicalis. Appl Environ Microbiol 69:5992–5999 Fickers P, Benetti PH, Waché Y, Marty A, Mauersberger S et al (2005) Hydrophobic substrate utilisation by the yeast Yarrowia lipolytica, and its potential applications. FEMS Yeast Res 5:527–543 Fukuda R (2013) Metabolism of hydrophobic carbon sources and regulation of it in n-alkaneassimilating yeast Yarrowia lipolytica. Biosci Biotechnol Biochem 77:1149–1154 Fukuda R, Ohta A (2013) Utilization of hydrophobic substrate by Yarrowia lipolytica. In: Barth G (ed) Yarrowia lipolytica genetics, genomics, and physiology. Springer, Heidelberg/New York/ Dordrecht/London, pp 111–119 Gatter M, Förster A, Bär K, Winter M, Otto C et al (2014) A newly identified fatty alcohol oxidase gene is mainly responsible for the oxidation of long-chain ω-hydroxy fatty acids in Yarrowia lipolytica. FEMS Yeast Res 14:858–872 Hirakawa K, Kobayashi S, Inoue T, Endoh-Yamagami S, Fukuda R, Ohta A (2009) Yas3p, an Opi1 family transcription factor, regulates cytochrome P450 expression in response to n-alkanes in Yarrowia lipolytica. J Biol Chem 284:7126–7137 Iida T, Ohta A, Takagi M (1998) Cloning and characterization of an n-alkane-inducible cytochrome P450 gene essential for n-decane assimilation by Yarrowia lipolytica. Yeast 14:1387–1397 Iida T, Sumita T, Ohta A, Takagi M (2000) The cytochrome P450ALK multigene family of an nalkane-assimilating yeast, Yarrowia lipolytica: cloning and characterization of genes coding for new CYP52 family members. Yeast 16:1077–1087 Iwama R, Kobayashi S, Ohta A, Horiuchi H, Fukuda R (2014) Fatty aldehyde dehydrogenase multigene family involved in the assimilation of n-Alkanes in Yarrowia lipolytica. J Biol Chem 289:33275–33286 Iwama R, Kobayashi S, Ohta A, Horiuchi H, Fukuda R (2015) Alcohol dehydrogenases and an alcohol oxidase involved in the assimilation of exogenous fatty alcohols in Yarrowia lipolytica. FEMS Yeast Res 15:fov014 Iwama R, Kobayashi S, Ishimaru C, Ohta A, Horiuchi H, Fukuda R (2016) Functional roles and substrate specificities of twelve cytochromes P450 belonging to CYP52 family in n-alkane assimilating yeast Yarrowia lipolytica. Fungal Genet Biol 91:43–54 Kemp GD, Dickinson FM, Ratledge C (1994) Occurrence of fatty alcohol oxidase in alkane-and fatty-acid-utilising yeasts and moulds. Appl Microbiol Biotechnol 40:873–875

Enzymes for Aerobic Degradation of Alkanes in Yeasts

13

Kim TH, Oh YS, Kim SJ (2000) The possible involvement of the cell surface in aliphatic hydrocarbon utilization by an oil-degrading yeast, Yarrowia lipolytica 180. J Microbiol Biotechnol 10:333–337 Kim D, Cryle MJ, De Voss JJ, Ortiz de Montellano PR (2007) Functional expression and characterization of cytochrome P450 52A21 from Candida albicans. Arch Biochem Biophys 464:213–220 Kobayashi S, Hirakawa K, Horiuchi H, Fukuda R, Ohta A (2013) Phosphatidic acid and phosphoinositides facilitate liposome association of Yas3p and potentiate derepression of ARE1 (alkaneresponsive element one)-mediated transcription control. Fungal Genet Biol 61:100–110 Kobayashi S, Tezaki S, Horiuchi H, Fukuda R, Ohta A (2015) Acidic phospholipid-independent interaction of Yas3p, an Opi1-family transcriptional repressor of Yarrowia lipolytica, with the endoplasmic reticulum. Yeast 32:691–701 Kurtzman CP, Fell JW, Boekhout T (eds) (2011) The yeasts: a taxonomic study, 5th edn. Elsevier, Amsterdam Liu CM, Johnson MJ (1971) Alkane oxidation by a particulate preparation from Candida. J Bacteriol 106:830–834 Mauersberger S, Drechsler H, Oehme G, Müller H-G (1992) Substrate specificity and stereoselectivity of fatty alcohol oxidase from the yeast Candida maltosa. Appl Microbiol Biotechnol 33:66–73 Mauersberger S, Ohkuma M, Schunck WH, Takagi M (1996) Candida maltosa. In: Wolf K (ed) Nonconventional yeasts in biotechnology. Springer, Berlin/Heidelberg/New York, pp 411–580 Mori K, Iwama R, Kobayashi S, Horiuchi H, Fukuda R, Ohta A (2013) Transcriptional repression by glycerol of genes involved in the assimilation of n-alkanes and fatty acids in yeast Yarrowia lipolytica. FEMS Yeast Res 13:233–240 Nakahara K, Ohkuni A, Kitamura T, Abe K, Naganuma T et al (2012) The Sjögren-Larsson syndrome gene encodes a hexadecenal dehydrogenase of the sphingosine 1-phosphate degradation pathway. Mol Cell 46:461–471 Nelson DR (2009) The cytochrome p450 homepage. Hum Genomics 4:59–65 Nicaud JM (2012) Yarrowia lipolytica. Yeast 29:409–418 Ohkuma M, Tanimoto T, Yano K, Takagi M (1991) CYP52 (cytochrome P450alk) multigene family in Candida maltosa: molecular cloning and nucleotide sequence of the two tandemly arranged genes. DNA Cell Biol 10:271–282 Ohkuma M, Muraoka S, Tanimoto T, Fujii M, Ohta A, Takagi M (1995) CYP52 (cytochrome P450alk) multigene family in Candida maltosa: identification and characterization of eight members. DNA Cell Biol 14:163–173 Ohkuma M, Zimmer T, Iida T, Schunck WH, Ohta A, Takagi M (1998) Isozyme function of n-alkane-inducible cytochromes P450 in Candida maltosa revealed by sequential gene disruption. J Biol Chem 273:3948–3953 Osumi M, Fukuzumi F, Yamada N, Nagatani T, Teranishi Y et al (1975) Surface structure of some Candida yeast cells grown on n-alkanes. J Ferment Technol 53:244–248 Panwar SL, Krishnamurthy S, Gupta V, Alarco AM, Raymond M et al (2001) CaALK8, an alkane assimilating cytochrome P450, confers multidrug resistance when expressed in a hypersensitive strain of Candida albicans. Yeast 18:1117–1129 Sagehashi Y, Horiuchi H, Fukuda R, Ohta A (2013) Identification and characterization of a gene encoding an ABC transporter expressed in the dicarboxylic acid-producing yeast Candida maltosa. Biosci Biotechnol Biochem 77:2502–2504 Sanglard D, Chen C, Loper JC (1987) Isolation of the alkane inducible cytochrome P450 (P450alk) gene from the yeast Candida tropicalis. Biochem Biophys Res Commun 144:251–257 Scheller U, Zimmer T, Becher D, Schauer F, Schunck WH (1998) Oxygenation cascade in conversion of n-alkanes to α, ω-dioic acids catalyzed by cytochrome P450 52A3. J Biol Chem 273:32528–32534

14

R. Fukuda and A. Ohta

Seghezzi W, Sanglard D, Fiechter A (1991) Characterization of a second alkane-inducible cytochrome P450-encoding gene, CYP52A2, from Candida tropicalis. Gene 106:51–60 Seghezzi W, Meili C, Ruffiner R, Kuenzi R, Sanglard D, Fiechter A (1992) Identification and characterization of additional members of the cytochrome P450 multigene family CYP52 of Candida tropicalis. DNA Cell Biol 11:767–780 Sophos NA, Pappa A, Ziegler TL, Vasiliou V (2001) Aldehyde dehydrogenase gene superfamily: the 2000 update. Chem Biol Interact 130–132:323–337 Soupene E, Kuypers FA (2008) Mammalian long-chain acyl-CoA synthetases. Exp Biol Med 233:507–521 Takai H, Iwama R, Kobayashi S, Horiuchi H, Fukuda R, Ohta A (2012) Construction and characterization of a Yarrowia lipolytica mutant lacking genes encoding cytochromes P450 subfamily 52. Fungal Genet Biol 49:58–64 Tanaka A, Fukui S (1989) Metabolism of n-Alkanes. In: The yeast. Academic, London/San Diego/New York/Berkeley/Boston/Sydney/Tokyo/Toronto, pp 261–287 Tenagy, Park JS, Iwama R, Kobayashi S, Ohta A et al (2015) Involvement of acyl-CoA synthetase genes in n-alkane assimilation and fatty acid utilization in yeast Yarrowia lipolytica. FEMS Yeast Res 15:fov031 Thevenieau F, Le Dall MT, Nthangeni B, Mauersberger S, Marchal R, Nicaud JM (2007) Characterization of Yarrowia lipolytica mutants affected in hydrophobic substrate utilization. Fungal Genet Biol 44:531–542 Ueda M, Tanaka A (1990) Long-chain aldehyde dehydrogenase of Candida yeast. Methods Enzymol 188:176–178 Van Bogaert IN, Saerens K, De Muynck C, Develter D, Soetaert W, Vandamme EJ (2007) Microbial production and application of sophorolipids. Appl Microbiol Biotechnol 76:23–34 Van Bogaert IN, De Mey M, Develter D, Soetaert W, Vandamme EJ (2009) Importance of the cytochrome P450 monooxygenase CYP52 family for the sophorolipid-producing yeast Candida bombicola. FEMS Yeast Res 9:87–94 Vanhanen S, West M, Kroon JT, Lindner N, Casey J et al (2000) A consensus sequence for longchain fatty-acid alcohol oxidases from Candida identifies a family of genes involved in lipid ω-oxidation in yeast with homologues in plants and bacteria. J Biol Chem 275:4445–4452 Verhoeven NM, Jakobs C, Carney G, Somers MP, Wanders RJ, Rizzo WB (1998) Involvement of microsomal fatty aldehyde dehydrogenase in the α-oxidation of phytanic acid. FEBS Lett 429:225–228 Wang J, Zhang B, Chen S (2011) Oleaginous yeast Yarrowia lipolytica mutants with a disrupted fatty acyl-CoA synthetase gene accumulate saturated fatty acid. Process Biochem 46:1436–1441 Watkins PA, Ellis JM (2012) Peroxisomal acyl-CoA synthetases. Biochim Biophys Acta 1822:1411–1420 Yadav JS, Loper JC (1999) Multiple p450alk (cytochrome P450 alkane hydroxylase) genes from the halotolerant yeast Debaryomyces hansenii. Gene 226:139–146 Yamada T, Nawa H, Kawamoto S, Tanaka A, Fukui S (1980) Subcellular localization of long-chain alcohol dehydrogenase and aldehyde dehydrogenase in n-alkane-grown Candida tropicalis. Arch Microbiol 128:145–151 Yamagami S, Morioka D, Fukuda R, Ohta A (2004) A basic helix-loop-helix transcription factor essential for cytochrome P450 induction in response to alkanes in yeast Yarrowia lipolytica. J Biol Chem 279:22183–22189 Zimmer T, Ohkuma M, Ohta A, Takagi M, Schunck WH (1996) The CYP52 multigene family of Candida maltosa encodes functionally diverse n-alkane-inducible cytochromes P450. Biochem Biophys Res Commun 224:784–789 Zimmer T, Iida T, Schunck WH, Yoshida Y, Ohta A, Takagi M (1998) Relation between evolutionary distance and enzymatic properties among the members of the CYP52A subfamily of Candida maltosa. Biochem Biophys Res Commun 251:244–247

Aerobic Degradation of Aromatic Hydrocarbons D. Pérez-Pantoja, B. González, and D.H. Pieper

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Catabolism of Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Peripheral Reactions Preparing Aromatic Hydrocarbons for Ring-Cleavage . . . . . . . . . 2.2 Side Chain Processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Central Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 CoA Dependent Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

3 4 4 11 18 31 33 34

Abstract

Aromatic hydrocarbons are widely distributed in nature. They are found as lignin components, aromatic amino acids and xenobiotic compounds, among others. Microorganisms, mostly bacteria, degrade an impressive variety of such chemical structures. The major principle of aromatic hydrocarbon biodegradation is that a broad range of aromatic hydrocarbons are transformed by peripheral reactions to a restricted range of central intermediates, which are subject to ring-cleavage and funneling into the Krebs cycle. Key enzymes in aerobic aromatic degradation are D. Pérez-Pantoja Departamento de Bioquímica y Biología Molecular, Facultad de Ciencias Biológicas, Universidad de Concepción, Concepción, Chile e-mail: [email protected] B. González Facultad de Ingeniería y Ciencias, Universidad Adolfo Ibáñez, Santiago, Chile e-mail: [email protected] D.H. Pieper (*) Microbial Interactions and Processes Research Group, HZI – Helmholtz Centre for Infection Research, Braunschweig, Germany e-mail: [email protected] # Springer International Publishing AG 2016 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-319-39782-5_10-1

1

2

D. Pérez-Pantoja et al.

oxygenases, preparing aromatics for ring-cleavage by the introduction of hydroxyl functions and catalyzing cleavage of the aromatic ring. The diverse monooxygenases and dioxygenases involved in hydroxylations, a significant proportion of them possessing relaxed substrate specificity, are discussed as well as the broad diversity of side chain processing transformations involved in the formation of ring-cleavage central intermediates. Ring cleavage dioxygenases, covering intradiol ring cleavage of ortho dihydroxylated intermediates, and a large number of diverse but mechanistically related extradiol dioxygenases participating in ring cleavage of ortho and para dihydroxylated intermediates are also discussed. CoA dependent aerobic routes to allow ringcleavage of aromatic hydrocarbons without involvement of dihydroxylated aromatic intermediates have been described in the last years and are also reviewed. The degradation of heteroarenes will not be described in this chapter.

1

Introduction

Aromatic hydrocarbons are very important building blocks of biomass and are widely distributed in nature, being produced by a variety of biological and biogeochemical processes, and range in size from low-molecular mass compounds to polymers. The most abundant fraction of aromatic hydrocarbons is formed by the lignin of higher plants, which in fact is the second most abundant polymer in nature after cellulose, comprising about 25% of the land-based biomass on Earth (Kirk and Farrell 1987). During the decomposition process, lignin degradation products – together with other plant-derived aromatic hydrocarbons – contribute to the formation of recalcitrant organic matter in soils. Other ubiquitous sources of aromatic hydrocarbons are the aromatic amino acids. Finally, the extensive use of natural and xenobiotic aromatic hydrocarbons in industrial processes, coupled with inadequate waste management strategies, has led to the positioning of these compounds among the most stable and persistent organic pollutants. The degradation of aromatic polymers is an important component of global biogeochemical cycles and is accomplished almost exclusively by microorganisms which have evolved diverse strategies to degrade aromatic hydrocarbons, and thereby derive carbon and energetic benefit from them. The large spectrum of aromatic substrates degradable by microbial communities is assured by a huge catabolic diversity present in different microbial species and the relaxed substrate specificity of some of the catabolic pathways (Pérez-Pantoja et al. 2008). The major principle of aromatic hydrocarbon biodegradation is that catabolic pathways involve two key steps: the activation of the thermodynamically stable benzene ring from structurally diverse aromatics, and its subsequent cleavage. In aerobic catabolism, oxygenases accomplish the main role in both steps (Duarte et al. 2014).

Aerobic Degradation of Aromatic Hydrocarbons

2

3

Catabolism of Aromatic Hydrocarbons

Aerobic microorganisms usually initiate degradation by activation of the aromatic nucleus through oxygenation reactions. A few central intermediates such as catechols, protocatechuates, gentisates and (hydroxy)benzoquinols, are produced by the introduction of hydroxyl groups, usually in ortho- or para-position to one another. These intermediates are subject to oxygenolytic ring cleavage followed by channeling of the ring-cleavage products into the central metabolism. Alternatively aromatic hydrocarbons, even under aerobic conditions, can be metabolized through the corresponding CoA thioesters and subject of non-oxygenolytic ring cleavage.

2.1

Peripheral Reactions Preparing Aromatic Hydrocarbons for Ring-Cleavage

2.1.1 Rieske Non-Heme Iron Oxygenases The so called Rieske non-heme iron oxygenases are one of the key families of enzymes important for aerobic activation and thus degradation of aromatic hydrocarbons such as benzoate, benzene, toluene, phthalate, naphthalene or biphenyl (Gibson and Parales 2000; Duarte et al. 2014). These multicomponent enzyme complexes, composed of a terminal oxygenase component and different electron transport proteins, usually catalyze the incorporation of two oxygen atoms into the aromatic ring to form arene- cis-dihydrodiols (although some members of this superfamily also catalyze monooxygenations), a reaction which is followed by a dehydrogenation usually catalyzed by cis-dihydrodiol dehydrogenases to give (substituted) catechols. Members of the Rieske non-heme iron oxygenases are since decades known to be involved in the degradation of benzoate (Gibson et al. 1968), converting it to 1-carboxy-1,2- cis-dihydroxycyclohexa-3,5-diene (benzoate- cis-dihydrodiol, see Fig. 1) (Reiner and Hegeman 1971). A benzoate dihydrodiol dehydrogenase catalyzes the dehydrogenation to a β-ketoacid, which spontaneously decarboxylates to catechol (Reiner 1972). The benzoate dioxygenases belong to the so-called benzoate subgroup of Rieske non-heme iron oxygenases and are composed of a reductase and an oxygenase component with an (αβ)3 quaternary structure (Wolfe et al. 2002), with each α-subunit containing a mononuclear non-heme iron active site and a Riesketype (2Fe-2S) cluster. Some of these benzoate dioxygenases have been studied in detail and it is well established that despite a significant degree in sequence identity, they differ significantly in substrate specificity (Reineke and Knackmuss 1978), with toluate dioxygenase of P. putida mt-2 being capable to transform meta- and parasubstituted benzoates whereas benzoate dioxygenase only transforms benzoate and meta-substituted benzoates. Ortho-substituted benzoates (2-chloro- and 2-methyl-) are poor substrates for both enzymes (Yamaguchi and Fujisawa 1980). Similar two-component enzyme systems (see Fig. 1) are responsible for 1,2-dioxygenation of anthranilate (Bundy et al. 1998), an intermediary metabolite of tryptophan degradation and a precursor for the Pseudomonas quinolone signal

Fig. 1 Dendrogram showing the relatedness among oxygenase α-subunits of Rieske non-heme iron oxygenases. Reactions catalyzed by enzymes indicated are given to the exterior of the figure, together with subsequent reactions resulting in the formation of central intermediates, which are subject to ring-cleavage reaction. Unstable intermediates are shown in brackets

4 D. Pérez-Pantoja et al.

Aerobic Degradation of Aromatic Hydrocarbons

5

(Farrow and Pesci 2007). Anthranilate dioxygenases catalyze the formation of catechol without requirement of a dehydrogenase due to spontaneous decarboxylation and deamination of 2-amino-1-carboxy-1,2-cis-dihydroxycyclohexa-3, 5-diene (anthranilate-cis-dihydrodiol, see Fig. 1). At least the enzyme system of Acinetobacter baylyi ADP1 also transforms benzoate (Eby et al. 2001), however, other ortho-substituted benzoates are only poorly converted, differentiating two-component anthranilate 1,2-dioxygenases from two-component 2-halobenzoate 1,2-dioxygenases. Rieske non-heme iron oxygenases are also involved in the degradation of p-cymene ( p-isopropyltoluene), a natural product identified in volatile oils from various plants. p-Cymene metabolism is initiated by oxidation of the methyl substituent with p-cumate as intermediate (Eaton 1997), which is attacked by a two-component p-cumate dioxygenase (Eaton 1996). In contrast to enzymes described above, p-cumate dioxygenases do not attack on the carboxysubstituted carbon atom but on the meta- and ortho- carbon atom to form 2,3-dihydroxy-2,3dihydro-p-cumate, followed by dehydrogenation to 2,3-dihydroxy-p-cumate (Fig. 1). Also aniline (aminobenzene) degradation seems to be mediated by related two-component dioxygenases and the α-subunits of the aniline dioxygenase system share significant identity with those of benzoate dioxygenases (see Fig. 1). However, in contrast to all enzyme systems described above, aniline dioxygenase consists of five protein components, all necessary for a functional enzyme (Fukumori and Saint 1997). Based on sequence similarities, it is proposed that three of them function as the large and small subunit of aniline dioxygenase and reductase, respectively. Additional proteins show similarity to glutamine synthetase and glutamine amidotransferase, respectively, and maybe are involved in transfer of the amino group or release of ammonia (Liang et al. 2005). Interestingly, enzyme systems catalyzing the transformation of anthranilate (and also of 2-halobenzoate), only distantly related to those of the benzoate subgroup of Rieske non-heme iron oxygenases, have also been described. Anthranilate dioxygenase of Burkholderia cepacia DBO1 is a three-component Rieske non-heme iron dioxygenase composed of a reductase, a ferredoxin and a two-subunit oxygenase which, besides anthranilate, also transforms salicylate (but not 2-chlorobenzoate) to catechol (Chang et al. 2003). Besides three component 2-halobenzoate 1,2-dioxygenases, the enzymes most closely related to three-component anthranilate dioxygenase have been characterized as salicylate 1-hydroxylases (see Fig. 1). The occurrence of three component salicylate 1-hydroxylases, contrasting the previously known single component flavoprotein monooxygenases was first reported in Sphingobium sp. strain P2, which synthesized three isoenzymes (Pinyakong et al. 2003). However, despite the similarity and identical products formed from salicylate, anthranilate dioxygenase from B. cepacia DBO1 and multicomponent salicylate 1-hydroxylases are quite distinct and whereas anthranilate dioxygenase catalyzes a dioxygenation of anthranilate, salicylate 1-hydroxylase catalyzes a monooxygenation of anthranilate with 2-aminophenol as product (Jouanneau et al. 2007).

6

D. Pérez-Pantoja et al.

Phylogenetic analyses (see Fig. 1) show that the α-subunits of three-component anthranilate dioxygenase, salicylate 1-hydroxylase and 2-halobenzoate 1,2-dioxygenase form a distinct group of enzymes together with salicylate 5-hydroxylases and terephthalate dioxygenases (Duarte et al. 2014). Salicylate 5-hydroxylase, transforming salicylate into gentisate, has initially been reported in Ralstonia sp. U2 (Fuenmayor et al. 1998). The encoding genes were identified to be in the same operon as those of the naphthalene dioxygenase and it was shown that naphthalene dioxygenase and salicylate 5-hydroxylase share the chain for the transport of electrons (Zhou et al. 2002), a ferredoxin reductase and ferredoxin, as in three component oxygenases. As mentioned above the α-subunits of terephthalate dioxygenases belong to the same subgroup of enzymes, all of them involved in the metabolism of carboxylated aromatics (see Fig. 1). Terephthalate dioxygenase catalyzes a 1,2-dioxygenation with 1,2-dihydroxy-3,5-cyclohexadiene-1,4-dicarboxylate as product, which is dehydrogenated, as described above for benzoate dihydrodiol, to a β-ketoacid which spontaneously decarboxylates to protocatechuate (Schläfli et al. 1994). Terephthahlate dioxygenase, at least from C. testosteroni T-2, seems to be of restricted substrate specificity and neither attacks benzoate nor isophthalate or phthalate (Schläfli et al. 1994). In contrast to salicylate 1- and 5-hydroxylase or anthranilate dioxygenase, terephthalate dioxygenase seems to be active as a two-component dioxygenase consisting of α- and β-subunits of the oxygenase and a reductase (Sasoh et al. 2006). The degradation of phthalate has initially been described in B. cepacia (Batie et al. 1987). In Proteobacteria, phthalate is subject to 4,5-dioxygenation giving rise to the dihydrodiol which is dehydrogenated to 4,5-dihydroxyphthalate (Fig. 1). As the attack does not involve a carboxylated carbon atom, no spontaneous decarboxylation is involved. Decarboxylation is catalyzed by a 4,5-dihydroxyphthalate decarboxylase yielding protocatechuate (Fig. 1) (Lee et al. 1994). The oxygenase of phthalate 4,5-dioxygenase differs from all above described Rieske non-heme iron oxygenases as being composed only of α-subunits, a feature shared with carbazole dioxygenases, 3-chlorobenzoate 4,5-dioxygenase and isophthalate dioxygenase from C. testosteroni YZW-D (Wang et al. 1995). Interestingly, phthalate 4,5-dioxygenases have only been described in Proteobacteria, whereas Actinobacteria such as Arthrobacter keyseri 12B degrade phthalate via 3,4-dioxygenation and through 3,4-dihydroxyphthalate, which is decarboxylated to protocatechuate (Fig. 1) as common intermediate of both the 4,5- and 3,4-dioxygenolytic pathways (Eaton 2001). In contrast to phthalate 4,5-dioxygenase, phthalate 3,4-dioxygenase is a three-component dioxygenase composed of a two-subunit terminal oxygenase, a ferredoxin and a ferredoxin reductase, with the α-subunit being closely related to α-subunits of naphthalene or phenanthrene dioxygenases from Actinobacteria (Fig. 1). However, not only the oxygenase systems of phthalate 4,5- and phthalate 3,4-dioxygenase are different. cis-3,4dihydro-3,4-dihydroxyphthalate dehydrogenase belongs to the aldo/keto reductase superfamily (Eaton 2001), and differs from cis-4,5-dihydro-4,5-dihydroxyphthalate dehydrogenase, which belongs to the GFO/IDH/MOCA family (Chang and Zylstra

Aerobic Degradation of Aromatic Hydrocarbons

7

1998). In addition, 3,4-dihydroxyphthalate decarboxylases are unrelated to 4,5-dihydroxyphthalate decarboxylases. Also the catabolism of 3-phenylpropionate (and cinnamate) can be initiated by dioxygenation through the action of a Rieske non-heme iron dioxygenase (Fig. 1). 3-Phenylpropionate dioxygenase, similarly to p-cumate dioxygenase, inserts oxygen into positions 2- and 3- of the aromatic ring yielding cis-3-(3-carboxyethyl)-3,5cyclohexadiene-1,2-diol, followed by dehydrogenation through a dehydrogenase of the short chain alcohol dehydrogenase family to 2,3-dihydroxyphenylpropionate (or 2,3-dihydroxycinnamate) (Diaz et al. 2001). 3-Phenylpropionate 2,3-dioxygenases are related to the benzene/toluene/isopropylbenzene/biphenyl subgroup (enzymes typically described to be capable to transform the aforementioned compounds) of Rieske non-heme iron oxygenases (Fig. 1), important for degradation of hydrophobic substrates.

2.1.2 Soluble Diiron Monooxygenases Enzymes capable to monooxygenate benzene/toluene to phenol/methylphenol and phenols to catechols belong to an evolutionary related family of soluble diiron monooxygenases (Leahy et al. 2003), which are enzyme complexes consisting of an electron transport system comprising a reductase (and in some cases a ferredoxin), a catalytic effector protein which contains neither organic cofactors nor metal ions and is assumed to play a role in assembly of an active oxygenase (Powlowski et al. 1997), and a terminal hydroxylase with a (αβγ)2 quaternary structure and a diiron center contained in each α-subunit. Theses monooxygenases are classified according to their α-subunits, which are assumed to be the site of substrate hydroxylation, into four different phylogenetic groups: the soluble methane monooxygenases, the alkene monooxygenase of Rhodococcus corallinus B-276, the phenol hydroxylases, and the four-component alkene/aromatic monooxygenases (Leahy et al. 2003). The four component alkene/aromatic monooxygenases comprise enzymes that oxidize non-hydroxylated compounds and their gene clusters usually encode a ferredoxin component (Leahy et al. 2003). Monooxygenases that hydroxylate toluene at all three possible positions, producing 2-methyl-, 3-methyl- or 4-methylphenol have been described (Olsen et al. 1994; Shields et al. 1989; Whited and Gibson 1991). However, later analysis has shown that the toluene monooxygenase of R. pickettii PK01, which had been reported previously to hydroxylate toluene at the meta position, producing primarily 3-methylphenol (Olsen et al. 1994), hydroxylates toluene predominantly at the para position producing 4-methylphenol (Fishman et al. 2004). Some of the enzymes of this subfamily, like the toluene monooxygenases of P. stutzeri OX1 (Bertoni et al. 1998) or R. pickettii PK01, or toluene 4-monooxygenase of P. mendocina KR1 have been shown to oxidize also phenol and methylphenols to the respective catechols and even further to 1,2,3-trihydroxybenzene (Tao et al. 2004b). The phenol hydroxylases comprise the multicomponent phenol hydroxylase of the methylphenol degrading Pseudomonas sp. strain CF600 (Shingler et al. 1992) and P. putida 35X (Ng et al. 1994) but also the toluene 2-monooxygenase of B. cepacia G4 (Newman and Wackett 1995), among others. The respective gene

8

D. Pérez-Pantoja et al.

clusters usually lack a ferredoxin gene (Leahy et al. 2003). Whereas all these monooxygenases share the capability to hydroxylate phenol and methylsubstituted derivatives, only a few enzymes of this group have been shown to hydroxylate the unactivated benzene nucleus. These enzymes, among them toluene 2-monooxygenase of strain G4, sequentially oxidize toluene to 2-methylphenol and further to 3-methylcatechol (Newman and Wackett 1995). Phenol hydroxylase of P. stutzeri OX1 was also shown to be capable to transform benzene or toluene, however, the specificity constant kcat/Km was 2–3 orders of magnitude lower compared to phenol as substrate, evidencing phenol to be the highly preferred substrate (Cafaro et al. 2004). It should be noted that in the recent years, the crystal structure of toluene monooxygenase from P. stutzeri OX1 has been solved and various mutagenesis studies on this group of enzymes have been performed, inter alia to elucidate amino acid residues determining regioselectivity (Fishman et al. 2005; Tao et al. 2004a), but also to identify and change residues critical for phenol hydroxylation (Tao et al. 2004b; Vardar and Wood 2004).

2.1.3 Single and Two Component Flavoprotein Monooxygenases Flavoprotein monooxygenases are involved in a wide variety of biological processes including biosynthesis of antibiotics and siderophores or biodegradation of aromatic hydrocarbons. The reactions use NAD(P)H and O2 as co-substrates and insert one atom of oxygen into the substrate. These enzymes utilize a general cycle in which NAD(P)H reduces the flavin, and the reduced flavin reacts with O2 to form a C4a-(hydro)peroxyflavin intermediate, which is the oxygenating agent. Hydroxylation of the substrate yields the flavin-C4a-hydroxide, from which, finally, water is eliminated (Ballou et al. 2005). This catalytic process has diverse requirements that are difficult to be satisfied by a single catalytic site. Two general strategies have evolved to deal with this complex chemical problem (Ballou et al. 2005). First, in the case of single-component flavin monooxygenases, the enzyme undergoes significant protein and flavin dynamics during catalysis. The second approach uses two components to separate the catalytic tasks, an oxidoreductase to generate reduced flavin, and an oxygenase to receive the reduced flavin, react with O2 and hydroxylate the substrate (Ballou et al. 2005). Flavin monooxygenases have been classified according to sequence and structural data in six classes (van Berkel et al. 2006), with classes A, D and E being of special importance for aromatic hydrocarbon degradation. Class A Single-Component Flavin Monooxygenases Class A flavin monooxygenases are encoded by a single gene, contain a tightly bound FAD as cofactor, depend on NADH or NADPH as cofactor and are structurally composed of one dinucleotide binding domain to bind FAD. They are widely distributed in different bacterial taxa as hydroxylases in ortho- or para-position of aromatic compounds that already contain a hydroxyl group (van Berkel et al. 2006) (Fig. 2). The 4-hydroxybenzoate 3-hydroxylase of P. fluorescens is one of the most thoroughly studied Class A enzymes (Entsch et al. 1987). This enzyme (encoded by the

Aerobic Degradation of Aromatic Hydrocarbons

9

Fig. 2 Dendrogram showing the relatedness of single component flavoprotein monooxygenases. Reactions catalyzed by enzymes indicated are given to the exterior of the figure

pobA gene) catalyzes the conversion of 4-hydroxybenzoate to protocatechuate. Typically pobA gene products show a narrow specificity and in addition to 4-hydroxybenzoate also hydroxylate 4-aminobenzoate to 4-amino-3hydroxybenzoate (Entsch and van Berkel 1995). The purification from several bacterial sources and the presence of pobA homologous genes in Actinomycetes, α, β, and γ- Proteobacteria is indicative of a broad distribution of this enzyme. In a dendrogram of single-component monooxygenases, PobA gene products are exclusively clustered without any additional sequences predicting a common evolutionary origin (Fig. 2). Single component salicylate 1-hydroxylases catalyze the transformation of salicylate to catechol and were the first flavin monooxygenase characterized (Yamamoto et al. 1965). This enzyme has also been purified and characterized from many microorganisms showing a relatively broader specificity including chloro- and methylsalicylates as substrates (Camara et al. 2007; Lehrbach et al. 1984). Salicylate 1-hydroxylases clustered very close in the dendrogram with the exception of NahW, an isoenzyme found in P. stutzeri AN10 (Bosch et al. 1999) (Fig. 2). Two distinct single component monooxygenases acting on 3-hydroxybenzoate have been described, 3-hydroxybenzoate 4-hydroxylase producing protocatechuate

10

D. Pérez-Pantoja et al.

(Hiromoto et al. 2006) and 3-hydroxybenzoate 6-hydroxylase producing gentisate (Wang et al. 1987). The latter enzymes are closely related to salicylate 1-hydroxylases (Fig. 2), and are, in genome sequencing projects often misleadingly annotated as salicylate hydroxylases. 3-Hydroxybenzoate 4-hydroxylases have so far only been reported in Comamonas strains and are only distantly related to other hydroxybenzoate hydroxylases. Another group of single component flavin monooxygenases belonging to class A, termed phenol hydroxylases, has been described from phenol degrading Pseudomonas strains, among them PheA from Pseudomonas sp. strain EST1001, which transforms phenol and 3-methylphenol (Nurk et al. 1991). This substrate specificity significantly differs from that described for a group of closely related enzymes termed 2,4-dichlorophenol hydroxylases. Various other closely related enzymes have confirmed or assumed activity as 2-hydroxybiphenyl 3-hydroxylase (HbpA of P. azelaica (Suske et al. 1997)), benzoquinol monooxygenases (chlorobenzoquinol monooxygenase of Pimelobacter simplex E3 (AY822041) and methylbenzoquinol monooxygenase of Burkholderia sp. NF100 (Tago et al. 2005)) or 2-hydroxyphenylpropionate 3-hydroxylase from Rhodococcus sp. V49 (Powell and Archer 1998)) (see Fig. 2). Enzymes of this group are commonly and misleadingly annotated as 2,4-dichlorophenol hydroxylases. Not only above mentioned benzoquinol monooxygenases can catalyze hydroxylation of dihydroxylated aromatic benzoquinal since a resorcinol (1,3-dihydroxybenzene) monooxygenase forming hydroxybenzoquinol, only distantly related to all above mentioned enzymes has been identified in Corynebacterium glutamicum (Huang et al. 2006). Whereas a class A 2-hydroxyphenylpropionate 3-hydroxylase forming 2,3-dihydroxyphenylpropionate has thus far only been identified in Rhodococcus (Powell and Archer 1998), class A 3-hydroxyphenylpropionate 2-hydroxylases forming the same product have been identified in Actinobacteria and Proteobacteria (Barnes et al. 1997; Ferrández et al. 1997) and clustered together in dendrogram (Fig. 2). Also 3-hydroxyphenylacetate is subject to monooxygenation by a 3-hydroxyphenylacetate 6-hydroxylase (Fig. 2), giving rise to homogentisate (Arias-Barrau et al. 2005). The MhaA 3-hydroxyphenylacetate 6-hydroxylase of P. putida, however, necessitates the presence of MhaB, described as an essential coupling protein, for activity. Thus it constitutes a novel type of two-component hydroxylase (AriasBarrau et al. 2005), distinct from the more classical two-component flavoprotein monooxygenases described below. Two-Component Flavin Monooxygenases During the past few years, several two-component aromatic hydroxylases consisting of an oxidoreductase and an oxygenase have been identified which have no structural or sequence similarities to the single-component enzymes and were classified as type D and E flavoprotein monooxygenases (Ballou et al. 2005). Class D flavoprotein monooxygenases (van Berkel et al. 2006) use FADH2 generated by the oxidoreductase as a coenzyme, show a structural resemblance

Aerobic Degradation of Aromatic Hydrocarbons

11

with the acyl-CoA dehydrogenase fold, and comprise 4-hydroxyphenylacetate 3-hydroxylase (e.g., from E. coli, Acinetobacter baumanii or P. putida), phenol hydroxylase from Bacillus thermoglucosidasius (van Berkel et al. 2006), and also trichlorophenol monooxygenases. Alternatively to 4-hydroxyphenylacetate 3-hydroxylase, a 1-hydroxylation of 4-hydroxyphenylacetate with homogentisate as reaction product has been described (Hareland et al. 1975), however, no information is available so far on the encoding gene nor on the detailed enzyme mechanism and thus on the class this enzyme belongs to. Styrene monooxygenase (StyA) has been identified from various Pseudomonas strains (Beltrametti et al. 1997), and was classified as Class E flavoprotein monooxygenase. An evolutionary link with the Class A flavoprotein monooxygenases was suggested (van Berkel et al. 2006). Styrene monooxygenases are highly enantioselective in oxidizing styrene and some of its derivatives to the respective epoxides.

2.2 2.2.1

Side Chain Processing

Oxidations of Methyl Groups in Methyl-Substituted Aromatic Hydrocarbons Some bacteria catabolize methyl-substituted aromatic hydrocarbons such as toluene, xylenes (Assinder and Williams 1990), p-cymene (DeFrank and Ribbons 1976), and p-cresol ( p-methylphenol) (Hopper and Taylor 1977) by oxidizing the methyl group to the corresponding acids (Fig. 3). The best known aromatic methyl-substituent oxidation pathway is that encoded by TOL plasmid, pWW0, in P. putida mt-2 (Assinder and Williams 1990), involving oxidation of toluene, m-xylene, and p-xylene to benzoate, m-toluate, and p-toluate, respectively (Fig. 3). The pathway is initiated by a monooxygenase, which catalyzes the oxidation of toluene (or m- or p-xylene) to benzyl alcohol. This monooxygenase is a two-component enzyme consisting of a XylA reductase subunit, which transfers electrons from NADH through FAD and a [2Fe-2S] center to the membraneassociated XylM hydroxylase subunit. There, one atom of activated molecular oxygen is inserted into the methyl group while the other oxygen atom is reduced to water. XylM shares significant amino acid identity (approx. 25%) with the integral-membrane non-heme diiron AlkB alkane hydroxylases. The conversion of benzyl alcohols to benzaldehydes is catalyzed by an NAD+linked alcohol dehydrogenase. It seems that dehydrogenation is the major route for this transformation, however, the alcohol is presumably oxidized by the monooxygenase to an unstable gem-diol intermediate which is recognizable as the hydrate of the corresponding benzaldehyde (Harayama et al. 1986). The aldehyde formed is then oxidized to benzoate (or m-toluate or p-toluate), by an NAD+-linked aldehyde dehydrogenase. Three enzymes – a two-component monooxygenase, an alcohol dehydrogenase, and an aldehyde dehydrogenase – have also been involved in the catabolism of p-cymene through p-cumate ( p-isopropylbenzoate) (DeFrank and Ribbons 1976; Eaton 1997) (Fig. 3). However, only the monooxygenase and

12

D. Pérez-Pantoja et al.

Fig. 3 Peripheral reactions involved in oxidation of methyl groups in methyl-substituted aromatic hydrocarbons. Reaction intermediates are shown in brackets

aldehyde dehydrogenase of the p-cymene catabolic pathway are related to the analogous enzymes of the TOL plasmid (Eaton 1997). The metabolism of p-cresol, as studied in P. putida NCIB 9866 (Hopper 1976), is initiated by a periplasmatic p-cresol methylhydroxylase (PCMH) to p-hydroxybenzaldehyde with the transient formation of p-hydroxybenzyl alcohol (Keat and Hopper 1978). This enzyme, a flavocytochrome, consists of an α subunit containing the active site and a FAD covalently linked to tyrosine, and a c-type cytochrome containing β subunit (McIntire et al. 1985). The product of PCMH, p-hydroxybenzaldehyde, is oxidized by a dehydrogenase to p-hydroxybenzoate (Cronin et al. 1999). 4-Ethylphenol methylenehydroxylase from P. putida JD1 is related to PCMH (Reeve et al. 1989) (Fig. 3). Catabolism of 4-ethylphenol in strain JD1 proceeds by hydroxylation to give 1-(40 -hydroxyphenyl)ethanol, followed by dehydrogenation to the ketone, 4-hydroxyacetophenone (Darby et al. 1987) (Fig. 3).

Aerobic Degradation of Aromatic Hydrocarbons

13

2.2.2 O-Demethylations Methoxylated aromatic hydrocarbons such as vanillate or syringate are important intermediate metabolites from lignin. Some demethylating systems have been described in aerobic bacteria to deal with such metabolites. Vanillate O-demethylase belongs to the Rieske non-heme iron oxygenases (see Fig. 1). It is a two-component oxygenase consisting of a reductase (VanB) and an oxygenase (VanA). The oxygenase is composed only of α-subunits and shares similarity with phthalate 4,5-dioxygenases (see Sect. 2.1.1 and Fig. 1). This type of demethylase is involved in vanillate degradation by most vanillate-degrading aerobic bacteria, such as Pseudomonas and Acinetobacter strains (Buswell and Ribbons 1988). Vanillate demethylases have a wide substrate specificity and vanillate analogs that are transformed share the common property of a methoxy- or methyl group in meta-position to the carboxyl group (such as m-anisate, 3,4-dimethoxybenzoate or m-toluate) (Morawski et al. 2000). The enzyme is not only able to demethylate methoxy groups but also to monohydroxylate methyl groups in the meta-position. Another type of demethylase is the tetrahydrofolate (THF) dependent demethylases, which have mainly been reported in anaerobic bacteria. However, THF dependent syringate and vanillate O-demethylases have also been reported in Sphingomonas paucimobilis SYK-6 (Abe et al. 2005; Masai et al. 2004) (Fig. 4). The deduced amino acid sequence of DesA syringate O-demethylase shows similarity to the THF-dependent aminomethyltransferase of E. coli involved in glycine cleavage. DesA converts syringate to 3-O-methylgallate only in the presence of THF, with the concomitant formation of 5-methyl-THF. Vanillate and 3-O-methylgallate are also used as substrates for DesA, however with poor activity (Masai et al. 2004). More recently, a second THF dependent O-demethylase termed LigM, showing 49% of amino acid sequence identity with DesA, was discovered in S. paucimobilis SYK-6. In the presence of THF, LigM converts vanillate and 3-O-methylgallate into protocatechuate and gallate, respectively (Fig. 4), whereas syringate was not transformed (Abe et al. 2005). Cytochrome P450 O-demethylase systems have been described for the demethylation of veratrole and guaiacol to catechol in Streptomyces setonii and Moraxella sp. respectively (Sauret-Ignazi et al. 1988; Sutherland 1986), and for demethylation of 2-ethoxyphenol and 4-methoxybenzoate in Rhodococcus rhodochrous (Karlson et al. 1993), however, identification of the cytochrome P450 genes is lacking. 2.2.3 Aromatic Acid Decarboxylations Decarboxylations are required for degradation of phthalate, 5-carboxyvanillate and 2,6-dihydroxybenzoate, among other aromatic acids. Decarboxylases involved in the elimination of the carboxyl group from the aromatic nucleus in these pathways have been reported as non-oxidative (reductive) decarboxylases that do not require the external addition of any cofactor for its activity. Phthalate is metabolized by two different dioxygenase-initiated pathways (see Sect. 2.1.1) either via 4,5-dihydroxyphthalate (in Proteobacteria) or 3,4-dihydroxyphthalate (in Actinobacteria) (Fig. 1). Both dihydroxyphthalate isomers are non-oxidatively decarboxylated to protocatechuate. 4,5-dihydroxyphthalate

14

D. Pérez-Pantoja et al.

Fig. 4 Metabolism of methoxylated aromatic hydrocarbons by Sphingomonas paucimobilis SYK-6. Enzymes catalyzing a given reaction only at slow rate are shown in italics

decarboxylases have been purified from P. fluorescens PHK (Pujar and Ribbons 1985) and C. testosteroni NH1000 (Nakazawa and Hayashi 1978) and show a narrow specificity with only 4,5-dihydroxyphthalate and 4-hydroxyphthalate as substrates (Nakazawa and Hayashi 1978). 4,5-Dihydroxyphthalate decarboxylases described thus far share >78% of sequence identity. 3,4-dihydroxyphthalate 2-decarboxylase from A. keyseri 12B is unrelated to the 4,5-dihydroxyphthalate decarboxylases and its deduced amino acid sequence most closely resembles that of aldolases which catalyze the cleavage of fuculose 1-phosphate (Eaton 2001). Similar 3,4-dihydroxyphthalate 2-decarboxylases (58–69% identity) were observed in other phthalate-degrading Actinobacteria.

Aerobic Degradation of Aromatic Hydrocarbons

15

In S. paucimobilis SYK-6, 5-carboxyvanillate is transformed to vanillate by non-oxidative 5-carboxyvanillate decarboxylases (Fig. 4). Two such decarboxylases, LigW and LigW2, have been identified, which share 37% amino acid sequence identity (Peng et al. 2002, 2005) but exhibit no homology with members of previously described non-oxidative decarboxylase families. It was recently proposed that they can be classified into a new family of non-oxidative aromatic acid decarboxylases, together with 2,6-dihydroxybenzoate decarboxylase of Rhizobium radiobacter (Yoshida et al. 2004), which catalyzes the reversible decarboxylation of 2,6- and 2,3-dihydroxybenzoate to resorcinol and catechol, respectively.

2.2.4

CoA-Dependent Peripheral Pathways for Phenylpropenoid Compounds Phenylpropenoid compounds constitute a common carbon source for plantassociated microorganisms since they are structural components of plant polymers, such as lignin and suberin. Among phenylpropenoid compounds, the largest group is hydroxycinnamates derivatives (i.e., ferulate, coumarate, caffeate and others). In A. baylyi ADP1, a chlorogenate esterase hydrolyzes the ester bond of chlorogenate, an abundant hydroxycinnamic compound, to produce quinate and caffeate (Smith et al. 2003). In this bacterium, like in most other bacteria characterized in the respect, hydroxycinnamate catabolism follows a CoA-dependent non β-oxidative route (Overhage et al. 1999). p-Coumarate, caffeate and ferulate are transformed to 4-hydroxybenzoate, protocatechuate and vanillate, respectively, through the action of enzymes with relatively broad substrate specificity, encoded in A. baylyi by the hcaABC genes (Fig. 5). Hydroxycinnamates are initially activated to the corresponding CoA esters by hydroxycinnamoyl-CoA synthase (HcaC) (also often termed feruloyl-CoA synthase), an ATP dependent (AMP forming) CoA ligase (Fig. 5). A bifunctional enoyl-CoA hydratase/aldolase (HcaA) catalyzes the hydratation but also acts as a lyase cleaving the hydrated derivatives of feruloyl-CoA, caffeoyl-CoA and p-coumaroyl-CoA to acetyl-CoA and vanillin, 3,4-dihydroxybenzaldehyde and 4-hydroxybenzaldehyde, respectively (Gasson et al. 1998). The aldehydes are oxidized by aldehyde dehydrogenases (HcaB) to vanillate, protocatechuate, and 4-hydroxybenzoate, respectively (Fig. 5). 4-Hydroxybenzoate and vanillate are transformed to protocatechuate by 4-hydroxybenzoate 3-hydroxylase or vanillate O-demethylase, respectively. Three alternative modes of ferulate degradation are additionally discussed in the literature, a non-oxidative decarboxylation, discovered mainly in fungi and yeast, side-chain reduction, typical for the anaerobic degradation of ferulate, and a CoA-independent deacetylation (Priefert et al. 2001). However, even in Delftia acidovorans, which had been proposed to carry out such a CoA-independent deacetylation, a CoA-dependent pathway was observed as the major route for ferulate degradation (Plaggenborg et al. 2001). Phenylpropanoid compounds, i.e., saturated derivatives of hydroxycinnamates, such as 4-hydroxyphenylpropionate and 3,4-dihydroxyphenylpropionate are

16

D. Pérez-Pantoja et al.

Fig. 5 Peripheral reactions in the bacterial degradation of ferulate, caffeate, p-coumarate, phenylalanine and tryptophan

assumed to be catabolized by A. baylyi ADP1, using a FAD-dependent acyl-CoA dehydrogenase (HcaD) which dehydrogenates the saturated propionyl-CoA side chain of the hydroxyphenylpropanoyl thioesters produced by HcaC to form

Aerobic Degradation of Aromatic Hydrocarbons

17

hydroxycinnamoyl-CoA thioesters that can be channeled to protocatechuate (Smith et al. 2003).

2.2.5

Peripheral Reactions in the Degradation of Aromatic Amino Acids In eukaryotes, the metabolism of the aromatic amino acids phenylalanine, tyrosine and tryptophan is initiated by tetrahydropterin dependent monooxygenases (Fitzpatrick 2003), where tetrahydropterin serves as electron source to reduce the second atom of oxygen to the level of water. Also in bacteria, phenylalanine is transformed by a pterin-dependent phenylalanine hydroxylase into tyrosine (Nakata et al. 1979) (Fig. 5). A tyrosine aminotransferase catalyzes the conversion of tyrosine into 4-hydroxyphenylpyruvate (Arias-Barrau et al. 2004; Gu et al. 1998), which is further transformed by a 4-hydroxyphenylpyruvate dioxygenase (HPPD) (Fitzpatrick 2003). HPPD is an Fe2+-dependent, non-heme oxygenase that catalyzes the conversion of 4-hydroxyphenylpyruvate to homogentisate (Fig. 5). This reaction involves decarboxylation, substituent migration and aromatic oxygenation in a single catalytic cycle. This enzyme is a member of the α-keto acid dependent oxygenases that typically require an α-keto acid (almost exclusively α-ketoglutarate) and molecular oxygen to either oxygenate or oxidize a third molecule. As an exception in this class of enzymes HPPD has only two substrates, does not use α-ketoglutarate, and incorporates both atoms of oxygen into the aromatic product, homogentisate (Moran 2005). Indications were also given that phenylalanine, in an alternative pathway can be metabolized via 3,4-dihydroxyphenylalanine and protocatechuate (Ranjith et al. 2007). In various bacteria, tryptophan is subject to non-oxidative degradation by a pyridoxal phosphate-dependent tryptophan indole-lyase (tryptophanase) yielding indole, pyruvate and ammonium (Vederas et al. 1978). In some bacteria, the oxidative degradation of exogenous tryptophan via anthranilate has been suggested (Kurnasov et al. 2003a), but details are still scarce. Experimental verification was achieved by functional expression of a Cupriavidus metallidurans putative kynBAU operon. Tryptophan is converted by a heme-containing specific tryptophan 2,3-dioxygenase (KynA) to N-formylkynurenine, from which the formyl group is removed by kynurenine formamidase (KynB) to kynurenine, and kynureninase (KynU) catalyzes the cleavage to anthranilate and alanine (Kurnasov et al. 2003a) (Fig. 5). However, further genome analyses revealed the presence of gene clusters encoding kynurenine monooxygenase together with kynureninase and a 3-hydroxyanthranilate 3,4-dioxygenase (see Sect. 2.3.5.4) (Kurnasov et al. 2003b) indicating kynurenine to be monooxygenated to 3-hydroxykynurenine followed by cleavage to alanine and 3-hydroxyanthranilate by kynureninase (Fig. 5), which accepts both kynurenine and 3-hydroxykynurenine as substrates (Kurnasov et al. 2003b). Thus, tryptophan can be degraded in bacteria via anthranilate, but also via 3-hydroxyanthranilate. How these different pathways contribute to tryptophan degradation in different taxa remains to be established.

18

D. Pérez-Pantoja et al.

2.3

Central Reactions

2.3.1

Intradiol Ring-Cleavage Pathways

The 3-Oxoadipate Pathway The 3-oxoadipate pathway is widely distributed among soil bacteria and plays a central role in the degradation of naturally occurring aromatic hydrocarbons (Harwood and Parales 1996) such as vanillin, p-coumarate, caffeate, mandelate or tryptophan. Two branches of the 3-oxoadipate pathway can be differentiated, the catechol branch and the protocatechuate branch (Fig. 6). In the catechol branch, the metabolism of catechol is initiated by ortho-cleavage catalyzed by catechol-1,2-dioxygenases resulting in the formation of cis,cismuconate, which is subsequently transformed by a muconate cycloisomerase to muconolactone. Muconolactone isomerase shifts the double bond to form 3-oxoadipate-enol-lactone (enol-lactone), the first common intermediate of the catechol and protocatechuate branch (Fig. 6). In the protocatechuate branch, protocatechuate is subject to ortho-cleavage by protocatechuate 3,4-dioxygenases. Like catechol 1,2-dioxygenases, protocatechuate 3,4-dioxygenases are non-heme Fe3+ containing dioxygenases (Fujisawa and Hayaishi 1968). However, in contrast to catechol 1,2-dioxygenases, which are composed of only one type of subunits, protocatechuate 3,4-dioxygenases are composed of two different subunits which, however, share substantial amino acid identity (Yoshida et al. 1976). Even though cycloisomerization is an important step in both branches of the 3-oxoadipate pathway, the enzymes catalyzing the respective reactions are different. Sequence analyses and kinetic studies showed that carboxymuconate cycloisomerases of the protocatechuate branch belong to the fumarase class II family (Williams et al. 1992), a group of enzymes catalyzing 1,2-addition–elimination reactions including aspartase and arginosuccinate lyase. They do not require any metal cofactors for catalytic activity and catalyze a syn-1,2addition-elimination with 4-carboxy-(S)-muconolactone as product. In contrast, the Mn2+ requiring muconate cycloisomerases belong to the enolase superfamily and catalyze an anti-1,2-addition-elimination with (R)-muconolactone as product (Babbitt et al. 1996). In the protocatechuate branch, the 4-carboxymuconolactone produced is transformed to enol-lactone by carboxymuconolactone decarboxylase. From the biochemical point of view, both branches merge at the stage of the enollactone (Fig. 6), however, a remarkable diversity of the 3-oxoadipate pathway in terms of gene organization, type of inducers and regulation mechanism has been observed (Harwood and Parales 1996), such that the pathways genetically converge at different points in different bacteria, or they never converge as in A. baylyi, which contains two independent set of genes encoding isofunctional enzymes for the last three steps of the pathway. The pcaC and pcaD genes, encoding 4-carboxymuconolactone decarboxylase and 3-oxoadipate enol-lactone hydrolase, respectively, catalyzing successive reactions, are, in some cases, fused in a unique pcaL gene. Sequence analysis of pcaL genes reveals that the N-terminal two thirds of the protein are homologous to the enol-lactone hydrolases, whereas the C-terminal

Aerobic Degradation of Aromatic Hydrocarbons

19

Fig. 6 Dendrogram showing the relatedness of intradiol dioxygenases (catechol 1,2-dioxygenases, protocatechuate 3,4-dioxygenases and hydroxybenzoquinol 1,2-dioxygenases). Reactions catalyzed by enzymes indicated are given to the exterior of the figure, together with subsequent reactions channeling the ring-cleavage products into the Krebs cycle. Among catechol 1,2-dioxygenases, two different lineages can be differentiated, observed in Proteobacteria and Actinobacteria, respectively (Eulberg et al. 1997). Reactions of catechol pathway enzymes with 4-methylcatechol resulting in the formation of 4-methylmuconolactone are indicated to the left of the figure

third is homologous to the decarboxylases (Eulberg et al. 1998). As these gene fusions are present in distantly related bacteria a biochemical advantage of these fused gene products is possible (Pérez-Pantoja et al. 2008). Enol-lactone is

20

D. Pérez-Pantoja et al.

hydrolyzed by enol-lactone hydrolases which are assumed to use a Ser-His-Asp catalytic triade (Schlömann 1994). 3-Oxoadipate in turn is transformed by 3-oxoadipate:succinyl-CoA transferase and 3-oxoadipyl-CoA thiolase to Krebs cycle intermediates (Gobel et al. 2002).

Metabolism of Methylaromatics via Intradiol Cleavage Usually, the 3-oxoadipate pathway is not suited for the degradation of methylaromatics because methylsubstituted muconolactones, formed by the action of catechol 1,2-dioxygenase and muconate cycloisomerase, accumulate as dead-end products (Catelani et al. 1971). In the case of transformation of 4-methylcatechol, 4-methylmuconolactone (4-ML) is formed (Fig. 6), which cannot be processed by enzymes of the 3-oxoadipate pathway as no proton is available to be abstracted by the muconolactone isomerase (Pieper et al. 1985). However, in C. necator JMP134 and R. rhodochrous N75, a 4-methylmuconolactone methylisomerase capable of converting 4-ML to 3-methylmuconolactone (3-ML) was described which compensates for the initial “incorrect” cycloisomerization of 3-methylmuconate (Bruce et al. 1989; Pieper et al. 1990). In C. necator JMP134, 3-ML is further metabolized by a methylmuconolactone isomerase and via 4-methyl-3-oxoadipate, with reactions analogous to those of the classical 3-oxoadipate pathway (Prucha et al. 1997) being encoded by the mml gene cluster (Marin et al. 2010). In R. rhodochrous N75 hydrolysis of the lactone ring obviously occurs from 3-methylmuconolactone-CoA (Cha et al. 1998).

Metabolism of 1,2,4-Trihydroxybenzene Hydroxybenzoquinol (1,2,4-trihydroxybenzene) is the central intermediate in the degradation of a variety of aromatic hydrocarbons such as resorcinol (Huang et al. 2006), 4-aminophenol (which is assumed to be degraded via 1,4-benzenediol, (Takenaka et al. 2003)) or 4-hydroxysalicylate (Armengaud et al. 1999) including a variety of particularly recalcitrant polychloro- and nitroaromatic pollutants (chapter 5, Vol. 2, Part 2). Hydroxybenzoquinol 1,2-dioxygenase is the key enzyme of hydroxybenzoquinol metabolism and catalyzes the intradiol cleavage to form 3-hydroxy-cis,cis-muconate and its tautomer, maleylacetate (Fig. 6). Hydroxybenzoquinol 1,2-dioxygenases have been purified and characterized from a variety of microorganisms (Takenaka et al. 2003), and also crystallized (Ferraroni et al. 2005). Hydroxybenzoquinol 1,2-dioxygenases are usually highly specific for hydroxybenzoquinol and do not, or relatively slowly, convert catechol. In accordance, in a dendrogram of intradiol dioxygenases, hydroxybenzoquinol and catechol 1,2-dioxygenases clustered in separated branches (Fig. 6). The next enzyme of the hydroxyquinol pathway, maleylacetate reductase, performs the reduction of the carbon-carbon double bond to channel maleylacetate into the 3-oxoadipate pathway. Maleylacetate reductases have previously been described as important key enzymes of chloroaromatics but also nitroaromatic degradation.

Aerobic Degradation of Aromatic Hydrocarbons

21

2.3.2 Catechol Meta-Cleavage Pathways The extradiol ring-cleavage of catechol and methylsubstituted catechols is typically catalyzed by type I extradiol dioxygenases (catechol 2,3-dioxygenases, C23O) which belong to the vicinal oxygen chelate family enzymes (Gerlt and Babbitt 2001). Type I extradiol dioxygenases are also involved in the degradation of biphenyl (2,3-dihydroxybiphenyl 1,2-dioxygenases) or naphthalene. All these extradiol dioxygenases, use non-heme Fe2+ for cleavage (Eltis and Bolin 1996; Harayama and Rekik 1989). However, Mn2+ dependent extradiol dioxygenases with high sequence similarity to the Fe2+ dependent enzymes have also been reported (Hatta et al. 2003). Eltis and Bolin (Eltis and Bolin 1996) analyzed in detail the phylogenetic relationships among type I extradiol dioxygenases and described them as a superfamily which can be divided into different families and subfamilies. Families I.2 and I.3 consist of two-domain iron-containing enzymes that show preferences for monocyclic and bicyclic substrates, respectively, whereas family I.1 comprises the small single domain enzymes identified in R. globerulus P6 and Sphingomonas sp. strain BN6. In the last years, the description of new members of this family increased significantly (Vaillancourt et al. 2006; Duarte et al. 2014). Extradiol dioxygenases can be subject to a rapid oxidation of the active site ferrous iron into its ferric form with concomitant loss of activity (Vaillancourt et al. 2002), specifically during turnover of certain substrates such as 4-methylcatechol. Small auxiliary ferredoxin proteins, whose genes are frequently encoded adjacently to the C23O genes, have been reported to have a reactivating function through reduction of the iron atom in the active site of the enzyme (Hugo et al. 1998). Two branches of the meta-cleavage pathway for catechols have been described, the hydrolytic and the oxalocrotonate branch, which are often encoded in one single gene cluster (Harayama et al. 1987). In the oxalocrotonate branch, 2-hydroxymuconic semialdehyde is oxidized to 2-hydroxymuconate by 2-hydroxymuconic semialdehyde dehydrogenase, followed by isomerization to oxalocrotonate through the action of oxalocrotonate isomerase and decarboxylation by oxalocrotonate decarboxylase to 2-hydroxypent-2,4-dienoate, the common intermediate of the hydrolytic and the 4-oxoalocrotonate branch (Fig. 7). Both 2-hydroxymuconic semialdehyde (from catechol) and 5-methyl-2hydroxymuconic semialdehyde (from 4-methylcatechol) are preferentially degraded via the oxalocrotonate branch (Harayama et al. 1987). Since 2-hydroxy-6-oxo-2,4heptadienoate, the ring-cleavage product of 3-methylcatechol is a ketone, rather than an aldehyde, it cannot be further oxidized by the 2-hydroxymuconic semialdehyde dehydrogenase and is exclusively metabolized via the hydrolytic route (Powlowski and Shingler 1994). Hydrolysis of 2-hydroxy-6-oxo-2,4-heptadienoate by 2-hydroxymuconic semialdehyde hydrolase gives rise to 2-hydroxypent-2, 4-dienoate and acetate (Fig. 7). The final steps of the catechol meta-cleavage pathway are catalyzed by 2-hydroxypent-2,4-dienoate hydratase (to give 4-hydroxy-2-oxovalerate), 4-hydroxy-2-oxovalerate aldolase (to give acetaldehyde and pyruvate) and acetaldehyde dehydrogenase (decycling) that converts acetaldehyde to acetyl-CoA.

Fig. 7 Extradiol ring-cleavage pathways involved in the degradation of catechol, 3-methylcatechol, protocatechuate, homoprotocatechuate, 2,3-dihydroxyphenylpropionate, 2,3-dihydroxycinnamate, 2-aminophenol, 3-hydroxyanthranilate and 4-amino-3-hydroxybenzoate. Unstable intermediates are shown in brackets. In case of catechol only metabolism via the oxalocrotonate branch is indicated

22 D. Pérez-Pantoja et al.

Aerobic Degradation of Aromatic Hydrocarbons

23

Fig. 8 Dendrogram showing the relatedness of type II (LigB) extradiol dioxygenases. The physiological function of BphC6 2,3-dihydroxybiphenyl 1,2-dioxygenase from R. rhodochrous K37 remains to be established, but is possibly involved in fluorene degradation. (Taguchi et al. 2004)

2.3.3 Protocatechuate Meta-Cleavage Pathways Like for catechol, protocatechuate can be metabolized via intradiol or extradiol cleavage pathways. As protocatechuate has an asymmetric structure, meta-cleavage can occur in the 2,3- but also in the 4,5-position. Most microorganisms seem to perform a 4,5-cleavage (Ono et al. 1970). A protocatechuate 2,3-dioxygenase has been described from Bacillus macerans (Wolgel et al. 1993), however, no further information on this pathway is yet available. Protocatechuate 4,5-cleavage is catalyzed by heteromultimeric protocatechuate 4,5-dioxygenases, with the two subunits being unrelated (Sugimoto et al. 1999). The active site comprises, like in C23O, a Fe2+ ion located in the β-subunit (Sugimoto et al. 1999). However, protocatechuate 4,5-dioxygenase is unrelated to above described C23Os and belongs to the type II or LigB superfamily of extradiol dioxygenases (Vaillancourt et al. 2006; Duarte et al. 2014) (Fig. 8). The protocatechuate 4,5-dioxygenolytic ring-cleavage product 4-carboxy-2hydroxymuconate-6-semialdehyde is non enzymatically converted to an intramolecular hemiacetal form and then oxidized by a 4-carboxy-2-hydroxymuconic semialdehyde dehydrogenase (Fig. 7) which, like cis-4,5-dihydro-4,5dihydroxyphthalate dehydrogenase (see Sect. 2.1.1), belongs to the GFO/IDH/

24

D. Pérez-Pantoja et al.

MOCA family (Chang and Zylstra 1998). The resulting intermediate, 2-pyrone-4,6dicarboxylate, is hydrolyzed by 2-pyrone-4,6-dicarboxylate hydrolase (Maruyama 1983) to yield the keto form and enol form (4-carboxy-2-hydroxymuconate) of 4-oxalomesaconate, which are in equilibrium. 2-Pyrone-4,6-dicarboxylate hydrolase was postulated to contain a catalytically active cysteine as part of a catalytic triad (Masai et al. 1999b), however, further analysis on related proteins indicates that this hydrolase might be a metal-dependent hydrolase (Halak et al. 2007). The recent elucidation of the crystal structure might shed some light on the enzyme mechanism. 4-Oxalomesaconate is converted to 4-carboxy-4-hydroxy-2-oxoadipate by 4-oxalomesaconate hydratase (Hara et al. 2000). Finally, 4-carboxy-4-hydroxy-2oxoadipate is cleaved by 4-carboxy-4-hydroxy-2-oxoadipate aldolase to produce pyruvate and oxaloacetate (Fig. 7).

2.3.4

Further Meta-Cleavage Routes Involving Type I or Type II Extradiol Dioxygenases

The Homoprotocatechuate Pathway Homoprotocatechuate (3,4-dihydroxyphenylacetate) is a central intermediate in the degradation of 4-hydroxyphenylacetate and the aromatic amines tyramine and dopamine. So far, degradation of homoprotocatechuate has been exclusively described by extradiol cleavage through homoprotocatechuate 2,3-dioxygenases. Proteobacterial homoprotocatechuate 2,3-dioxygenases as the one described from E. coli C (Roper and Cooper 1990) belong to the type II or LigB superfamily of extradiol dioxygenases (Fig. 8; Duarte et al. 2014). In contrast, actinobacterial homoprotocatechuate 2,3-dioxygenases like the Fe2+ dependent enzyme from Brevibacterium fuscum or the Mn2+ dependent enzyme from Arthrobacter globiformis belong to the type I extradiol dioxygenases (Vetting et al. 2004). Independent of the type of reaction, 5-carboxymethyl-2-hydroxymuconic semialdehyde is the reaction product (Fig. 7). In E. coli, the further metabolism follows a dehydrogenative route with dehydrogenation to the acid by 5-carboxymethyl-2-hydroxymuconic semialdehyde dehydrogenase, which exhibits significant sequence identity (40%) to the respective 2-hydroxymuconic semialdehyde dehydrogenases involved in catechol degradation (Diaz et al. 2001). Isomerization of 5-carboxymethyl-2-hydroxymuconate to 5-oxopent-3-ene-1,2,5-tricarboxylic acid is catalyzed by an isomerase in a reaction similar to that performed by 4-oxalocrotonate tautomerase (see Sect. 2.3.2), however, the two enzymes do not have any apparent sequence similarity (Diaz et al. 2001). A bifunctional decarboxylase/isomerase catalyzes the magnesium-dependent decarboxylation of 5-oxopent-3-ene-1,2,5-tricarboxylate to 2-oxo-hept-3-ene-1,7-dioate. This reaction is followed by a hydratase giving rise to 2,4-dihydroxyhept-2-ene-1,7-dioate and an aldolase forming pyruvate and succinic semialdehyde (Fig. 7).

Aerobic Degradation of Aromatic Hydrocarbons

25

The 2,3-Dihydroxyphenylpropionate Pathway Extradiol ring-cleavage is also involved in the metabolism of 3-hydroxyphenylpropionate and 3-hydroxycinnamate via 2,3-dihydroxyphenylpropionate or 2,3-dihydroxycinnamate, respectively. Like proteobacterial 3,4-dihydroxyphenylacetate 2,3-dioxygenases, proteobacterial 2,3-dihydroxyphenylpropionate 1,2-dioxygenases belong to the LigB superfamily of extradiol dioxygenases (Diaz et al. 2001; Duarte et al. 2014) (Fig. 8). Similarly, actinobacterial 2,3-dihydroxyphenylpropionate 1,2-dioxygenases also belongs to this superfamily (Barnes et al. 1997). 2,3-dihydroxyphenylpropionate 1,2-dioxygenases show a broad specificity with 2,3-dihydroxycinnamate as a good substrate (Spence et al. 1996). Also catechol and methylcatechols are usually accepted as substrates (Barnes et al. 1997; Diaz et al. 2001). The ring-cleavage product 2-hydroxy-6-ketonona-2,4-diene-1,9-dioate is further degraded through a hydrolytic route generating succinate and 2-hydroxypent-2,4dienoate (Fig. 7). The respective hydrolases show some substrate selectivity for the carboxylate of the side chain with only slow turnover of the ring fission products of 3-methylcatechol or catechol (Diaz et al. 2001). However, the ring fission product of 2,3-dihydroxycinnamate was a fairly efficient substrate, generating fumarate and 2-hydroxypent-2,4-dienoate (Barnes et al. 1997; Lam and Bugg 1997) (Fig. 7). Significant sequence similarity has been detected between 2-hydroxy-6-ketonon2,4-diene-1,9-dioate hydrolases and other C-C bond hydrolases cleaving vinylogous 1,5-diketones such as those involved in the degradation of 2,3-dihydroxybiphenyl or catechol (see Sect. 2.3.2). 2-Hydroxy-6-ketonon-2,4-diene-1,9-dioate hydrolases of Proteobacteria seem to be most closely related to hydrolases involved in 2,3-dihydroxybiphenyl degradation, and share only approx. 30% sequence identity with 2-hydroxy-6-ketonon-2,4-diene-1,9-dioate hydrolases of Actinobacteria. Further metabolism of 2-hydroxypent-2,4-dienoate occurs as described above (see Sect. 2.3.2). Degradation of Gallate Only poor information is available so far on the metabolism of gallate (3,4,5trihydroxybenzoate). In S. paucimobilis SYK-6, it has been described that syringate is metabolized via 3-O-methylgallate (Masai et al. 1999a). The protocatechuate 4,5-dioxygenase of this strain was reported to catalyze ring-cleavage also of 3-O-methylgallate with the direct formation of 2-pyrone-4,6-dicarboxylate, a metabolite of protocatechuate degradation (see Sect. 2.3.3) (Fig. 4). However, further analysis revealed the presence of a novel extradiol dioxygenase of the LigB family termed DesZ to be responsible for 3-O-methylgallate metabolism (Kasai et al. 2004). This enzyme also transforms gallate, but is practically inactive with protocatechuate (Fig. 4). However, 3-O-methylgallate formed from syringate can be subject to an initial demethylation in strain SYK-6, and a third extradiol dioxygenase of the LigB family, termed DesB, could be identified as being highly specific for cleavage of gallate, and being inactive with either 3-O-methylgallate or protocatechuate (Kasai et al. 2005). Oxalomesaconate was identified as ring-cleavage product (Fig. 4). A similar gallate dioxygenase was also identified in P. putida KT2440 (Nogales

26

D. Pérez-Pantoja et al.

et al. 2005) (Fig. 8). A more detailed analysis of the primary structure of gallate dioxygenases revealed that the N-terminal regions showed a significant amino acid sequence identity with the β-subunit of protocatechuate 4,5-dioxygenases, whereas the C-terminal region has similarity to the corresponding small α-subunit (Nogales et al. 2005). It was therefore suggested that gallate dioxygenases are two-domain proteins that have evolved from the fusion of large and small subunits of protocatechuate 4,5-dioxygenases. Degradation of 2-aminophenol Usually, extradiol dioxygenases necessitate the presence of two neighbored hydroxyl-substituents on the substrate. However, analysis of the metabolism of 2-aminophenol revealed that this substrate can be directly cleaved by extradiol dioxygenases of the LigB superfamily, termed 2-aminophenol 1,6-dioxygenases (Takenaka et al. 1997) (Fig. 8). These enzymes are composed of two subunits, which share sequence similarity (Fig. 8) but it appears that only the β-subunit contains an active site. Catechol is only a poor substrate for the enzyme (Takenaka et al. 1997). The further metabolism of the formed 2-aminomuconic semialdehyde occurs in analogy to the metabolism of 2-hydroxymuconic semialdehyde produced during catechol extradiol cleavage (Fig. 7). 2-Aminomuconic semialdehyde dehydrogenases share significant sequence similarity (up to 60%) with 2-hydroxymuconic semialdehyde dehydrogenases and the enzyme of P. pseudoalcaligenes JS45 was shown to be capable to transform 2-hydroxymuconic semialdehyde (He et al. 1998). Aminomuconate is hydrolyzed by aminomuconate deaminase to 4-oxalocrotonate in strain JS45, which indicates that deamination is carried out via an imine intermediate (He and Spain 1998). Aminomuconate deaminase was also observed in the degradation of 2-aminophenol by other Pseudomonas strains (Takenaka et al. 2000). Further degradation of 4-oxalocrotonate proceeds through reactions as described above (see Sect. 2.3.2) (Fig. 7). A different aminomuconate deaminating activity was recently observed in Comamonas strain CNB-1 (Liu et al. 2007) and it was suggested that 2-hydroxymuconate rather than oxalocrotonate is the deamination product. Extradiol Cleavage of Benzoquinol Pathways that involve the cleavage of benzoquinol have been suggested to be involved in the degradation of 4-hydroxyphenoxyacetate (Crawford 1978), 4-ethylphenol and 4-hydroxyacetophenone (Darby et al. 1987), but also of chloroor nitroaromatics. Chlorobenzoquinol (and also benzoquinol) produced during the catabolism of γ-HCH (hexachlorocyclohexane) in S. japonicum UT26 is subject to direct ring cleavage by LinE extradiol dioxygenase, a type I extradiol dioxygenase. In contrast, benzoquinol 1,2-dioxygenase from the 4-hydroxyacetophenonedegrading P. fluorescens ACB has been shown to be an α2β2 heterotetramer where the α- and β-subunits displayed no significant sequence identity with known dioxygenases. The enzyme is thus the prototype of a novel class of Fe2+-dependent dioxygenases (Moonen et al. 2008b). The enzyme not only cleaves benzoquinol to

Aerobic Degradation of Aromatic Hydrocarbons

27

form 4-hydroxymuconic semialdehyde but also a wide range of substituted benzoquinols to the corresponding 4-hydroxymuconic semialdehyde derivatives. In P. fluorescens ACB, the subsequent conversion of 4-hydroxymuconic semialdehyde to maleylacetate is accomplished by a 4-hydroxymuconic semialdehyde dehydrogenase, which exhibits moderate sequence identity (37–43%) to the respective 2-hydroxymuconic semialdehyde dehydrogenases involved in meta-cleavage pathways of catechol (Moonen et al. 2008a). Maleylacetate is transformed to 3-oxoadipate by maleylacetate reductase to be channeled to Krebs cycle intermediates.

2.3.5

Pathways Involving Extradiol Ring-Cleavage by Enzymes of the Cupin Superfamily Proteins of the cupin superfamily share a common architecture and the term cupin (from the latin term “cupa,” for a small barrel or cask) has been given to a beta barrel structural domain (Dunwell et al. 2000). Members of this superfamily share two histidine containing sequence motifs that identify the binding site of the metal. Various extradiol dioxygenases of aromatic degradation pathways (termed type III) have been described to belong to this superfamily (Duarte et al. 2014). It should be noted that even though belonging to different families, all three types of extradiol dioxygenases share similar active sites and all type I, type II and various type III enzymes have the same iron ligands, two histidine and one glutamate, that constitute the 2-His-1-carboxylate structural motif (Vaillancourt et al. 2006). The Gentisate Pathway Gentisate and substituted gentisates serve as the focal point in the aerobic biodegradation of a large number of simple and complex aromatic hydrocarbons such as salicylate, 3-hydroxybenzoate, 3,5- or 2,5-xylenol or naphthalene. Consequently, the gentisate pathway is distributed throughout the bacterial world. In this pathway, gentisate 1,2-dioxygenase, a member of the cupin superfamily, cleaves the aromatic ring between the carboxyl substituent and the proximal hydroxyl group to yield maleylpyruvate (Crawford et al. 1975) (Fig. 9). Gentisate 1,2-dioxygenases have been purified and characterized from various Proteobacteria and Actinobacteria (Crawford et al. 1975; Suemori et al. 1993), and even archaea (Fu and Oriel 1998), and are described as two-domain bicupins (Dunwell et al. 2000). The gentisate 1,2-dioxygenases reported to date have demonstrated reasonably broad substrate tolerance in terms of substitutions on the aromatic ring catalyzing the turnover of a range of alkyl- and halo-substituted gentisates (Harpel and Lipscomb 1990). Rates similar to that observed with gentisate are reported with C-3 substituted gentisates (methyl, ethyl, 2-propyl, bromo, fluoro), whereas C-4 substituted gentisates are turned over at reduced rates (Crawford et al. 1975; Harpel and Lipscomb 1990). All three functional groups of gentisate appear to be required for efficient turnover. However derivatives having a substitution of the carboxyl-group by hydroxyl, acetyl, or aldehyde functions are slowly metabolized (Lipscomb and Orville 1992). Maleylpyruvate produced by gentisate 1,2-dioxygenase is converted to Krebs cycle intermediates via two downstream routes, a direct hydrolytic cleavage to

28

D. Pérez-Pantoja et al.

Fig. 9 Metabolism of gentisate, homogentisate and salicylate via 1,2-dioxygenolytic cleavage.

pyruvate and maleate by maleylpyruvate hydrolase (Hopper et al. 1971) or isomerization to fumarylpyruvate and subsequent hydrolytic cleavage to fumarate and pyruvate by fumarylpyruvate hydrolase (Crawford and Frick 1977) (Fig. 9). In the latter pathway, isomerization of maleylpyruvate to fumarylpyruvate is catalyzed by either a glutathione (GSH)-dependent maleylpyruvate isomerase almost exclusively found in gram negative bacteria (Crawford et al. 1975), or a GSH-independent maleylpyruvate isomerase that has been characterized in various gram-positive bacteria (Crawford and Frick 1977; Shen et al. 2005). Sequence analysis of the GSH-dependent and -independent maleylpyruvate isomerases revealed that the two isomerases were neither homologous nor phylogenetically related (Shen et al. 2005). The Homogentisate Pathway Homogentisate is the central metabolite formed during degradation of aromatic amino acids phenylalanine and tyrosine in several microorganisms (see Sect. 2.2.5). Its further metabolism is initiated by homogentisate 1,2-dioxygenase, which perform the ring cleavage between the acetyl substituent and the proximal hydroxyl group to yield maleylacetoacetate in a manner analogous to that of gentisate 1,2-dioxygenase (Harpel and Lipscomb 1990; Titus et al. 2000) (Fig. 9). Like gentisate 1,2-dioxygenase, also homogentisate 1,2-dioxygenase is a type III extradiol dioxygenases of the two-domain bicupins, that usually contain a single active site in one of two domains, with the other domain remaining as a non-catalytic vestigial remnant (Vaillancourt et al. 2004). The downstream catabolism of maleylacetoacetate is analogous to that described for maleylpyruvate in the gentisate

Aerobic Degradation of Aromatic Hydrocarbons

29

pathway (Fig. 9). It can be hydrolyzed directly to acetoacetate and maleate by maleylacetoacetate hydrolase (Crawford 1976) or, obviously more commonly, be isomerized by a GSH-independent (Suemori et al. 1996) or GSH-dependent isomerase (Crawford and Frick 1977) to fumarylacetoacetate which is finally hydrolyzed to acetoacetate and fumarate by fumarylacetoacetate hydrolase (Arias-Barrau et al. 2004; Crawford and Frick 1977). Direct Cleavage of Salicylate by Salicylate 1,2-Dioxygenase A ring fission dioxygenase which cleaves salicylate between the carboxyl group and the hydroxyl group to form 2-oxohepta-3,5-dienedioate has been described in the naphthalenesulfonate-degrading strain Pseudaminobacter salicylatoxidans BN12 (Hintner et al. 2001). Similarly, 1-hydroxy-2-naphthoate dioxygenase from Nocardioides sp. KP7 (Iwabuchi and Harayama 1998) involved in the degradation of phenanthrene by this strain was shown to be capable to cleave between a carboxyl and a hydroxyl group, contradicting a generally accepted paradigm that the enzymatic ring fission of the aromatic nucleus by bacteria requires the presence of two hydroxyl groups or one amino and one hydroxyl group (Fig. 9). In addition to salicylate, salicylate 1,2-dioxygenase also converts gentisate and a wide range of substituted salicylates (Hintner et al. 2001). The deduced amino acid sequence revealed that salicylate-1,2-dioxygenase also belongs to the type III extradiol dioxygenases with a subunit topology characteristic of the bicupin beta-barrel folds (Matera et al. 2008). The crystal structure revealed, however, that this enzyme does not contain the classical 2-His-1-carboxylate metal-binding motif but a mononuclear iron center involving three histidine ligands, the iron coordination being completed by water molecules (Matera et al. 2008). The downstream pathway of 2-oxohepta-3,5-dienedioate is still elusive. The 3-Hydroxyanthranilate Pathway 3-Hydroxyanthranilate is a central intermediate of tryptophan degradation via the kynurenine pathway (see Sect. 2.2.5) (Fig. 5) and of the biosynthetic pathway from tryptophan to quinolinate, the universal de novo precursor to the pyridine ring of nicotinamide adenine dinucleotide. In this pathway, 3-hydroxyanthranilate 3,4-dioxygenase catalyzes the conversion of 3-hydroxyanthranilate to 2-amino-3carboxymuconic semialdehyde (Muraki et al. 2003) (Fig. 7). 3-Hydroxyanthranilate 3,4-dioxygenase also belongs to the type III extradiol dioxygenases, but in contrast to gentisate 1,2-dioxygenase, it is composed of a single cupin domain (Zhang et al. 2005). The ring-cleavage product 2-amino-3-carboxymuconic semialdehyde is decarboxylated to 2-aminomuconic semialdehyde (Muraki et al. 2003), a common intermediate with the 2-aminophenol metabolic pathway (see Sect. 2.3.4.4) (Fig. 7). The 4-Amino-3-Hydroxybenzoate Pathway A 4-amino-3-hydroxybenzoate 2,3-dioxygenase that catalyzes the ring fission between C2 and C3 yielding 2-amino-5-carboxymuconic semialdehyde has been isolated from Bordetella sp. 10d (Takenaka et al. 2002). This Fe2+ dependent enzyme is highly specific and neither 2-aminophenol, nor its methyl-, hydroxyl- or carboxyl- derivatives, including 3-hydroxyanthranilate are substrates. The deduced

30

D. Pérez-Pantoja et al.

amino acid sequence shows significant identity (28%) with 3-hydroxyanthranilate 3,4-dioxygenases indicating that this enzyme also belongs to the cupin superfamily (Murakami et al. 2004). Further metabolism of 2-amino-5-carboxymuconic semialdehyde is assumed to proceed via enzyme catalyzed deamination to 2-hydroxy-5-carboxymuconic semialdehyde followed by spontaneous decarboxylation to yield 2-hydroxymuconic semialdehyde (Orii et al. 2004). Subsequent catabolism of 2-hydroxymuconic semialdehyde occurs via a dehydrogenative route as described above (see Sect. 2.3.2) (Fig. 7).

2.4

CoA Dependent Pathways

The involvement of CoA-dependent reactions in aromatic hydrocarbon degradation is known since decades. However, such an involvement was thought to be restricted to an oxidation of side-chains or reactions funneling ring-cleavage products into the Krebs cycle. The cleavage of the aromatic ring of CoA-substituted derivatives was assumed to be restricted to anaerobic pathways. However, an aerobic route for degradation of aromatic hydrocarbons without involvement of dihydroxylated aromatic intermediates was initially reported for phenylacetate degradation in E. coli W (Ferrandez et al. 1998) and P. putida U (Olivera et al. 1998) (Fig. 10). The initial step of the pathway involves the activation of phenylacetate into phenylacetyl-CoA by a phenylacetate-CoA ligase (Mohamed 2000). Like other CoA ligases (see Sect. 2.2.4), this enzyme belongs to the AMP-forming acyl-CoA ligases, which catalyze thioesterification via a two-step process in which an acyladenosine monophosphate (AMP) intermediate is formed in the first step, followed by formation of the acyl-CoA ester and release of AMP. Phenylacetyl-CoA is attacked by a ring-oxygenase/reductase (the PaaABCDE gene products), generating a hydroxylated and reduced derivative of phenylacetyl-CoA, probably 1,2-dihydroxy-1,2-dihydrophenylacetyl-CoA, which is not re-oxidized to a dihydroxylated aromatic intermediate, as in other known aromatic pathways (Ismail et al. 2003) (Fig. 10). Sequence comparisons of the paaABCDE gene products strongly suggest that the oxygenase belongs to the bacterial diiron multicomponent oxygenases family and suggest that PaaACD might constitute the α, β, and y subunits of the heteromultimeric diiron oxygenase component of the oxygenase. PaaB and PaaE may be the effector protein and the oxidoreductase, respectively, that mediate electron transfer from NAD(P)H (Fernandez et al. 2006). Interestingly, although all bacterial diiron multicomponent oxygenases described so far are monooxygenases, the proposed product of the reaction catalyzed by the oxygenase is a dihydrodiol, and therefore this enzyme could be a hydroxylating dioxygenase (Ismail et al. 2003). It has been proposed that 1,2-dihydroxy-1,2dihydrophenylacetyl-CoA, is further metabolized in a complex reaction sequence comprising enoyl-CoA isomerization/hydration, non-oxygenolytic ring opening and dehydrogenation, which is catalyzed by the PaaG and PaaZ gene products. The resulting aliphatic CoA dicarboxylate compound is further catabolized by a

Aerobic Degradation of Aromatic Hydrocarbons

31

Fig. 10 Metabolism of phenylacetate, benzoate and 2-aminobenzoate by CoA-dependent aerobic pathways. Reaction intermediates are shown in brackets. The transformation of 3-hydroxyadipylCoA to 3-oxoadipyl-CoA has been proven to be involved in phenylacetate degradation. Tentative reactions are indicated by a question mark

32

D. Pérez-Pantoja et al.

β-oxidation-like pathway via β-ketoadipyl-CoA (Ismail et al. 2003) (Fig. 10) and a β-ketoadipyl-CoA thiolase that catalyses the last step of the phenylacetate catabolic pathway, i.e., the thiolytic cleavage of beta-ketoadipyl-CoA to succinyl-CoA and acetyl-CoA (Nogales et al. 2007). Also benzoate, a strategic intermediate in aerobic aromatic hydrocarbon metabolism, can be metabolized aerobically via benzoyl-CoA, involving also non-oxygenolytic ring cleavage (Altenschmidt et al. 1993) (Fig. 10). The benzoate-CoA ligase of B. xenovorans LB400, which also belongs to the AMP-forming acyl-CoA ligases, has been analyzed in detail and shows some activity with 2-aminobenzoate, but is inactive with phenylacetate (Bains and Boulanger 2007). Benzoyl-CoA is hydroxylated by benzoyl-CoA oxygenase/reductase, a two component benzoyl-CoA dioxygenase, forming 2,3-dihydro-2,3-dihydroxybenzoyl-CoA (Zaar et al. 2004) (Fig. 10). Benzoyl-CoA dioxygenase is composed of an ironsulfur-flavoprotein reductase (BoxA) and an oxygenase (BoxB) which shows low similarity to PaaA, the supposed α subunit of the heteromultimeric diiron phenylacetyl-CoA oxygenase. The dihydrodiol is the substrate for ring fission catalyzed by dihydrodiol lyase (BoxC) (Gescher et al. 2005), a member of the enoyl-CoA hydratase/isomerase superfamily. This homodimeric enzyme does not require oxygen and catalyzes the transformation to 3,4-dehydroadipyl-CoA semialdehyde. The latter intermediate is subsequently oxidized by 3,4-dehydroadipylCoA semialdehyde dehydrogenase (BoxD) to 3,4-dehydroadipyl-CoA (Gescher et al. 2006) (Fig. 10). The further metabolism is thought to lead to 3-oxoadipylCoA, which is finally cleaved into succinyl-CoA and acetyl-CoA (Zaar et al. 2004). An on the first view similar pathway has also been reported for the aerobic metabolism of anthranilate (2-aminobenzoate) via 2-aminobenzoyl-CoA (Altenschmidt and Fuchs 1992). Even though thioesterification is catalyzed by a 2-aminobenzoate CoA ligase with similarity to benzoate CoA ligase, oxygenation is catalyzed by a 2-aminobenzoyl-CoA monooxygenase/reductase rather than a diiron oxygenase (Buder and Fuchs 1989). This enzyme catalyzes both monooxygenation and hydrogenation of 2-aminobenzoyl-CoA to form 2-amino-5-oxocyclohex-1enecarboxyl-CoA via 2-amino-5-oxocyclohex-1,3-dienecarboxyl-CoA (Schuhle et al. 2001) (Fig. 10). Sequence analysis revealed that the N-terminal part shows similarities to single component flavin monooxygenases and the C-terminal part to NADH dependent, flavin-containing oxidoreductases of the old-yellow-enzyme type (Schuhle et al. 2001). Further metabolism is assumed to proceed by β-oxidation, however, the metabolic pathway remains to be elucidated. Interestingly, novel CoA dependent pathways are still being discovered. In Streptomyces WA-46 salicylate was described to be subject to initial thioesterification and the formed salicylyl-CoA hydroxylated by salicylyl-CoA 5-hydroxylase to gentisyl-CoA (Ishiyama et al. 2004). Gentisyl-CoA was supposed to spontaneously decompose to CoA and gentisate, which was subject to ring-cleavage by a gentisate 1,2-dioxygenase. However, also gentisyl-CoA thioesterases were recently described (Zhuang et al. 2004).

Aerobic Degradation of Aromatic Hydrocarbons

3

33

Research Needs

Various main routes for microbial aerobic degradation of aromatic hydrocarbons are known. However, novel catabolic pathways are still being discovered, indicating the broad and still poorly understood diversity of microbial capabilities. Also the links between aerobic and anaerobic degradation of aromatic hydrocarbons are poorly described. Changing environments are common for microorganisms, especially bacteria, therefore oxygen availability may control the way a particular aromatic hydrocarbon is being degraded. In this context, CoA dependent pathways may play an important role. Another important research need is to broaden the knowledge on the range and type of peripheral reactions that microorganisms can perform. Interestingly, even the metabolism of abundant aromatics, such as the amino acid tryptophan is still poorly understood. In view of the impressive variety of natural aromatic products, especially those produced by plants, this unexplored diversity can be an important source of enzymes for transformation to valuable products. Special efforts should be directed towards a better understanding of O-demethylation reactions, because the number of methoxylated aromatic hydrocarbons known is far greater than the methoxylated aromatic hydrocarbons known to be degraded. It would be also important to better understand why some compounds are degraded by different peripheral or central pathways. For example, benzoate can be degraded by the classical ortho ring cleavage pathway, but also by a CoA dependent pathway; toluene can be degraded by direct oxygenation of the aromatic ring and also by oxidation of the methyl substituent; catechol and protocatechuate can be degraded through ortho or meta ring cleavage pathways. Is the particular pathway controlled by physiological constraints at the cell level? Is it controlled at the species or population level or by environmental factors, such as oxygen or iron availability, as suggested for benzoate degradation pathways? As it is thoroughly demonstrated in this chapter, an impressive diversity of oxygenases plays a significant role at different stages in aerobic aromatic hydrocarbon degradation. Several aspects concerning oxygenases should be addressed. Substrate specificity is a key to allow these enzymes to use different compounds as substrates. Narrow specificity decreases the impact of a particular oxygenase in aerobic degradation whereas broad specificity, in principle, provides an advantage allowing the microorganism to degrade a wider range of (potential) carbon and energy sources. However, this potential advantage contrast with problems associated to dead-end product formation and, more important, intermediate misrouting. Additional biochemical studies, especially those related with regioselectivity, and genetic studies, i.e., inducer of the genes encoding oxygenases, are clearly required. In addition, ongoing microbial genome sequencing projects clearly indicate the presence of sequences putatively encoding oxygenases that do not match, or cannot be associated with the pathways, which have been already reported. Although a number of these putative sequences may be related to biosynthesis or even degradation of non-aromatic compounds, it is highly expected that a significant fraction of them, would be involved in aerobic aromatic hydrocarbon degradation. Metabolic

34

D. Pérez-Pantoja et al.

reconstruction studies linking in vivo with in silico catabolic properties and transcriptional studies would help to address this point. Moreover, most of the current knowledge on biochemistry and genetics of aromatic aerobic microbial metabolism has been obtained with bacterial strains isolated by traditional culture dependent approaches. Taking into account the significant increase in knowledge on strategies to degrade aromatics, which is still obtained by new isolates, it seems obvious that our current knowledge cover only a small proportion of the broad microbial degradative potential.

References Abe T, Masai E, Miyauchi K, Katayama Y, Fukuda M (2005) A tetrahydrofolate-dependent Odemethylase, LigM, is crucial for catabolism of vanillate and syringate in Sphingomonas paucimobilis SYK-6. J Bacteriol 187:2030–2037 Altenschmidt U, Fuchs G (1992) Novel aerobic 2-aminobenzoate metabolism. Purification and characterization of 2-aminobenzoate-CoA ligase, localization of the genes on a 8-kbp plasmid, and cloning and sequencing of the genes from a denitrifying Pseudomonas sp. Eur J Biochem 205:721–727 Altenschmidt U, Oswald B, Steiner E, Herrmann H, Fuchs G (1993) New aerobic benzoate oxidation pathway via benzoyl-Coenzyme A and 3-hydroxybenzoyl-Coenzyme A in a denitrifying Pseudomonas sp. J Bacteriol 175:4851–4858 Arias-Barrau E, Olivera ER, Luengo JM, Fernandez C, Galan B, Garcia JL, Diaz E, Minambres B (2004) The homogentisate pathway: a central catabolic pathway involved in the degradation of L-phenylalanine, L-tyrosine, and 3-hydroxyphenylacetate in Pseudomonas putida. J Bacteriol 186:5062–5077 Arias-Barrau E, Sandoval A, Naharro G, Olivera ER, Luengo JM (2005) A two-component hydroxylase involved in the assimilation of 3-hydroxyphenylacetate in Pseudomonas putida. J Biol Chem 280:26435–26447 Armengaud J, Timmis KN, Wittich RM (1999) A functional 4-hydroxysalicylate/hydroxyquinol degradative pathway gene cluster is linked to the initial dibenzo-p-dioxin pathway genes in Sphingomonas sp strain RW1. J Bacteriol 181:3452–3461 Assinder SJ, Williams PA (1990) The TOL plasmids: determinants of the catabolism of toluene and the xylenes. Adv Microb Physiol 31:1–69 Babbitt PC, Hasson MS, Wedekind JE, Palmer DRJ, Barrett WC, Reed GH, Rayment I, Ringe D, Kenyon GL, Gerlt JA (1996) The enolase superfamily: a general strategy for enzyme-catalyzed abstraction of the alpha-protons of carboxylic acids. Biochemistry 35:16489–16501 Bains J, Boulanger MJ (2007) Biochemical and structural characterization of the paralogous benzoate CoA ligases from Burkholderia xenovorans LB400: defining the entry point into the novel benzoate oxidation (box) pathway. J Mol Biol 373:965–977 Ballou DP, Entsch B, Cole LJ (2005) Dynamics involved in catalysis by single-component and two-component flavin-dependent aromatic hydroxylases. Biochem Biophys Res Commun 338:590–598 Barnes MR, Duetz WA, Williams PA (1997) A 3-(3-hydroxyphenyl)propionic acid catabolic pathway in Rhodococcus globerulus PWD1: cloning and characterization of the hpp operon. J Bacteriol 179:6145–6153 Batie CJ, LaHaie E, Ballou DP (1987) Purification and characterization of phthalate oxygenase and phthalate oxygenase reductase from Pseudomonas cepacia. J Biol Chem 262:1510–1518 Beltrametti F, Marconi AM, Bestetti G, Colombo C, Galli E, Ruzzi M, Zennaro E (1997) Sequencing and functional analysis of styrene catabolism genes from Pseudomonas fluorescens ST. Appl Environ Microbiol 63:2232–2239

Aerobic Degradation of Aromatic Hydrocarbons

35

Bertoni G, Martino M, Galli E, Barbieri P (1998) Analysis of the gene cluster encoding toluene/o-xylene monooxygenase from Pseudomonas stutzeri OX1. Appl Environ Microbiol 64:3626–3632 Bosch R, Moore ERB, GarciaValdes E, Pieper DH (1999) NahW, a novel, inducible salicylate hydroxylase involved in mineralization of naphthalene by Pseudomonas stutzeri AN10. J Bacteriol 181:2315–2322 Bruce NC, Cain RB, Pieper DH, Engesser K-H (1989) Purification and characterization of 4-methylmuconolactone methyl-isomerase, a novel enzyme of the modified 3-oxoadipate pathway in nocardioform actionomycetes. Biochem J 262:303–312 Buder R, Fuchs G (1989) 2-Aminobenzoyl-CoA monooxygenase/reductase, a novel type of flavoenzyme. Purification and some properties of the enzyme. Eur J Biochem 185:629–635 Bundy BM, Campbell AL, Neidle EL (1998) Similarities between the antABC-encoded anthranilate dioxygenase and the benABC-encoded benzoate dioxygenase of Acinetobacter sp. strain ADP1. J Bacteriol 180:4466–4474 Buswell JA, Ribbons DW (1988) Vanillate O-demethylase from Pseudomonas species. Methods Enzymol 161:294–301 Cafaro V, Izzo V, Scognamiglio R, Notomista E, Capasso P, Casbarra A, Pucci P, Di Donato A (2004) Phenol hydroxylase and toluene/o-xylene monooxygenase from Pseudomonas stutzeri OX1: interplay between two enzymes. Appl Environ Microbiol 70:2211–2219 Camara B, Bielecki P, Kaminski F, dos Santos VM, Plumeier I, Nikodem P, Pieper DH (2007) A gene cluster involved in degradation of substituted salicylates via ortho cleavage in Pseudomonas sp. strain MT1 encodes enzymes specifically adapted for transformation of 4-methylcatechol and 3-methylmuconate. J Bacteriol 189:1664–1674 Catelani D, Fiecchi A, Galli E (1971) (+)-γ-Carboxymethyl-γ-methyl-Δα-butenolide. A 1,2 ringfission product of 4-methylcatechol by Pseudomonas desmolyticum. Biochem J 121:89–92 Cha CJ, Cain RB, Bruce NC (1998) The modified beta-ketoadipate pathway in Rhodococcus rhodochrous N75: enzymology of 3-methylmuconolactone metabolism. J Bacteriol 180:6668–6673 Chang HK, Zylstra GJ (1998) Novel organization of the genes for phthalate degradation from Burkholderia cepacia DBO1. J Bacteriol 180:6529–6537 Chang HK, Mohseni P, Zylstra GJ (2003) Characterization and regulation of the genes for a novel anthranilate 1,2-dioxygenase from Burkholderia cepacia DBO1. J Bacteriol 185:5871–5881 Crawford RL (1976) Degradation of homogentisate by strains of Bacillus and Moraxella. Can J Microbiol 22:276–280 Crawford RL (1978) Hydroxylation of 4-hydroxyphenoxyacetate by a Bacillus sp. FEMS Microbiol Lett 4:233–234 Crawford RL, Frick TD (1977) Rapid spectrophotometric differentiation between glutathionedependent and glutathione-independent gentisate and homogentisate pathways. Appl Environ Microbiol 34:170–174 Crawford RL, Hutton SW, Chapman PJ (1975) Purification and properties of gentisate 1,2-dioxygenase from Moraxella osloensis. J Bacteriol 121:794–799 Cronin CN, Kim J, Fuller JH, Zhang X, McIntire WS (1999) Organization and sequences of p-hydroxybenzaldehyde dehydrogenase and other plasmid-encoded genes for early enzymes of the p-cresol degradative pathway in Pseudomonas putida NCIMB 9866 and 9869. DNA Seq 10:7–17 Darby JM, Taylor DG, Hopper DJ (1987) Hydroquinone as a ring-fission substrate in the catabolism of 4-ethylphenol and 4-hydroxyacetophenone by Pseudomonas putida JD1. J Gen Microbiol 133:2137–2146 DeFrank JJ, Ribbons DW (1976) The p-cymene pathway in Pseudomonas putida PL: isolation of a dihydrodiol accumulated by a mutant. Biochem Biophys Res Commun 70:1129–1135 Diaz E, Ferrandez A, Prieto MA, Garcia JL (2001) Biodegradation of aromatic compounds by Escherichia coli. Microbiol Mol Biol Rev 65:523–569 Duarte M, Jauregui R, Vilchez-Vargas R, Junca H, Pieper DH (2014) AromaDeg, a novel database for phylogenomics of aerobic degradation of aromatics. Database 2014; bau118

36

D. Pérez-Pantoja et al.

Dunwell JM, Khuri S, Gane PJ (2000) Microbial relatives of the seed storage proteins of higher plants: conservation of structure and diversification of function during evolution of the cupin superfamily. Microbiol Mol Biol Rev 64:153–179 Eaton RW (1996) p-Cumate catabolic pathway in Pseudomonas putida F1: cloning and characterization of DNA carrying the cmt operon. J Bacteriol 178:1351–1362 Eaton RW (1997) p-Cymene catabolic pathway in Pseudomonas putida F1: cloning and characterization of DNA encoding conversion of p-cymene to p-cumate. J Bacteriol 179:3171–3180 Eaton RW (2001) Plasmid-encoded phthalate catabolic pathway in Arthrobacter keyseri 12B. J Bacteriol 183:3689–3703 Eby DM, Beharry ZM, Coulter ED, Kurtz DM, Neidle EL (2001) Characterization and evolution of anthranilate 1,2-dioxygenase from Acinetobacter sp. strain ADP1. J Bacteriol 183:109–118 Eltis LD, Bolin JT (1996) Evolutionary relationships among extradiol dioxygenases. J Bacteriol 178:5930–5937 Entsch B, van Berkel WJ (1995) Structure and mechanism of para-hydroxybenzoate hydroxylase. FASEB J 9:476–483 Entsch B, Massey V, Claiborne A (1987) p-Hydroxybenzoate hydroxylase containing 6-hydroxyFAD is an effective enzyme with modified reaction mechanisms. J Biol Chem 262:6060–6068 Eulberg D, Golovleva LA, Schlomann M (1997) Characterization of catechol catabolic genes from Rhodococcus erythropolis 1CP. J Bacteriol 179:370–381 Eulberg D, Lakner S, Golovleva LA, Schlomann M (1998) Characterization of a protocatechuate catabolic gene cluster from Rhodococcus opacus 1CP: evidence for a merged enzyme with 4-carboxymuconolactone-decarboxylating and 3-oxoadipate enol-lactone-hydrolyzing activity. J Bacteriol 180:1072–1081 Farrow JM, Pesci EC (2007) Two distinct pathways supply anthranilate as a precursor of the Pseudomonas quinolone signal. J Bacteriol 189:3425–3433 Fernandez C, Ferrandez A, Minambres B, Diaz E, Garcia JL (2006) Genetic characterization of the phenylacetyl-coenzyme A oxygenase from the aerobic phenylacetic acid degradation pathway of Escherichia coli. Appl Environ Microbiol 72:7422–7426 Ferrández A, Garciá JL, Díaz E (1997) Genetic characterization and expression in heterologous hosts of the 3-(3-hydroxyphenyl)propionate catabolic pathway of Escherichia coli K-12. J Bacteriol 179:2573–2581 Ferrandez A, Minambres B, Garcia B, Olivera ER, Luengo JM, Garcia JL, Diaz E (1998) Catabolism of phenylacetic acid in Escherichia coli – Characterization of a new aerobic hybrid pathway. J Biol Chem 273:25974–25986 Ferraroni M, Seifert J, Travkin VM, Thiel M, Kaschabek S, Scozzafava A, Golovleva L, Schlomann M, Briganti F (2005) Crystal structure of the hydroxyquinol 1,2-dioxygenase from Nocardioides simplex 3E, a key enzyme involved in polychlorinated aromatics biodegradation. J Biol Chem 280:21144–21154 Fishman A, Tao Y, Wood TK (2004) Toluene 3-monooxygenase of Ralstonia pickettii PKO1 is a para-hydroxylating enzyme. J Bacteriol 186:3117–3123 Fishman A, Tao Y, Rui L, Wood TK (2005) Controlling the regiospecific oxidation of aromatics via active site engineering of toluene para-monooxygenase of Ralstonia pickettii PKO1. J Biol Chem 280:506–514 Fitzpatrick PF (2003) Mechanism of aromatic amino acid hydroxylation. Biochemistry 42:14083–14091 Fu WJ, Oriel P (1998) Gentisate 1,2-dioxygenase from Haloferax sp. D1227. Extremophiles 2:439–446 Fuenmayor SL, Wild M, Boyes AL, Williams PA (1998) A gene cluster encoding steps in conversion of naphthalene to gentisate in Pseudomonas sp. strain U2. J Bacteriol 180:2522–2530 Fujisawa H, Hayaishi O (1968) Protocatechuate 3,4-dioxygenase. I. Crystallization and characterization. J Biol Chem 243:2673–2681

Aerobic Degradation of Aromatic Hydrocarbons

37

Fukumori F, Saint CP (1997) Nucleotide sequences and regulational analysis of genes involved in conversion of aniline to catechol in Pseudomonas putida UCC22(pTDN1). J Bacteriol 179:399–408 Gasson MJ, Kitamura Y, McLauchlan WR, Narbad A, Parr AJ, Lindsay E, Parsons H, Payne J, Rhodes MJC, Walton NJ (1998) Metabolism of ferulic acid to vanillin – A bacterial gene of the enoyl-SCoA hydratase/isomerase superfamily encodes an enzyme for the hydration and cleavage of a hydroxycinnamic acid SCoA thioester. J Biol Chem 273:4163–4170 Gerlt JA, Babbitt PC (2001) Divergent evolution of enzymatic function: mechanistically diverse superfamilies and functionally distinct suprafamilies. Annu Rev Biochem 70:209–246 Gescher J, Eisenreich W, Worth J, Bacher A, Fuchs G (2005) Aerobic benzoyl-CoA catabolic pathway in Azoarcus evansii: studies on the non-oxygenolytic ring cleavage enzyme. Mol Microbiol 56:1586–1600 Gescher J, Ismail W, Olgeschlager E, Eisenreich W, Worth J, Fuchs G (2006) Aerobic benzoylcoenzyme A (CoA) catabolic pathway in Azoarcus evansii: conversion of ring cleavage product by 3,4-dehydroadipyl-CoA semialdehyde dehydrogenase. J Bacteriol 188:2919–2927 Gibson DT, Parales RE (2000) Aromatic hydrocarbon dioxygenases in environmental biotechnology. Curr Opin Biotechnol 11:236–243 Gibson DT, Koch JR, Kallio RE (1968) Oxidative degradation of aromatic hydrocarbons by microorganisms. 1. Enzymatic formation of catechol from benzene. Biochemistry 7:2653–2662 Gobel M, Kassel-Cati K, Schmidt E, Reineke W (2002) Degradation of aromatics and chloroaromatics by Pseudomonas sp. strain B13: cloning, characterization, and analysis of sequences encoding 3-oxoadipate: succinyl-coenzyme A (CoA) transferase and 3-oxoadipylCoA thiolase. J Bacteriol 184:216–223 Gu W, Song J, Bonner CA, Xie G, Jensen RA (1998) PhhC is an essential aminotransferase for aromatic amino acid catabolism in Pseudomonas aeruginosa. Microbiology 144:3127–3134 Halak S, Basta T, Burger S, Contzen M, Wray V, Pieper DH, Stolz A (2007) 4-Sulfomuconolactone hydrolases from Hydrogenophaga intermedia S1 and Agrobacterium radiobacter S2. J Bacteriol 189:6998–7006 Hara H, Masai E, Katayama Y, Fukuda M (2000) The 4-oxalomesaconate hydratase gene, involved in the protocatechuate 4,5-cleavage pathway, is essential to vanillate and syringate degradation in Sphingomonas paucimobilis SYK-6. J Bacteriol 182:6950–6957 Harayama S, Rekik M (1989) Bacterial aromatic ring-cleavage enzymes are classified into two different gene families. J Biol Chem 264:15328–15333 Harayama S, Leppik RA, Rekik M, Mermod M, Lehrbach PR, Reineke W, Timmis KN (1986) Gene order of the TOL catabolic plasmid upper pathway operon and oxidation of both toluene and benzyl alcohol by the xylA product. J Bacteriol 167:455–461 Harayama S, Mermod N, Rekik M, Lehrbach PR, Timmis KN (1987) Roles of the divergent branches of the meta-cleavage pathway in the degradation of benzoate and substituted benzoates. J Bacteriol 169:558–564 Hareland WA, Crawford RL, Chapman PJ, Dagley S (1975) Metabolic function and properties of 4-hydroxyphenylacetic acid 1-hydroxylase from Pseudomonas acidovorans. J Bacteriol 121:272–285 Harpel MR, Lipscomb JD (1990) Gentisate 1,2-dioxygenase from Pseudomonas. Substrate coordination of the active site Fe2+ and mechanism of turnover. J Biol Chem 265:22187–22196 Harwood CS, Parales RE (1996) The beta-ketoadipate pathway and the biology of self-identity. Annu Rev Microbiol 50:553–590 Hatta T, Mukerjee-Dhar G, Damborsky J, Kiyohara H, Kimbara K (2003) Characterization of a novel thermostable Mn(II)-dependent 2,3-dihydroxybiphenyl 1,2-dioxygenase from a polychlorinated biphenyl- and naphthalene-degrading Bacillus sp. JF8. J Biol Chem 278:21483–21492 He ZG, Spain JC (1998) A novel 2-Aminomuconate deaminase in the nitrobenzene degradation pathway of Pseudomonas pseudoalcaligenes JS45. J Bacteriol 180:2502–2506

38

D. Pérez-Pantoja et al.

He ZQ, Davis JK, Spain JC (1998) Purification, characterization, and sequence analysis of 2-aminomuconic 6-semialdehyde dehydrogenase from Pseudomonas pseudoalcaligenes JS45. J Bacteriol 180:4591–4595 Hintner JP, Lechner C, Riegert U, Kuhm AE, Storm T, Reemtsma T, Stolz A (2001) Direct ring fission of salicylate by a salicylate 1,2- dioxygenase activity from Pseudaminobacter salicylatoxidans. J Bacteriol 183:6936–6942 Hiromoto T, Fujiwara S, Hosokawa K, Yamaguchi H (2006) Crystal structure of 3-hydroxybenzoate hydroxylase from Comamonas testosteroni has a large tunnel for substrate and oxygen access to the active site. J Mol Biol 364:878–896 Hopper DJ (1976) The hydroxylation of p-cresol and its conversion to p-hydroxybenzaldehyde in Pseudomonas putida. Biochem Biophys Res Commun 69:462–468 Hopper D, Taylor DG (1977) The purification and properties of p-cresol-(acceptor) oxidoreductase (hydroxylating), a flavocytochrome from Pseudomonas putida. Biochem J 187:155–162 Hopper D, Chapman PJ, Dagley S (1971) The enzymatic degradation of alkyl-substituted gentisates, maleates and malates. Biochem J 122:29–40 Huang Y, Zhao KX, Shen XH, Chaudhry MT, Jiang CY, Liu SJ (2006) Genetic characterization of the resorcinol catabolic pathway in Corynebacterium glutamicum. Appl Environ Microbiol 72:7238–7245 Hugo N, Armengaud J, Gaillard J, Timmis KN, Jouanneau Y (1998) A novel [2Fe-2S] ferredoxin from Pseudomonas putida mt-2 promotes the reductive reactivation of catechol 2,3-dioxygenase. J Biol Chem 273:9622–9629 Ishiyama D, Vujaklija D, Davies J (2004) Novel pathway of salicylate degradation by Streptomyces sp. strain WA46. Appl Environ Microbiol 70:1297–1306 Ismail W, El-Said Mohamed M, Wanner BL, Datsenko KA, Eisenreich W, Rohdich F, Bacher A, Fuchs G (2003) Functional genomics by NMR spectroscopy. Phenylacetate catabolism in Escherichia coli. Eur J Biochem 270:3047–3054 Iwabuchi T, Harayama S (1998) Biochemical and molecular characterization of 1-hydroxy-2naphthoate dioxygenase from Nocardioides sp. KP7. J Biol Chem 273:8332–8336 Jouanneau Y, Micoud J, Meyer C (2007) Purification and characterization of a three-component salicylate 1-hydroxylase from Sphingomonas sp. strain CHY-1. Appl Environ Microbiol 73:7515–7521 Karlson U, Dwyer DF, Hooper SW, Moore ERB, Timmis KN, Eltis LD (1993) Two independently regulated cytochromes P-450 in a Rhodococcus rhodochrous strain that degrades 2-ethoxyphenol and 4-methoxybenzoate. J Bacteriol 175:1467–1474 Kasai D, Masai E, Miyauchi K, Katayama Y, Fukuda M (2004) Characterization of the 3- O-methylgallate dioxygenase gene and evidence of multiple 3- O-methylgallate catabolic pathways in Sphingomonas paucimobilis SYK-6. J Bacteriol 186:4951–4959 Kasai D, Masai E, Miyauchi K, Katayama Y, Fukuda M (2005) Characterization of the gallate dioxygenase gene: three distinct ring cleavage dioxygenases are involved in syringate degradation by Sphingomonas paucimobilis SYK-6. J Bacteriol 187:5067–5074 Keat MJ, Hopper DJ (1978) p-Cresol and 3,5-xylenol methylhydroxylases in Pseudomonas putida N.C.I.B. 9869. Biochem J 175:649–658 Kirk TK, Farrell RL (1987) Enzymatic “combustion”: the microbial degradation of lignin. Annu Rev Microbiol 41:465–505 Kurnasov O, Jablonski L, Polanuyer B, Dorrestein P, Begley T, Osterman A (2003a) Aerobic tryptophan degradation pathway in bacteria: novel kynurenine formamidase. FEMS Microbiol Lett 227:219–227 Kurnasov O, Goral V, Colabroy K, Gerdes S, Anantha S, Osterman A, Begley TP (2003b) NAD biosynthesis: identification of the tryptophan to quinolinate pathway in bacteria. Chem Biol 10:1195–1204 Lam WWY, Bugg TDH (1997) Purification, characterization, and stereochemical analysis of a C-C hydrolase: 2-hydroxy-6-keto-nona-2,4-diene-1,9-dioic acid 5,6-hydrolase. Biochemistry 36:12242–12251

Aerobic Degradation of Aromatic Hydrocarbons

39

Leahy JG, Batchelor PJ, Morcomb SM (2003) Evolution of the soluble diiron monooxygenases. FEMS Microbiol Rev 27:449–479 Lee J-H, Omori T, Kodama T (1994) Identification of the metabolic intermediates of phthalate by Tn5 mutants of Pseudomonas testosteroni and analysis of the 4,5-dihydroxyphthalate decarboxylase gene. J Ferment Bioeng 77:583–590 Lehrbach PR, Zeyer J, Reineke W, Knackmuss HJ, Timmis KN (1984) Enzyme recruitment in vitro: use of cloned genes to extend the range of haloaromatics degraded by Pseudomonas sp. strain B13. J Bacteriol 158:1025–1032 Liang Q, Takeo M, Chen M, Zhang W, Xu Y, Lin M (2005) Chromosome-encoded gene cluster for the metabolic pathway that converts aniline to TCA-cycle intermediates in Delftia tsuruhatensis AD9. Microbiology 151:3435–3446 Lipscomb JD, Orville AM (1992) Mechanistic aspects of dihydroxybenzoate dioxygenases. In: Sigel H, Sigel A (eds) Metal ions in biological systems. Marcel Dekker Inc., New York, pp 243–298 Liu L, Wu JF, Ma YF, Wang SY, Zhao GP, Liu SJ (2007) A novel deaminase involved in chloronitrobenzene and nitrobenzene degradation with Comamonas sp. strain CNB-1. J Bacteriol 189:2677–2682 Marin M, Perez-Pantoja D, Donoso R, Wray V, Gonzalez B, Pieper DH (2010). Modified 3oxoadipate pathway for the biodegradation of methylaromatics in Pseudomonas reinekei MT1. J Bacteriol 192:1543–1552 Maruyama K (1983) Purification and properties of 2-pyrone-4,6-dicarboxylate hydrolase. J Biochem 93:557–565 Masai E, Katayama Y, Nishikawa S, Fukuda M (1999a) Characterization of Sphingomonas paucimobilis SYK-6 genes involved in degradation of lignin-related compounds. J Ind Microbiol Biotechnol 23:364–373 Masai E, Shinohara S, Hara H, Nishikawa S, Katayama Y, Fukuda M (1999b) Genetic and biochemical characterization of a 2-pyrone-4,6-dicarboxylic acid hydrolase involved in the protocatechuate 4,5-cleavage pathway of Sphingomonas paucimobilis SYK-6. J Bacteriol 181:55–62 Masai E, Sasaki M, Minakawa Y, Abe T, Sonoki T, Miyauchi K, Katayama Y, Fukuda M (2004) A novel tetrahydrofolate-dependent O-demethylase gene is essential for growth of Sphingomonas paucimobilis SYK-6 with syringate. J Bacteriol 186:2757–2765 Matera I, Ferraroni M, Bürger S, Scozzafava A, Stolz A, Briganti F (2008) Salicylate 1,2-dioxygenase from Pseudaminobacter salicylatoxidans: crystal structure of a peculiar ringcleaving dioxygenase. J Mol Biol 380:856–868 McIntire W, Hopper DJ, Singer TP (1985) p-Cresol methylhydroxylase. Biochem J 228:325–335 Mohamed ME (2000) Biochemical and molecular characterization of phenylacetate-coenzyme A ligase, an enzyme catalyzing the first step in aerobic metabolism of phenylacetic acid in Azoarcus evansii. J Bacteriol 182:286–294 Moonen MJ, Kamerbeek N, Westphal AH, Boeren SA, Janssen DB, Fraaije MW, van Berkel WJ (2008a) Elucidation of the 4-hydroxyacetophenone catabolic pathway in Pseudomonas fluorescens ACB. J Bacteriol 190:5190–5198 Moonen MJ, Synowsky SA, van den Berg WA, Westphal AH, Heck AJ, van den Heuvel RH, Fraaije MW, van Berkel WJ (2008b) Hydroquinone dioxygenase from Pseudomonas fluorescens ACB: a novel member of the family of nonheme-iron(II)-dependent dioxygenases. J Bacteriol 190:5199–5209 Moran GR (2005) 4-Hydroxyphenylpyruvate dioxygenase. Arch Biochem Biophys 433:117–128 Morawski B, Segura A, Ornston LN (2000) Substrate range and genetic analysis of Acinetobacter vanillate demethylase. J Bacteriol 182:1383–1389 Murakami S, Sawami Y, Takenaka S, Aoki K (2004) Cloning of a gene encoding 4-amino-3hydroxybenzoate 2,3-dioxygenase from Bordetella sp. 10d. Biochem Biophys Res Commun 314:489–494

40

D. Pérez-Pantoja et al.

Muraki T, Taki M, Hasegawa Y, Iwaki H, Lau PCK (2003) Prokaryotic homologs of the eukaryotic 3-hydroxyanthranilate 3,4-dioxygenase and 2-amino-3-carboxymuconate-6-semialdehyde decarboxylase in the 2-nitrobenzoate degradation pathway of Pseudomonas fluorescens strain KU-7. Appl Environ Microbiol 69:1564–1572 Nakata H, Yamauchi T, Fujisawa H (1979) Phenylalanine hydroxylase from Chromobacterium violaceum Purification and characterization. J Biol Chem 254:1829–1833 Nakazawa T, Hayashi E (1978) Phthalate and 4-hydroxyphthalate metabolism in Pseudomonas testosteroni: purification and properties of 4,5-dihydroxyphthalate decarboxylase. Appl Environ Microbiol 36:264–269 Newman LM, Wackett LP (1995) Purification and characterization of toluene 2-monooxygenase from Burkholderia cepacia G4. Biochemistry 34:14066–14076 Ng LC, Shingler V, Sze CC, Poh CL (1994) Cloning and sequences of the first eight genes of the chromosomally encoded (methyl) phenol degradation pathway from Pseudomonas putida P35X. Gene 151:29–36 Nogales J, Canales A, Jimenez-Barbero J, Garcia JL, Diaz E (2005) Molecular characterization of the gallate dioxygenase from Pseudomonas putida KT2440. The prototype of a new subgroup of extradiol dioxygenases. J Biol Chem 280:35382–35390 Nogales J, Macchi R, Franchi F, Barzaghi D, Fernández C, García JL, Bertoni G, Díaz E (2007) Characterization of the last step of the aerobic phenylacetic acid degradation pathway. Microbiology 153:357–365 Nurk A, Kasak L, Kivisaar M (1991) Sequence of the gene (pheA) encoding phenol monooxygenae from Pseudomonas sp. EST1001: expression in Escherichia coli and Pseudomonas putida. Gene 102:13–18 Olivera ER, Minambres B, Garcia B, Muniz C, Moreno MA, Ferrandez A, Diaz E, Garcia JL, Luengo JM (1998) Molecular characterization of the phenylacetic acid catabolic pathway in Pseudomonas putida U: the phenylacetyl-CoA catabolon. Proc Natl Acad Sci U S A 95:6419–6424 Olsen RH, Kukor JJ, Kaphammer B (1994) A novel toluene-3-monooxygenase pathway cloned from Pseudomonas pickettii PKO1. J Bacteriol 176:3749–3756 Ono K, Nozaki M, Hayaishi O (1970) Purification and some properties of protocatechuate 4,5-dioxygenase. Biochim Biophys Acta 220:224–238 Orii C, Takenaka S, Murakami S, Aoki K (2004) A novel coupled enzyme assay reveals an enzyme responsible for the deamination of a chemically unstable intermediate in the metabolic pathway of 4-amino-3-hydroxybenzoic acid in Bordetella sp. strain 10d. Eur J Biochem 271:3248–3254 Overhage J, Priefert H, Steinbuchel A (1999) Biochemical and genetic analyses of ferulic acid catabolism in Pseudomonas sp strain HR199. Appl Environ Microbiol 65:4837–4847 Peng X, Masai E, Kitayama H, Harada K, Katayama Y, Fukuda M (2002) Characterization of the 5-carboxyvanillate decarboxylase gene and its role in lignin-related biphenyl catabolism in Sphingomonas paucimobilis SYK-6. Appl Environ Microbiol 68:4407–4415 Peng X, Masai E, Kasai D, Miyauchi K, Katayama Y, Fukuda M (2005) A second 5-carboxyvanillate decarboxylase gene, ligW2, is important for lignin-related biphenyl catabolism in Sphingomonas paucimobilis SYK-6. Appl Environ Microbiol 71:5014–5021 Perez-Pantoja D, De la Iglesia R, Pieper DH, Gonzalez B (2008) Metabolic reconstruction of aromatic compounds degradation from the genome of the amazing pollutant degrading bacterium Cupriavidus necator JMP134. FEMS Microbiol Rev 32:736–794 Pieper DH, Engesser K-H, Don RH, Timmis KN, Knackmuss H-J (1985) Modified ortho-cleavage pathway in Alcaligenes eutrophus JMP134 for the degradation of 4-methylcatechol. FEMS Microbiol Lett 29:63–67 Pieper DH, Stadler-Fritzsche K, Knackmuss H-J, Engesser KH, Bruce NC, Cain RB (1990) Purification and characterization of 4-methylmuconolactone methylisomerase, a novel enzyme of the modified 3-oxoadipate pathway in the Gram-negative bacterium Alcaligenes eutrophus JMP 134. Biochem J 271:529–534

Aerobic Degradation of Aromatic Hydrocarbons

41

Pinyakong O, Habe H, Yoshida T, Nojiri H, Omori T (2003) Identification of three novel salicylate 1-hydroxylases involved in the phenanthrene degradation of Sphingobium sp strain P2. Biochem Biophys Res Commun 301:350–357 Plaggenborg R, Steinbuchel A, Priefert H (2001) The coenzyme A-dependent, non-beta-oxidation pathway and not direct deacetylation is the major route for ferulic acid degradation in Delftia acidovorans. FEMS Microbiol Lett 205:9–16 Powell JA, Archer JA (1998) Molecular characterisation of a Rhodococcus ohp operon. Antonie Van Leeuwenhoek 74:175–188 Powlowski J, Shingler V (1994) Genetics and biochemistry of phenol degradation by Pseudomonas sp. CF600. Biodegradation 5:219–236 Powlowski J, Sealy J, Shingler V, Cadieux E (1997) On the role of DmpK, an auxiliary protein associated with multicomponent phenol hydroxylase from Pseudomonas sp. CF600. J Biol Chem 272:945–951 Priefert H, Rabenhorst J, Steinbüchel A (2001) Biotechnological production of vanillin. Appl Microbiol Biotechnol 56:296–314 Prucha M, Peterseim A, Pieper DH (1997) Evidence for an isomeric muconolactone isomerase involved in the metabolism of 4-methylmuconolactone by Alcaligenes eutrophus JMP134. Arch Microbiol 168:33–38 Pujar BG, Ribbons DW (1985) Phthalate metabolism in Pseudomonas fluorescens PHK: purification and properties of 4,5-dihydroxyphthalate decarboxylase. Appl Environ Microbiol 49:374–376 Ranjith NK, Sasikala C, Ramana Ch V (2007) Catabolism of L-phenylalanine and L-tyrosine by Rhodobacter sphaeroides OU5 occurs through 3,4-dihydroxyphenylalanine. Res Microbiol 158:506–511 Reeve D, Carver MA, Hopper DJ (1989) The purification and characterization of 4-ethylphenol methylenehydroxylase, a flavocytochrome from Pseudomonas putida JD1. Biochem J 263:431–437 Reineke W, Knackmuss H-J (1978) Chemical structure and biodegradability of halogenated aromatic compounds. Substituent effects on 1,2-dioxygenation of benzoic acid. Biochim Biophys Acta 532:412–423 Reiner AM (1972) Purification and properties of the catechol-forming enzyme 3,5-cyclohexadiene1,2-diol-1-carboxylic acid (NAD+) oxidoreductase (decarboxylating). J Biol Chem 247:4960–4965 Reiner A, Hegeman G (1971) Metabolism of benzoic acid by bacteria. Accumulation of ()-3,5cyclohexadiene-1,2-diol-1-carboxylic acid by mutant strain of Alcaligenes eutrophus. Biochemistry 10:2530–2536 Roper DI, Cooper RA (1990) Purification, some properties and nucleotide sequence of 5-carboxymethyl-2-hydroxymuconate isomerase of Escherichia coli C. Fed Eur Biochem Soc Lett 266:63–66 Sasoh M, Masai E, Ishibashi S, Hara H, Kamimura N, Miyauchi K, Fukuda M (2006) Characterization of the terephthalate degradation genes of Comamonas sp. strain E6. Appl Environ Microbiol 72:1825–1832 Sauret-Ignazi G, Dardas A, Pelmont J (1988) Purification and properties of cytochrome P-450 from Moraxella sp. Biochimie 70:1385–1395 Schläfli HR, Weiss MA, Leisinger T, Cook AM (1994) Terephthalate 1,2-dioxygenase system from Comamonas testosteroni T-2: purification and some properties of the oxygenase component. J Bacteriol 176:6644–6652 Schlömann M (1994) Evolution of chlorocatechol catabolic pathways. Biodegradation 5:301–321 Schuhle K, Jahn M, Ghisla S, Fuchs G (2001) Two similar gene clusters coding for enzymes of a new type of aerobic 2-aminobenzoate (anthranilate) metabolism in the bacterium Azoarcus evansii. J Bacteriol 183:5268–5278

42

D. Pérez-Pantoja et al.

Shen XH, Jiang CY, Huang Y, Liu ZP, Liu SJ (2005) Functional identification of novel genes involved in the glutathione-independent gentisate pathway in Corynebacterium glutamicum. Appl Environ Microbiol 71:3442–3452 Shields MS, Montgomery SO, Chapman PJ, Cuskey SM, Pritchard PH (1989) Novel pathway of toluene catabolism in the trichloroethylene-degrading bacterium G4. Appl Environ Microbiol 55:1624–1629 Shingler V, Powlowski J, Marklund U (1992) Nucleotide sequence and functional analysis of the complete phenol/3,4-dimethylphenol catabolic pathway of Pseudomonas sp. strain CF600. J Bacteriol 174:711–724 Smith MA, Weaver VB, Young DM, Ornston LN (2003) Genes for chlorogenate and hydroxycinnamate catabolism (hca) are linked to functionally related genes in the dca-pcaqui-pob-hca chromosomal cluster of Acinetobacter sp strain ADP1. Appl Environ Microbiol 69:524–532 Spence EL, Kawamukai M, Sanvoisin J, Braven H, Bugg TD (1996) Catechol dioxygenases from Escherichia coli (MhpB) and Alcaligenes eutrophus (MpcI): sequence analysis and biochemical properties of a third family of extradiol dioxygenases. J Bacteriol 178:5249–5256 Suemori A, Kurane R, Tomizuka N (1993) Purification and properties of gentisate 1,2-dioxygenase from Rhodococcus erythropolis S-1. Biosci Biotechnol Biochem 57:1781–1783 Suemori A, Nakajima K, Kurane R, Nakamura Y (1996) Purification and characterization of o-hydroxyphenylacetate 5-hydroxylase, m-hydroxyphenylacetate 6-hydroxylase and p-hydroxyphenylacetate 1-hydroxylase from Rhodococcus erythropolis. J Ferment Bioeng 81:133–137 Sugimoto K, Senda T, Aoshima H, Masai E, Fukuda M, Mitsui Y (1999) Crystal structure of an aromatic ring opening dioxygenase LigAB, a protocatechuate 4,5-dioxygenase, under aerobic conditions. Structure 7:953–965 Suske WA, Held M, Schmid A, Fleischmann T, Wubbolts MG, Kohler HPE (1997) Purification and characterization of 2-hydroxybiphenyl 3-monooxygenase, a novel NADH-dependent, FAD-containing aromatic hydroxylase from Pseudomonas azelaica HBP1. J Biol Chem 272:24257–24265 Sutherland JB (1986) Demethylation of veratrole by cytochrome P-450 in Streptomyces setonii. Appl Environ Microbiol 52:98–100 Tago K, Sato J, Takesa H, Kawagishi H, Hayatsu M (2005) Characterization of methylhydroquinone-metabolizing oxygenase genes encoded on plasmid in Burkholderia sp. NF100. J Biosci Bioeng 100:517–523 Taguchi K, Motoyama M, Kudo T (2004) Multiplicity of 2,3-dihydroxybiphenyl dioxygenase genes in the Gram-positive polychlorinated biphenyl degrading bacterium Rhodococcus rhodochrous K37. Biosci Biotechnol Biochem 68:787–795 Takenaka S, Murakami S, Shinke R, Hatakeyama K, Yukawa H, Aoki K (1997) Novel genes encoding 2-aminophenol 1,6-dioxygenase from Pseudomonas species AP-3 growing on 2-aminophenol and catalytic properties of the purified enzyme. J Biol Chem 272:14727–14732 Takenaka S, Murakami S, Kim YJ, Aoki K (2000) Complete nucleotide sequence and functional analysis of the genes for 2-aminophenol metabolism from Pseudomonas sp. AP-3. Arch Microbiol 174:265–272 Takenaka S, Asami T, Orii C, Murakami S, Aoki K (2002) A novel meta-cleavage dioxygenase that cleaves a carboxyl-group substituted 2-aminophenol – Purification and characterization of 4-amino-3-hydroxybenzoate 2,3-dioxygenase from Bordetella sp. strain 10d. Eur J Biochem 269:5871–5877 Takenaka S, Okugawa S, Kadowaki M, Murakami S, Aoki K (2003) The metabolic pathway of 4-aminophenol in Burkholderia sp. strain AK-5 differs from that of aniline and aniline with C-4 substituents. Appl Environ Microbiol 69:5410–5413 Tao Y, Fishman A, Bentley WE, Wood TK (2004a) Altering toluene 4-monooxygenase by activesite engineering for the synthesis of 3-methoxycatechol, methoxyhydroquinone, and methylhydroquinone. J Bacteriol 186:4705–4713

Aerobic Degradation of Aromatic Hydrocarbons

43

Tao Y, Fishman A, Bentley WE, Wood TK (2004b) Oxidation of benzene to phenol, catechol, and 1,2,3-trihydroxybenzene by toluene 4-monooxygenase of Pseudomonas mendocina KR1 and toluene 3-monooxygenase of Ralstonia pickettii PKO1. Appl Environ Microbiol 70:3814–3820 Titus GP, Mueller HA, Burgner J, Rodríguez de Córdoba S, Peñalva MA, Timm DE (2000) Crystal structure of human homogentisate dioxygenase. Nat Struct Biol 7:542–546 Vaillancourt FH, Labbe G, Drouin NM, Fortin PD, Eltis LD (2002) The mechanism-based inactivation of 2,3-dihydroxybiphenyl 1,2-dioxygenase by catecholic substrates. J Biol Chem 277:2019–2027 Vaillancourt FH, Bolin JT, Eltis LD (2004) Ring-cleavage dioxygenases. In: Ramos JL (ed) Pseudomonas. Kluwer Academic/Plenum Publishers, New York, pp 359–395 Vaillancourt FH, Bolin JT, Eltis LD (2006) The ins and outs of ring-cleaving dioxygenases. Crit Rev Biochem Mol Biol 41:241–267 van Berkel WJ, Kamerbeek NM, Fraaije MW (2006) Flavoprotein monooxygenases, a diverse class of oxidative biocatalysts. J Biotechnol 124:670–689 Vardar G, Wood TK (2004) Protein engineering of toluene- o-xylene monooxygenase from Pseudomonas stutzeri OX1 for synthesizing 4-methylresorcinol, methylhydroquinone, and pyrogallol. Appl Environ Microbiol 70:3253–3262 Vederas JC, Schleicher E, Tsai MD, Floss HG (1978) Stereochemistry and mechanism of reactions catalyzed by tryptophanase Escherichia coli. J Biol Chem 253:5350–5354 Vetting MW, Wackett LP, Que L, Lipscomb JD, Ohlendorf DH (2004) Crystallographic comparison of manganese- and iron-dependent homoprotocatechuate 2,3-dioxygenases. J Bacteriol 186:1945–1958 Wang L-H, Hamzah RY, Yu Y, Tu S-C (1987) Pseudomonas cepacia 3-hydroxybenzoate 6-hydroxylase: induction, purification, and characterization. Biochemistry 26:1099–1104 Wang YZ, Zhou Y, Zylstra GJ (1995) Molecular analysis of isophthalate and terephthalate degradation by Comamonas testosteroni YZW-D. Environ Health Perspect 103(Suppl 5):9–12 Whited GM, Gibson DT (1991) Toluene-4-monooxygenase, a three-component enzyme system that catalyzes the oxidation of toluene to p-cresol in Pseudomonas mendocina KR1. J Bacteriol 173:3010–3016 Williams SE, Woodridge EM, Ransom SC, Landro JA, Babbitt PC, Kozarich JW (1992) 3-carboxycis, cis-muconate lactonizing enzymes from Pseudomonas putida is homologous to the class 2 fumarase family: a new reaction in the evolution of a mechanistic motif. Biochemistry 31:9768–9776 Wolfe MD, Altier DJ, Stubna A, Popescu CV, Munck E, Lipscomb JD (2002) Benzoate 1,2-dioxygenase from Pseudomonas putida: single turnover kinetics and regulation of a two-component Rieske dioxygenase. Biochemistry 41:9611–9626 Wolgel SA, Dege JE, Perkins-Olson PE, Juarez-Garcia CH, Crawford RL, Münck E, Lipscomb JD (1993) Purification and characterization of protocatechuate 2,3-dioxygenase from Bacillus macerans: a new extradiol catecholic dioxygenase. J Bacteriol 175:4414–4426 Yamaguchi M, Fujisawa H (1980) Purification and characterization of an oxygenase component in benzoate 1,2-dioxygenase system from Pseudomonas arvilla C-1. J Biol Chem 255:5058–5063 Yamamoto S, Katagiri M, Maeno H, Hayaishi O (1965) Salicylate hydroxylase, a monooxygenase requiring flavin adenine dinucleotide. I. Purification and general properties. J Biol Chem 240:3408–3413 Yoshida R, Hori K, Fujiwara M, Saeki Y, Kagamiyama H (1976) Non-identical subunits of protocatechuate 3,4-dioxygenase. Biochemistry 15:4048–4053 Yoshida T, Hayakawa Y, Matsui T, Nagasawa T (2004) Purification and characterization of 2,6-dihydroxybenzoate decarboxylase reversibly catalyzing nonoxidative decarboxylation. Arch Microbiol 181:391–397 Zaar A, Gescher J, Eisenreich W, Bacher A, Fuchs G (2004) New enzymes involved in aerobic benzoate metabolism in Azoarcus evansii. Mol Microbiol 54:223–238

44

D. Pérez-Pantoja et al.

Zhang Y, Colabroy KL, Begley TP, Ealick SE (2005) Structural studies on 3-hydroxyanthranilate3,4-dioxygenase: the catalytic mechanism of a complex oxidation involved in NAD biosynthesis. Biochemistry 44:7632–7643 Zhou NY, Al-Dulayymi J, Baird MS, Williams PA (2002) Salicylate 5-hydroxylase from Ralstonia sp. strain U2: a monooxygenase with close relationships to and shared electron transport proteins with naphthalene dioxygenase. J Bacteriol 184:1547–1555 Zhuang Z, Song F, Takami H, Dunaway-Mariano D (2004) The BH1999 protein of Bacillus halodurans C-125 is gentisyl-coenzyme A thioesterase. J Bacteriol 186:393–399

Aerobic Degradation of Chloroaromatics D. H. Pieper, B. González, B. Cámara, D. Pérez-Pantoja, and W. Reineke

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Metabolism of Chloroaromatics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Peripheral Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Degradation of Chlorocatechols and Chloroprotocatechuates . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Ring Cleavage of (Chloro)Benzoquinols and (Chloro)Hydroxybenzoquinols . . . . . . . 3 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 2 3 15 22 23 24

D.H. Pieper (*) Microbial Interactions and Processes Research Group, HZI – Helmholtz Centre for Infection Research, Braunschweig, Germany e-mail: [email protected] B. González Facultad de Ingeniería y Ciencias, Universidad Adolfo Ibáñez, Santiago, Chile e-mail: [email protected] B. Cámara Laboratorio de Microbiología Molecular y Biotecnología Ambiental, Departamento de Química & Centro de Biotecnología, Universidad Técnica Federico Santa María, Valparaíso, Chile e-mail: [email protected] D. Pérez-Pantoja Departamento de Bioquímica y Biología Molecular, Facultad de Ciencias Biológicas, Universidad de Concepción, Concepción, Chile e-mail: [email protected] W. Reineke Chemical Microbiology, Bergische Universität Wuppertal, Wuppertal, Germany e-mail: [email protected] # Springer International Publishing AG 2016 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-319-39782-5_13-1

1

2

D.H. Pieper et al.

Abstract

Microorganisms are key players in the global carbon cycle. In addition, it appears that most xenobiotic industrial chemicals can be degraded by microorganisms, either by a combination of cometabolic steps or by serving as growth substrate, leading to the mineralization of at least part of the molecule. Here, we present the principles of the microbial aerobic degradation of chloroaromatic compounds. The so-called peripheral sequences of the oxidative degradation of chloroaromatic compounds in aerobic bacteria yield central intermediates with a diphenolic structure such as catechols or hydroxybenzoquinols. These compounds are subsequently cleaved by enzymes that use molecular oxygen and further metabolized by central pathway sequences. The broad variety of mechanisms resulting in dechlorination that occur in these peripheral or central sequences is specifically discussed.

1

Introduction

Chlorinated hydrocarbons comprise a large spectrum of compounds that are or have been of enormous industrial and economic importance. The introduction of chlorine atom(s) into a hydrocarbon significantly influences its physicochemical and biochemical properties and the tendency for bioaccumulation and environmental persistence. Acting in combination with possible (eco)toxicological effects, these properties have pushed the chlorochemistry into the focus of considerable debate and governmental regulatory action. For decades, it is known that microorganisms have the capability to mineralize various chlorinated hydrocarbons. Under anaerobic conditions, chlorinated hydrocarbons can function as alternative electron acceptors in a process termed dehalorespiration. Under aerobic conditions, chlorinated hydrocarbons can function as carbon and energy source, which necessitates dechlorination. This chapter deals with the aerobic degradation and the use of the chloroaromatics as carbon and energy source involving elimination of chloride from the substrate or metabolites.

2

Metabolism of Chloroaromatics

Aerobic microorganisms usually initiate degradation by activation of the aromatic nucleus through oxygenation reactions. A few central intermediates such as catechols, protocatechuate, gentisate, and hydroxybenzoquinols are produced by the introduction of hydroxyl groups, usually in ortho- or para-position to one another (peripheral reactions). These intermediates are subject to oxygenolytic ring cleavage followed by the channeling of the ring-cleavage products into the central metabolism. Despite the fact that various specific dehalogenating enzymes have been identified, microorganisms capable of mineralizing chloroaromatics often employ

Aerobic Degradation of Chloroaromatics

3

peripheral reactions, which have their function in the degradation of naturally occurring aromatics such as benzoate, salicylate, or biphenyl (Reineke 2001). These enzyme systems are predominantly of relaxed-substrate specificity and tolerate lower chlorinated substrate analogs.

2.1

Peripheral Reactions

A broad set of peripheral pathways for chloroaromatic degradation has been described, mainly responsible for transforming chloroaromatics to central catechol or benzoquinol intermediates. Figure 1 gives an overview on reactions leading to (chloro) catechols.

2.1.1 Rieske Non-Heme Iron Oxygenases The so called Rieske non-heme iron oxygenases are one of the key families of enzymes important for aerobic activation and thus degradation of aromatics such as benzoate, benzene, toluene, phthalate, naphthalene, or biphenyl (Gibson and Parales 2000; Duarte et al. 2014) (see Fig. 1, reaction 1). These multicomponent enzyme complexes, composed of a terminal oxygenase component and different electron transport proteins, usually catalyze the incorporation of two oxygen atoms into the aromatic ring to form arene- cis-dihydrodiols, a reaction that is followed by a dehydrogenation catalyzed by cis-dihydrodiol dehydrogenases to give (substituted) catechols. Comparison of the amino acid sequences of the terminal oxygenase α-subunits revealed that they form a family of diverse but evolutionarily related sequences. Although none of the enzymes is completely specific, a broad correlation between the grouping in toluene/biphenyl, naphthalene, benzoate, or phthalate families, and the native substrates oxidized by the family members can be observed (Gibson and Parales 2000). Enzyme engineering studies of biphenyl, benzene, chlorobenzene, and naphthalene dioxygenases showed that the α-subunit of the terminal oxygenase determines substrate specificity and that only slight differences in the amino acid sequence can be associated with dramatic changes in substrate specificity or regioselectivity (Beil et al. 1998; Furukawa et al. 2004). However, as a general rule Rieske non-heme iron oxygenases can transform lower chlorinated substrate analogs and dioxygenation is usually directed to carbon atoms proximal to substituents. 2.1.2

Oxygenolytic Dehalogenations by Rieske Non-Heme Iron Oxygenases Different bacteria capable of degrading 2-chlorobenzoate have been described. All these organisms catalyze a 1,2-dioxygenation such that one of the vic-hydroxyl groups in the cis-dihydrodiol is bound to the same carbon as the chloro-substituent. From such an unstable vic-dihydrodiol, the chloro-substituent is spontaneously eliminated to form catechols (Fig. 2). Two distinct 2-chlorobenzoate-degrading dioxygenase systems have been described. The two-component 2-halobenzoate 1,2-dioxygenases (oxygenase consisting of α- and β-subunits and a reductase as the one from strain Burkholderia

Fig. 1 Overview of peripheral reactions in the degradation of chloroaromatics channeling to catechols. (1) Rieske non-heme iron oxygenases (1a) and dehydrogenases (1b) involved in (chloro)benzene and (chloro)benzoate metabolism; (2) aromatic monooxygenases (soluble diiron monooxygenases); (3) phenol hydroxylases (soluble diiron monooxygenases or single component flavin monooxygenases); (4) salicylate 1-hydroxylases (single component flavin monooxygenases); (5) α-ketoglutarate-dependent dioxygenases; (6) 2,4,5-trichloro- and 2,4-dichlorophenoxyacetate monooxygenases; (7) oxidation of side chains (xylene monooxygenases, 7a; benzylalcohol dehydrogenases, 7b; benzaldehyde dehydrogenases, 7c)

4 D.H. Pieper et al.

Aerobic Degradation of Chloroaromatics

5

Fig. 2 Oxygenolytic dehalogenation of 2-chlorobenzoate by Rieske non-heme iron oxygenases

cepacia 2CBS (Fetzner et al. 1992)), are similar to two-component toluate and benzoate 1,2-dioxygenases of the Rieske non-heme iron oxygenases. They are characterized by their high activity against 2-halosubstituted benzoates but have negligible activity with 4-chloro-, or 2,5-dichlorobenzoate. In contrast, the broad specificity 2-chlorobenzoate dioxygenase of P. aeruginosa strain 142 is a threecomponent dioxygenase system (oxygenase consisting of α- and β-subunits, ferredoxin and reductase) (Romanov and Hausinger 1994), the α-subunit of which exhibits only 22% sequence identity with that of strain 2CBS, but a significant similarity (42%) to salicylate 5-hydroxylase NagG from Pseudomonas sp. strain U2. Thus, 2-chlorobenzoate dioxygenases are functionally similar, but represent two different lineages with distinct activities. Oxidative dehalogenation is not restricted to 2-halobenzoate 1,2-dioxygenases but has also been described for tetrachlorobenzene dioxygenase TecA of Ralstonia sp. PS12 (Beil et al. 1998), biphenyl 2,3-dioxygenase BphA of Burkholderia xenovorans LB400 (Seeger et al. 1995), or 3-chlorobenzoate 4,5-dioxygenase CbaA from Comamonas testosteroni BR60 (Nakatsu and Wyndham 1993). Dehalogenation always involves dioxygenolytic attack on a chlorosubstituted carbon atom and its unsubstituted neighbor to give unstable intermediates, which spontaneously rearrange with elimination of chloride. However, in all cases mentioned here, only higher chlorinated substrate analogs are dechlorinated. CbaA catalyzes the 4,5-dioxygenation of 3,4-dichlorobenzoate resulting in an unstable dihydrodiol, which spontaneously eliminates chloride to form 5-chloroprotocatechuate, whereas 4,5-dioxygenation of 3-chlorobenzoate yields 5-chloroprotocatechuate after dehydrogenation. TecA catalyzes the dehalogenation of 1,2,4,5-tetrachlorobenzene to form 3,4,6-trichlorocatechol, whereas lower chlorinated benzenes were transformed to the corresponding dihydrodiols. BphA catalyzes the dehalogenation of 2,20 - or 2,40 -dichlorobiphenyl among others. For both TecA and BphA, amino acid residues crucial for dehalogenation were identified (Beil et al. 1998; Furukawa et al. 2004).

2.1.3 Soluble Diiron Monooxygenases Enzymes attacking the nonactivated benzene nucleus by monooxygenation belong to an evolutionary-related family of soluble diiron monooxygenases (Leahy et al. 2003) that are enzyme complexes consisting of an electron transport system comprising a reductase (and in some cases a ferredoxin), a catalytic effector protein, and a terminal hydroxylase with a (αβγ)2 quaternary structure and a diiron center

6

D.H. Pieper et al.

contained in each α-subunit. The monooxygenases are classified according to their α-subunits, which are assumed to be the site of substrate hydroxylation, into four different phylogenetic groups, comprising the (multicomponent) phenol hydroxylases, and the four-component alkene/aromatic monooxygenases (Leahy et al. 2003). The multicomponent phenol hydroxylases (Fig. 1, reaction 3) such as the phenol hydroxylase of Pseudomonas sp. CF600 share the capability to hydroxylate phenol and methylsubstituted derivatives (Shingler et al. 1992), and a few enzymes of this group, such as phenol hydroxylase of P. stutzeri OX1, also have been shown to hydroxylate the nonactivated benzene nucleus and thus can catalyze two sequential hydroxylations (Cafaro et al. 2004). The available information on the transformation of chlorophenols by multicomponent phenol hydroxylases is insufficient; however, preliminary data indicate that such a capability is spread among this group of enzymes (Teramoto et al. 1999). The four-component alkene/aromatic monooxygenases comprise enzymes that oxidize nonhydroxylated compounds (Leahy et al. 2003) (Fig. 1, reaction 2). Toluene 4-monooxygenase of P. mendocina KR1 also transforms chlorobenzene with 4-chlorophenol as product (Fishman et al. 2005). Regioselectivity of attack is mainly controlled by the α-subunit of the enzyme, but also by the effector protein (Mitchell et al. 2002), in such a way that the enzyme could be modulated also to produce 2-chloro- and 3-chlorophenol. Tbc2 toluene 2-monooxygenase of strain Burkholderia cepacia JS150 was shown to transform chlorobenzene into 2-chlorophenol (Kahng et al. 2001).

2.1.4 Single Component Flavoprotein Monooxygenases The oxidation of various phenolic aromatic compounds is catalyzed by singlecomponent flavoprotein monooxygenases (class A flavin monooxygenases (van Berkel et al. 2006)). These enzymes catalyze hydroxylation in ortho- or para-position to the preexisting hydroxyl-group and usually display a narrow substrate specificity (van Berkel et al. 2006). Only a few of these enzymes have been analyzed for their capability to transform chlorinated substrate analogues. Single-component salicylate 1-hydroxylases (Fig. 3) have been reported to catalyze the transformation of salicylate to catechol (Fig. 1, reaction 4), and enzymes analyzed in this aspect were capable of transforming chlorinated salicylates into the respective chlorocatechols with high activity (Bosch et al. 1999; Cámara et al. 2007). In accordance with these properties, mineralization of chlorosalicylates by natural or engineered organisms involving salicylate 1-hydroxylase has been reported (Cámara et al. 2007; Lehrbach et al. 1984). Another group of single component flavin monooxygenases, termed phenol hydroxylases (Fig. 1, reaction 3), has been described from phenol degrading Pseudomonas strains, among them PheA from Pseudomonas sp. strain EST1001, which transforms phenol and 3-methylphenol (Nurk et al. 1991). This substrate specificity significantly differs from that described for a group of enzymes termed 2,4dichlorophenol hydroxylases and involved in the degradation of 2,4-D or

Aerobic Degradation of Chloroaromatics

7

Fig. 3 Dendrogram showing the relatedness of single component flavin monooxygenases (2,4-dichlorophenol hydroxylase, TfdB, ClpB; phenol hydroxylase, PheA; salicylate 1-hydroxylase, SalA, NahG, NahW; pentachlorophenol monooxygenase, PcpB; 2-hydroxybiphenyl 3-monooxygenase, HbpA; chlorobenzoquinol monooxygenase, ChqA; unknown function, CphC-II)

2,4-dichlorophenol via 3,5-dichlorocatechol, with which phenol hydroxylases share 50% sequence identity (Fig. 3). In contrast to the substrate specificity of phenol hydroxylases, all the five characterized dichlorophenol hydroxylases, TfdB of Burkholderia cepacia 2a (Beadle and Smith 1982), ClpB from Defluvibacter lusatiensis S1 (Makdessi and Lechner 1997), TfdBa from Bradyrhizobium sp. RD5-C2 (Huong et al. 2007), and both TfdBI and TfdBII dichlorophenol hydroxylases from C. necator JMP134 (Ledger et al. 2006), are highly active with 2,4-dichloro-, 4-chloro-2-methylphenol and 4-chlorophenol, but activity with phenol and 3-chlorophenol was either low or absent. Despite the obvious similarity in substrate specificity, these dichlorophenol hydroxylases are just distantly related. Various 2,4-D degraders contain dichlorophenol hydroxylases sequences highly related to that of B. cepacia 2a or

8

D.H. Pieper et al.

TfdBII from strain JMP134 (e.g., A. denitrificans EST4002, D. acidovorans P4a, Burkholderia sp. RASC or V. paradoxus TV1). In contrast, the closest homolog of TfdBI from strain JMP134 is the enzyme of D. lusatiensis (only 76% identity). However, partial amino acid sequences indicate that enzymes highly similar to TfdBI are present in other 2,4-D degrading β-proteobacterial strains (Vallaeys et al. 1999). In addition to ClpB from D. lusatiensis and TfdBa from Bradyrhizobium sp., a third lineage of dichlorophenol hydroxylases represented by TfdB of S. herbicidovorans MH (Müller et al. 2004) has been observed in α-proteobacteria. All these three lineages have only approximately 60% amino acid identity (Fig. 3) and similar identity to TfdB proteins from β-proteobacteria, indicating an ancient divergence and evolution without recent horizontal gene transfer in α-proteobacteria.

2.1.5

Initial Side-Chain Processing

Oxidation of Methylgroups Aromatic compounds that bear alkyl substituents on the aromatic ring may undergo oxidation of the side chain before ring activation. The most intensely described example is the TOL plasmid encoded degradation pathway of toluene and xylenes by Pseudomonas putida mt-2, which proceeds via benzoate or the respective methylbenzoates, and catechol or methylcatechols as intermediates (Worsey and Williams 1975) (Fig. 1, reactions 7). The respective enzymes are encoded by the xyl genes, which are organized in two functional units localized on the TOL plasmid pWW0 (Greated et al. 2002). The upper pathway encodes three enzymes, xylene monooxygenase, benzylalcohol dehydrogenase, and benzaldehyde dehydrogenase, that oxidize toluene and xylenes to benzoate and toluates, respectively. The degree of transformation of chlorotoluenes by xylene monooxygenase depends on the position of the chlorine substituent. The substrate analogs 3-chloro- and 4-chlorotoluene are transformed at high rates, while no or only low activity has been found with other chlorotoluenes. Substituents in ortho position impaired substrate binding (Brinkmann and Reineke 1992). Benzylalcohol and benzaldehyde dehydrogenases show broader specificities. But again, a chlorine substituent in ortho position leads to a drastic decrease in the substrate conversion. Transfer of the TOL plasmid into strains capable of mineralizing chlorocatechols (see below) allowed the isolation of 3-chloro- and 4-chlorotoluene-degrading organisms (Brinkmann and Reineke 1992). α-Ketoglutarate-Dependent Dioxygenases Bacterial metabolism of 2,4-D and MCPA is initiated by the cleavage of the side chain, resulting in the formation of 2,4-dichloro- and 4-chloro-2-methylphenol, respectively (Fig. 1, reaction 5). In C. necator JMP134 (pJP4), the degradation of 2,4-D is catalyzed by 2,4-dichlorophenoxyacetate/α-ketoglutarate dioxygenase, TfdA (Fukumori and Hausinger 1993). TfdA belongs to the large superfamily of α-ketoacid-dependent, mononuclear non-heme iron oxygen activating enzymes that catalyze the oxidation of aliphatic C–H bonds (Schofield and Zhang 1999). During

Aerobic Degradation of Chloroaromatics

9

Fig. 4 Dendrogram showing the relatedness of 2,4-D/α-ketoglutarate dioxygenases (TfdA), (S)mecoprop/and (R)-mecoprop/α-ketoglutarate dioxygenases (SdpA and RdpA, respectively). TauD taurine α-ketoglutarate dioxygenase from E. coli is shown as outgroup

2,4-D transformation, one oxygen atom is involved in the oxidative decarboxylation of α-ketoglutarate, whereas the other oxygen atom is introduced into the β-carbon of the aliphatic side chain to produce an unstable hemiacetal intermediate that decomposes to glyoxylate, and 2,4-DCP (Fukumori and Hausinger 1993). TfdA is able to hydroxylate several phenoxyacetates, including 2,4,5-trichlorophenoxyacetate (2,4,5-T) and MCPA; however, 2,4-D is the preferred substrate. Representatives of tfdA genes have been described thus far from α, β, and γ subgroups of the proteobacteria (McGowan et al. 1998). Sequence alignment among β-proteobacterial representatives reveals the presence of three distinct classes of tfdA gene sequences with slight variations in each class (Fig. 4); however, all share >80% sequence identity and all are capable of transforming several phenoxyacetate compounds. tfdA-like genes were also observed in 2,4-D degrading α-proteobacteria belonging to the Bradyrhizobium- Agromonas- Nitrobacter- Afipia cluster (Itoh et al. 2002), and these tfdAα sequences show 56–60% identity to the β-proteobacterial counterparts. tfdA-like genes were also detected in 2,4-D degrading sphingomonads showing 46–51% of nucleotide sequence identity with either tfdA or tfdAα-genes (Itoh et al. 2004) (Fig. 4).

10

D.H. Pieper et al.

Fig. 5 Degradation of 4-chlorobenzoate involving hydrolytic dehalogenation

Dichlorprop ((RS)-2-(2,4-dichlorophenoxy)propionate) and mecoprop ((RS)-2(4-chloro-2-methylphenoxy)propionate) are chiral molecules that are poor substrates for the TfdA enzymes already mentioned. Experiments with pure enantiomers of mecoprop revealed that S. herbicidovorans MH and D. acidovorans MC1 can degrade both enantiomers. In both organisms, enantioselective α-ketoglutaratedependent dioxygenases termed SdpA and RdpA were observed (Müller et al. 2006; Westendorf et al. 2003) with the SdpA proteins (64% sequence identity) being highly specific in transforming only the (S)-enantiomers of mecoprop and dichlorprop and the (identical) RdpA proteins active only against the (R)-enantiomers. Like TfdA, SdpA and RdpA are α-ketoglutarate-dependent dioxygenases; however SdpA and RdpA share only 25% of protein sequence identity and only less than 35% of identity with previously characterized TfdA proteins (Fig. 4).

2.1.6 Hydrolytic Dehalogenation of 4-Chlorobenzoate Various Arthrobacter, Pseudomonas and Alcaligenes strains degrading 4-chlorobenzoate (4CB) by a pathway involving an initial dehalogenation event have been described. The 4-chlorobenzoate dehalogenase system consists of three distinct enzymes, a 4CB-coenzyme A (CoA) ligase, 4CB-CoA dehalogenase, and 4-hydroxybenzoate (4HB)-CoA thioesterase (Fig. 5) encoded by the fcbA, fcbB and fcbC genes, respectively. In 4CB degrading actinobacteria, a fcbABC gene order is conserved (Schmitz et al. 1992), whereas 4CB degrading proteobacterial strains exhibit a fcbBAC gene order (Savard et al. 1992). In some cases, the gene cluster also encodes putative transport proteins between the fcbB and fcbA genes. In addition to differences in the gene order, significant differences in protein sequence were observed. Actinobacterial 4CB-CoA ligases are highly similar (>95% sequence identity), but differ significantly (38–44% identity) from proteobacterial 4CB-CoA ligases; however, all ligases share significant sequence similarity with proteins catalyzing similar chemistry in the β-oxidation pathway (Babbitt et al. 1992). The following dehalogenation by 4CB-CoA dehalogenase forms 4-hydroxybenzoyl-CoA in a hydrolytic substitution reaction (Liang et al. 1993). This group of enzymes utilizes a unique form of catalysis in which an active site carboxylate bonds to C-4 of the benzoyl ring of the bound substrate to form a Meisenheimer-like complex. Expulsion of the chloride from the Meisenheimer complex with concomitant

Aerobic Degradation of Chloroaromatics

11

rearomatization of the benzoyl ring generates an arylated enzyme as the second reaction intermediate (Dong et al. 2002). Hydrolysis of the arylated enzyme occurs by addition of a water molecule to the acyl carbonyl carbon to form a tetrahedral intermediate, which expels the hydroxylbenzoyl group to generate the catalytic carboxylate residue and form 4-hydroxybenzoyl-CoA. As observed for the ligases, the actinobacterial dehalogenases are highly similar (>98% sequence identity) but differ significantly (approximately 50% sequence identity) from proteobacterial dehalogenases. The last reaction step to form 4HB is carried out by the 4HB-CoA thioesterase. The absence of serine, cysteine, or histidine catalytic residues in the Pseudomonas sp. strain CBS-3 enzyme had distinguished this protein from previously characterized thioesterases. However, crystallographic investigation (Benning et al. 1998) revealed a high similarity of its three-dimensional structure to that of β-hydroxydecanoyl thiol ester dehydrase from E. coli. Both enzymes contained the so-called Hotdog fold, which now was found to be shared by numerous thioesterases and dehydrase proteins (Dillon and Bateman 2004). An active site aspartate was suggested to participate in nucleophilic catalysis (Zhuang et al. 2002). Other proteobacterial 4HB-CoA thioesterases belong to the same subfamily of Hotdog fold proteins, whereas the respective thioesterases from Arthrobacter strains do not share significant sequence similarity with the proteobacterial counterparts. However, they also belong to the same superfamily and an active site glutamate was suggested as catalytic nucleophile (Dillon and Bateman 2004).

2.1.7

Reactions Leading to (Chloro)Benzoquinols and (Chloro) Hydroxybenzoquinols The degradation of various, specially higher chlorinated aromatic pollutants, proceeds via (chloro)benzoquinols or (chloro)hydroxybenzoquinols as central intermediates (Fig. 6). Degradation of Lindane One of the most important xenobiotic compounds metabolized through (chloro) benzoquinol is lindane or γ-hexachlorocyclohexane (γ-HCH), the biodegradation of which has been mainly studied in Sphingobium japonicum UT26 (Endo et al. 2005) (Fig. 6). However, other γ-HCH degrading strains have recently been found to possess nearly identical genes for degradation (Nagata et al. 2007). LinA γ-HCH dehydrochlorinase catalyzes two steps of dehydrochlorination to 1,3,4,6-tetrachloro1,4-cyclohexadiene (1,4-TCDN) via γ-pentachlorocyclohexene (Nagata et al. 2007). 1,4-TCDN is further transformed by LinB 1,4-TCDN chlorohydrolase: a reaction competing with spontaneous rearrangement to the dead-end 1,2,4-trichlorobenzene product. Unlike LinA, which seems to be a unique dehydrochlorinase, LinB belongs to the haloalkane dehalogenase family and catalyzes two successive hydrolytic dehalogenations to form 2,5-dichloro-2,5-cyclohexadiene-1,4-diol via 2,4,5-trichloro-2,5-cyclohexadiene-1-ol (Nagata et al. 2007). Further metabolism of 2,5-dichloro-2,5-cyclohexadiene-1,4-diol is achieved by LinC or LinX 2,5-dichloro-2,5-cyclohexadiene-1,4-diol dehydrogenase, members of the shortchain alcohol dehydrogenase superfamily (Nagata et al. 1994), forming

Fig. 6 Catabolic pathways leading to (chloro)benzoquinols and (chloro)hydroxybenzoquinols as central intermediates. The metabolism of γ-hexachlorocyclohexane by S. japonicum UT26, of pentachlorophenol by S. chlorophenolicum ATCC39723, of 2,4,6-trichlorophenol by C. necator JMP134 and of 2,4,5-trichlorophenol by B. phenoliruptrix AC1100 are shown. Unstable intermediates are enclosed in brackets. Broken arrows indicate reactions of the lower pathway detailed in Fig. 10

12 D.H. Pieper et al.

Aerobic Degradation of Chloroaromatics

13

Fig. 7 Diversity of pathways for the degradation/transformation of chlorocatechols. C12O catechol 1,2-dioxygenase, CC12O chlorocatechol 1,2-dioxygenase, MCI muconate cycloisomerase, MCIP proteobacterial muconate cycloisomerase, CMCIP proteobacterial chloromuconate cycloisomerase, CMCIA actinobacteral chloromuconate cycloisomerase, MLI muconolactone isomerase, CMLIA actinobacterial chloromuconolactone isomerase, DLHP proteobacterial dienelactone hydrolase, DLHA actinobacterial dienelactone hydrolase, tDLH trans-dienelactone hydrolase, MAR maleylacetate reductase. The unstable intermediate is enclosed in brackets

2,5-dichloro- p-benzoquinol, which is subject to a reductive dechlorination by 2,5dichloro- p-benzoquinol dechlorinase (LinD). LinD belongs to the glutathione transferases family, and catalyzes a rapid dechlorination to chlorobenzoquinol and a subsequent slow conversion to further dechlorinate and forms benzoquinol (Miyauchi et al. 1998). Transformation of Chlorinated Phenols and Phenoxyacetates to (Chloro) Benzoquinols and (Chloro)Hydroxybenzoquinols Like HCH, pentachlorophenol (PCP) is degraded through (chloro)benzoquinol and the pathway has been elucidated in detail in Sphingobium chlorophenolicum ATCC

14

D.H. Pieper et al.

39723 (Crawford et al. 2007) (Fig. 6). The oxidation of PCP is catalyzed by PCP 4-monooxygenase (PcpB), a flavin monooxygenase with functional domains similar to those of other bacterial flavoprotein monooxygenases specific to phenolic substrates (see Fig. 3). PcpB can catalyze attack on a broad range of substituted phenols, hydroxylating the para position and removing halogen, nitro, amino, and cyano groups (Xun et al. 1992a). The generated tetrachloro- p-benzoquinone is reduced to tetrachloro- p-benzoquinol by a NADPH-dependent tetrachlorobenzoquinone reductase (PcpD) (Dai et al. 2003). The following TeCH reductive dehalogenase (PcpC)catalyzed reactions, reductive dehalogenation to 2,3,6-trichloro- p-benzoquinol and then to 2,6-dichloro- p-benzoquinol (DiCH) (Xun et al. 1992b) resemble those catalyzed by LinD of the HCH degradation pathway. However, in contrast, two chlorine atoms remain on the aromatic ring before ring-cleavage. The broadly used herbicide 2,4,5-T is also metabolized through the (chloro) hydroxybenzoquinol pathway, which has been well studied in Burkholderia phenoliruptrix AC1100 (Hübner et al. 1998). The transformation to 2,4,5trichlorophenol (2,4,5-TCP) is performed by a two-component monooxygenase (TftAB) (Danganan et al. 1995), which exhibits strong homology to terminal oxygenase subunits of benzoate and toluate dioxygenases and thus contrasts the degradation of 2,4-D, which is typically catalyzed by 2,4-D/α-ketoglutarate dioxygenases (see section “α-Ketoglutarate-Dependent Dioxygenases”). However, 2,4-D monooxygenase genes (cadAB) with significant identity (approximately 45% on the amino acid level) with TftAB of B. phenoliruptrix AC1100 were observed in Bradyrhizobium sp. strain HW13 capable of degrading 2,4-D (Kitagawa et al. 2002). The substrate profiles of the 2,4,5-T oxygenase and 2,4-D oxygenase are similar (Danganan et al. 1995; Kitagawa et al. 2002) with high activity against 2,4-D and 2,4,5-T. Additional experiments indicated the presence of respective genes in various 2,4-D degrading Sphingomonas (B6–10, TFD26, TFD44) and Bradyrhizobium (HWK12, BTH) strains (Kitagawa et al. 2002). Further metabolism of 2,4,5-TCP (Fig. 6) is achieved by TftCD 2,4,5-TCP monooxygenase, a class D monooxygenase (van Berkel et al. 2006) comprising a FADH2-utilizing monooxygenase (TfdD) and a NADH:FAD oxidoreductase (TfdC). TfdC supplies FADH2 to TftD (Gisi and Xun 2003) converting 2,4,5-TCP via 2,5-dichlorobenzoquinol to 5-chloro-2-hydroxybenzoquinol. 5-Chloro-2hydroxybenzoquinol is dechlorinated to hydroxybenzoquinone by a dechlorinase (TftG) and then reduced to hydroxybenzoquinol by a quinone reductase the gene of which is presently unknown. A similar pathway has been reported for the degradation of 2,4,6-trichlorophenol (2,4,6-TCP) in C. necator JMP134 (Fig. 6). In this bacterium, 2,4,6-TCP monooxygenase (TcpA with 65% sequence identity to TftD) catalyzes the oxidative conversion of 2,4,6-TCP to 2,6-dichlorobenzoquinone, followed by hydrolytic dechlorination to produce 6-chloro-2-hydroxybenzoquinone (Xun and Webster 2004). Like TftC, TcpX NADH:FAD oxidoreductase provides FADH2 for TcpA catalysis (Matus et al. 2003). 6-Chloro-2-hydroxybenzoquinone is reduced to 6-chlorohydroxybenzoquinol by a quinone reductase (TcpB) (Belchik and Xun 2008). Recently, evidence was accumulated that also 2,4-dichlorophenol (Ferraroni

Aerobic Degradation of Chloroaromatics

15

Fig. 8 Dendrogram showing the relatedness of muconate cycloisomerases (CatB) and chloromuconate cycloisomerases (ClcB, TfdD, TcbD, TetD). Mandelate racemase MdlA of P. putida is shown as outgroup

et al. 2005) and 4-chlorophenol (Nordin et al. 2005) can be degraded via (chloro) hydroxybenzoquinols as intermediate. The downstream catabolism of (chloro) hydroxybenzoquinols is described in Sect. 2.3.

2.2

Degradation of Chlorocatechols and Chloroprotocatechuates

2.2.1 Metabolism of Chlorocatechols via the 3-Oxoadipate Pathway The chromosomally encoded 3-oxoadipate pathway is widely distributed in soil bacteria, specifically proteobacteria and actinobacteria. The function of the catechol branch of this pathway is to transform catechol to Krebs cycle intermediates (Fig. 7). From phylogenetic analyses, proteobacterial catechol 1,2-dioxygenases (C12O) and actinobacterial C12Os form separate branches with low sequence identity to one another; however, all those enzymes seem to be characterized by similar kinetic properties and out of the chlorocatechols, only 4-chlorocatechol is transformed at a reasonable rate (Dorn and Knackmuss 1978; Matsumura et al. 2004).

16

D.H. Pieper et al.

Also proteobacterial and actinobacterial muconate cycloisomerases (MCI) constitute separate evolutionary branches with low sequence identity to one another (Fig. 8). Proteobacterial MCIs catalyze the cycloisomerization of 3-chloromuconate at reasonable rates (Cámara et al. 2007). This reaction is assumed to proceed via 4-chloromuconolactone as intermediate (Fig. 7), which either spontaneously or enzyme-catalyzed rearranges with concomitant dehalogenation and decarboxylation to form protoanemonin (Blasco et al. 1995; Nikodem et al. 2003), a compound of high toxicity. The cycloisomerization product of 3-chloromuconate by actinobacterial MCIs has not yet been identified. In the case of 2-chloromuconate turnover, proteobacterial MCIs catalyze cycloisomerization only, to form both 2-chloro- and 5-chloromuconolactone as stable products (Vollmer et al. 1994) (Fig. 7). Even though only few studies are available on actinobacterial MCIs, some studies suggested that they differ in catalytic properties from their proteobacterial counterparts and form exclusively 5-chloromuconolactone (Solyanikova et al. 1995). Both 5-chloro- and 2-chloromuconolactone are substrates of muconolactone isomerases of the 3-oxoadipate pathway. 5-Chloromuconolactone is transformed predominantly to cis-dienelactone probably via abstraction of the C4 proton followed by spontaneous chloride elimination (Prucha et al. 1996) and 2-chloromuconolactone into protoanemonin, probably by the elimination of CO2 and chloride from chlorosubstituted 3-oxoadipate enol-lactone (Skiba et al. 2002) (Fig. 7). Mineralization of chlorocatechols via the 3-oxoadipate pathway is thus prevented by (1) restricted substrate specificity of the enzymes, (2) formation of protoanemonin as toxic dead-end product, or (3) formation of cis-dienelactone as dead-end product.

2.2.2 The Chlorocatechol Pathway In most organisms isolated thus far, chlorocatechols are degraded by a chlorocatechol ortho-cleavage pathway (Reineke 2001). This pathway is initiated by a broad specificity chlorocatechol 1,2-dioxygenase with consumption of molecular oxygen to produce the corresponding chloro-cis,cis-muconates (Dorn and Knackmuss 1978). This reaction is identical to the one of the 3-oxoadipate pathway, and the main difference between the enzymes is their substrate specificity. A key enzyme in the pathway is chloromuconate cycloisomerase (CMCI), which differs in various aspects from muconate cycloisomerase. Proteobacterial CMCIs catalyze a specific cycloisomerization of 2-chloro-cis,cis-muconate (the metabolite of 3-chlorocatechol degradation) to form preferentially 5-chloromuconolactone, which is dehalogenated to trans-dienelactone (Vollmer and Schlömann 1995). Thus, proteobacterial CMCIs are specific dehalogenases. The formation of transdienelactone is due to the fact that, after cycloisomerization, the lactone ring has to rotate in the catalytic center to achieve proton abstraction and thus dehalogenation (Schell et al. 1999) (Fig. 7). In contrast, chloride from the 3-position of 3-chloro-cis, cis-muconate (the metabolite of 4-chlorocatechol degradation) seems to be directly eliminated during cycloisomerization to form cis-dienelactone (Kaulmann et al. 2001) (Fig. 7). Similarly, 2,4-dichloro-cis,cis-muconate, the metabolite of

Aerobic Degradation of Chloroaromatics

17

3,5-dichlorocatechol degradation, is converted to 2-chloro-cis-dienelactone (Pieper et al. 1991). The dienelactones are converted into the respective maleylacetates by dienelactone hydrolases (DLH) (Schmidt and Knackmuss 1980). Evidently, for the degradation of various chloroaromatics, an important prerequisite is that the dienelactone hydrolase is of relaxed substrate specificity and accepts both cis- and trans-isomers as substrates. The following enzyme, maleylacetate reductase, plays a major role in the degradation of chloroaromatic compounds either in the ortho-cleavage chlorocatechol pathway (Kaschabek and Reineke 1992) or as part of the benzoquinol pathway. The original function is the reduction of the double bond to channel maleylacetate into the 3-oxoadipate pathway. Maleylacetates with chlorine substituents in the 2-position are subject to reductive dechlorination, e.g., 2-chloromaleylacetate, the intermediate in 3,5-dichlorocatechol degradation, is initially transformed to maleylacetate followed by reduction to 3-oxoadipate (Kaschabek and Reineke 1992). Obviously, enzymatic attack on the C2-carbon results in an intermediate, which spontaneously eliminates chloride. Probably, dehalogenation of 2-chloromaleylacetate is a general capability of maleylacetate reductases. The ortho-cleavage chlorocatechol pathway described tolerates substitution at the aromatic ring of up to four chlorine atoms. Two dechlorination steps have been described up to now. The genes encoding the chlorocatechol pathway usually form clusters and the structures of the corresponding operons of chlorobenzoate degraders such as P. knackmussii B13 or C. necator NH9 or chlorobenzene degraders such as Pseudomonas sp. P51 or P. chlororaphis RW71 are nearly identical, in spite of the geographically distinct origins of the bacteria or the difference in their phylogenetic position (Pieper 2005). Interestingly, the genetic organization of microorganisms isolated based on their capability to degrade 2,4-D usually differs substantially from that of chlorobenzoate and chlorobenzene degraders already described and in most strains the module comprises a gene encoding a phenol hydroxylase (tfdB), a protein probably involved in 2,4-D transport (tfdK) and a 2,4-D/α-ketoglutarate dioxygenase (tfdA). These gene clusters are typically missing tfdD and tfdF genes encoding chloromuconate cycloisomerases and maleylacetate reductases, respectively. However, in strains analyzed in this aspect, tfdD genes are located upstream of the tfdR regulator gene, whereas tfdF genes are located downstream of tfdA (Pieper 2005). Even though analysis of chlorocatechol gene operons in proteobacteria had concentrated on β- and γ-Proteobacteria, chlorocatechol genes have been also characterized also in members of the α- Proteobacteria (Müller et al. 2004). Both types of newly discovered chlorocatechol dioxygenases from Sphingomonas isolates represent deep-branching lines in the dendrogram and suggest a different reservoir and reduced transfer of the respective genes in α-Proteobacteria compared with the ones in β- and γ-Proteobacteria. Only poor information is available on chlorocatechol genes in Actinobacteria, and thus far, only two modules from Rhodococcus opacus 1CP have been analyzed in detail (Maltseva et al. 1994a; Moiseeva et al. 2002).

18

D.H. Pieper et al.

Usually, chlorocatechol 1,2-dioxygenases are characterized by their broad substrate specificity and can convert several higher chlorinated catechols. As an example, the chlorocatechol gene products of P. knackmussii B13 were capable of dealing with higher chlorinated catechols such as 3,5-dichloro- or 3,6-dichlorocatechol, whereas 3,4-dichlorocatechol was only a poor substrate (Dorn and Knackmuss 1978; Oltmanns et al. 1988). In contrast, the enzyme of Pseudomonas sp. strain P51 was capable of transforming 3,4-dichlorocatechol (van der Meer et al. 1991). Recently, amino acids determining such differences in substrate specificity could be identified (Liu et al. 2005). In the case of chlorocatechol 1,2-dioxygenases from Rhodococcus, only one of the thus-far described enzymes exhibits broad substrate specificity (Moiseeva et al. 2002), whereas 4-chlorocatechol 1,2-dioxygenase prefers 4-chlorocatechol and has only negligible activity with 3-chlorocatechol (Maltseva et al. 1994a). All proteobacterial CMCIs obviously share common features, which discriminate them from the Rhodococcus enzymes, but also from MCIs. In agreement with their special biochemical properties, a phylogenetic comparison depicted proteobacterial CMCIs to constitute a separate subfamily, whereas the rhodococcal CMCIs, like the respective chlorocatechol 1,2-dioxygenases, appear to represent isolated evolutionary lines (Fig. 8). As already described, proteobacterial CMCIs are specific dehalogenases, and capable of dehalogenating even higher substituted chloromuconates. In contrast, both CMCIs described from R. opacus 1CP exhibited extraordinary properties and catalyzed the formation of 5-chloromuconolactone from 2-chloromuconate but were not capable of dehalogenating (Moiseeva et al. 2002; Solyanikova et al. 1995) (Fig. 7). Only some of the DLHs encoded in chlorocatechol gene clusters such as TfdEI from C. necator JMP134, and especially ClcD from P. knackmussii B13, have been described in more detail. These enzymes are active against both the cis- and transisomer, a feature that is indispensable for the mineralization of differently substituted chloroaromatics by proteobacteria as already described. Structure elucidation of the P. knackmussii B13 enzyme revealed that the protein belongs to the α/β hydrolase fold enzymes (Ollis and Nai 1985). Analysis of DLHs from various chloroaromatic degraders showed that even though some of these DLHs share only low sequence identity (down to 13%), they all belong to the same family with a conserved CysHis-Asp triad and highly conserved residues flanking the triad; however they all differ from enol-lactone hydrolases (Schlömann 1994). Unfortunately, only poor data are available on the substrate specificity of those DLHs; however, both DLHs from Rhodococcus are active only against the cis-isomer (Maltseva et al. 1994b; Moiseeva et al. 2002), contrasting DLHs from proteobacterial chlorobenzoate or chlorobenzene degraders.

2.2.3

The Alternative Pathway of 3-Chlorocatechol Degradation in Rhodococcus opacus 1CP Among the Gram-positive organisms, chlorocatechol catabolism has mainly been investigated in R. opacus 1CP. 4-chloro- and 3,5-dichlorocatechol are transformed similar to the metabolism in proteobacteria, whereas 2-chloromuconate formed from

Aerobic Degradation of Chloroaromatics

19

3-chlorocatechol is not dehalogenated during cycloisomerization. In contrast, dehalogenation is catalyzed by a muconolactone isomerase-related enzyme (Fig. 7). Despite its sequence similarity to muconolactone isomerase, the Rhodococcus enzyme does not show muconolactone-isomerizing activity and thus represents an enzyme dedicated to its new function as a 5-chloromuconolactone dehalogenase forming cis-dienelactone (Moiseeva et al. 2002). Thus, a major difference between chlorocatechol degradation in proteobacteria and in Rhodococcus lies in the fact that cis-dienelactone is the intermediate of both 4-chloro- and 3-chlorocatechol degradation in Rhodococcus, whereas cis- and trans-dienelactone, respectively, are formed by proteobacteria, therefore requiring a DLH with broad substrate specificity (Fig. 7).

2.2.4 The Alternative Pathway of 4-Halocatechol Degradation Another variant of the chlorocatechol pathway has been described in Pseudomonas reinekei MT1 (Nikodem et al. 2003). In this strain 4-chlorocatechol formed from 4-chloro- and 5-chlorosalicylate is channeled into the 3-oxoadipate pathway by a catechol 1,2-dioxygenase and a MCI that showed unusual kinetic properties as being adapted for turnover of 4-chlorocatechol and 3-chloromuconate, respectively (Cámara et al. 2007). A trans-dienelactone hydrolase capable of transforming the trans- but not the cis-dienelactone isomer has been observed to be crucial for 4-chlorocatechol degradation by this strain. It was shown that the enzyme interacts with the MCI-catalyzed transformation of 3-chloromuconate and hydrolyzes the cycloisomerization intermediate 4-chloromuconolactone, thereby preventing the formation of protoanemonin in favor of maleylacetate (Nikodem et al. 2003) (Fig. 7). Maleylacetate is then transformed by maleylacetate reductase. DLHs active against trans-dienelactone only and not able to transform cis-dienelactone, the intermediate of 4-chlorocatechol degradation by enzymes of the chlorocatechol pathway, have initially been described to be involved in 4-fluorocatechol degradation by C. necator 335 and C. necator JMP222 and probably responsible for further degradation of intermediate 4-fluoromuconolactone (Schlömann et al. 1990). 2.2.5 Chlorocatechol Degradation via the Meta-Cleavage Pathway Catechol meta-cleavage routes are widespread and usually involved in the degradation of methylsubstituted compounds such as toluene or methylphenols. For a long time the presence of such a meta-cleavage pathway was assumed to severely interfere with the degradation of chloroaromatics. One of the reasons of interference was assumed to be the formation of a reactive acyl chloride, e.g., from 3-chlorocatechol by the catechol 2,3-dioxygenase of P. putida mt-2 (XylE) (Bartels et al. 1984), resulting in irreversible inactivation of the ring cleavage enzyme (Fig. 9). In other cases, reversible inactivation was shown to be due to a rapid oxidation of the active site ferrous iron to its ferric form with concomitant loss of activity (Vaillancourt et al. 2002), and a general mechanism for the inactivation of extradiol dioxygenases during catalytic turnover involving the dissociation of superoxide from the enzyme-catechol-dioxygen ternary complex was suggested. In contrast to 3-chlorocatechol, 4-chlorocatechol is a moderate substrate for various

20

D.H. Pieper et al.

Fig. 9 Meta-cleavage pathways for catechol, 3-chlorocatechol, protocatechuate and 5-chloroprotocatechuate. The acylchlorides are shown in brackets. The 3-chlorocatechol ringcleavage product can react either with the ring cleavage enzyme resulting in suicide inactivation, or be rapidly hydrolyzed to give 2-hydroxymuconate. The 5-chloroprotocatechuate ring-cleavage product undergoes intramolecular rearrangement and dehalogenation

catechol 2,3-dioxygenases (Bartels et al. 1984; Murray et al. 1972), among them catechol 2,3-dioxygenases of family I.2.A (Eltis and Bolin 1996), which have been commonly observed to be involved in the degradation of methylaromatics. However, despite high sequence identity, members of this subfamily exhibit very different substrate specificities, and the capability to transform 4-chlorocatechol is not a general characteristic of catechol 2,3-dioxygenases of family I.2.A (Kitayama et al. 1996). Some publications postulate that compounds degraded via catechols chlorinated in the 4-position might be mineralized via a catechol degrading metacleavage pathway (Arensdorf and Focht 1995) but information about the way in which the products are dechlorinated is missing. In 1997, the first isolate, P. putida GJ31, capable of degrading a chloroaromatic compound (chlorobenzene) via 3-chlorocatechol using a meta-cleavage pathway

Aerobic Degradation of Chloroaromatics

21

was described (Mars et al. 1997). The CbzE chlorocatechol 2,3-dioxygenase of strain GJ31 productively converts 3-chlorocatechol, and stoichiometric displacement of chloride leads to the production of 2-hydroxymuconate (Kaschabek et al. 1998), which is further converted through the meta-cleavage pathway (Fig. 9). 3-Chlorocatechol was found to be the preferred substrate of CbzE. Additional pseudomonads using a meta-cleavage route for 3-chlorocatechol degradation were isolated in the meantime from various contaminated environments (Göbel et al. 2004), suggesting that productive meta-cleavage of 3-chlorocatechol is not atypical for chloroaromatic degradation. All the three isolates harbor chlorocatechol 2,3-dioxygenases, highly resistant to inactivation during 3-chlorocatechol turnover, sharing 97% amino acid sequence identity with the enzyme from strain GJ31, thus forming a distinct subfamily of catechol 2,3-dioxygenases in what was termed family I.2.C. Other extradiol dioxygenases of family 1.2.C do not share the capability of effectively transforming 3-chlorocatechol. In all the chlorobenzene-degrading strains already mentioned, cbzT genes encoding ferredoxins are located upstream of cbzE. Similar ferredoxins have been observed to be encoded in various meta-cleavage pathways (Hugo et al. 2000). Catechol 2,3-dioxygenases are oxygen sensitive and unstable in vitro, particularly in the presence of substituted catechol substrates. This instability was shown to be due to the oxidation of active site Fe(II)–Fe(III) (Vaillancourt et al. 2002), and CbzT-like ferredoxins reactivate catechol 2,3-dioxygenases through the reduction of the iron atom in the active site of the enzyme (Hugo et al. 2000). It was thus proposed that the ability of strain GJ31 to metabolize both chlorobenzene and toluene might depend on the regeneration of the chlorocatechol dioxygenase activity mediated by CbzT (Tropel et al. 2002).

2.2.6

Degradation of 5-Chloroprotocatechuate via a Meta-Cleavage Pathway A productive meta-cleavage without suicide effect has been known for many years. The extradiol cleavage of 5-chloroprotocatechuate by protocatechuate 4,5-dioxygenase (Kersten et al. 1985) results in the formation of 2-pyrone-4,6dicarboxylate by nucleophilic displacement of chloride (Fig. 9). This indicates that cyclization entailing nucleophilic displacement of halogen provides an effective alternative to the enzyme suicide inactivation that occurs when a nucleophilic group of the extradiol dioxygenase undergoes acylation. An important aspect of this mechanism is that the ring fission product remains bound to the enzyme during the complete configuration change that precedes nucleophilic displacement. 2-Pyrone-4,6-dicarboxylate is a metabolite of the protocatechuate 4,5-dioxygenase pathway typically formed after the dehydrogenation of the ring-cleavage product (see Fig. 9), such that both protocatechuate and 5-chloroprotocatechuate degradation merge at this metabolic intermediate. So far, only the mineralization of 3-chlorobenzoate by C. testosteroni BR60 (Nakatsu et al. 1997) has been proposed to occur via such a route.

22

D.H. Pieper et al.

Fig. 10 Catabolic pathways for (chloro)benzoquinols and (chloro)hydroxybenzoquinols. Unstable intermediates are enclosed in brackets. Broken arrows indicate reactions of the upper pathway detailed in Fig. 6

2.3

Ring Cleavage of (Chloro)Benzoquinols and (Chloro) Hydroxybenzoquinols

Ring cleavage dioxygenases involved in the turnover of (chloro) benzoquinols and (chloro) hydroxybenzoquinols has been identified from various microorganisms degrading γ-HCH, or chlorophenols (Fig. 10). Chlorobenzoquinol (but also benzoquinol) produced during the catabolism of γ-HCH in S. japonicum UT26 is subject to direct ring cleavage by a novel type of extradiol dioxygenase, designated chlorobenzoquinol/benzoquinol 1,2-dioxygenase (LinE), which preferentially cleaves aromatic rings with two hydroxyl groups at para positions (Miyauchi et al. 1999) and produces maleylacetate from either chlorobenzoquinol or benzoquinol (Fig. 10). Highly similar or identical dioxygenases (>99% sequence identity) have been identified in other γ-HCH-degrading organisms (Dogra et al. 2004). More distantly related extradiol dioxygenases (51–53% sequence identity) and designated 2,6-dichlorobenzoquinol 1,2-dioxygenases (PcpA) were observed in PCP degrading strains such as S. chlorophenolicum ATCC 39723 (Xu et al. 1999) and produce

Aerobic Degradation of Chloroaromatics

23

2-chloromaleylacetate (2-CMA) from 2,6-dichlorobenzoquinol (Fig. 10). A protein highly similar to PcpA proteins and with activity with 2,6-dichlorobenzoquinol was also observed in S. japonicum UT26 (LinEb exhibiting 92% sequence identity to PcpA proteins (Endo et al. 2005)). However, its involvement in γ-HCH degradation responsible for cleavage of 2,5-dichlorobenzoquinol is unlikely, because such a cleavage would result in the formation of 3-chloromaleylacetate, which cannot be dehalogenated by maleylacetate reductase. In contrast, members of the intradiol dioxygenase family are involved in the degradation of hydroxybenzoquinols formed from trichlorophenols, 2,4-dichloroor 4-chlorophenol, and these (chloro)hydroxybenzoquinol dioxygenases form a defined phylogenetic group inside the intradiol dioxygenase family. Hydroxybenzoquinol is cleaved by TftH hydroxybenzoquinol 1,2-dioxygenase to yield maleylacetate during the mineralization of 2,4,5-T by B. phenoliruptrix AC1100 (Daubaras et al. 1996) (Fig. 10). In 2,4,6-TCP degradation by C. necator JMP134, TcpC 6-chlorohydroxyquinol 1,2-dioxygenase is responsible for direct cleavage of 6-chlorohydroxybenzoquinol producing 2-chloromaleylacetate (Louie et al. 2002) (Fig. 10). An important difference between the pathways of 2,4,5-TCP and 2,4,6TCP degradation is the fact that 6-chlorohydroxybenzoquinol is directly cleaved instead of a preceding reductive dechlorination forming hydroxybenzoquinol. Interestingly, TftH hydroxybenzoquinol 1,2-dioxygenase from B. phenoliruptrix AC1100, which shares 53% amino acid sequence identity with TcpC, is unable to use 5-chloro- or 6-chlorohydroxybenzoquinol (Daubaras et al. 1996). The catabolism of (chloro)benzoquinols and (chloro)hydroxybenzoquinols is completed by the conversion of MA or 2-CMA to 3-oxoadipate (Fig. 10) by the action of maleylacetate reductases as already described (see Sect. 2.2.2).

3

Research Needs

Most of the current knowledge on biochemistry and genetics of chloroaromatic aerobic microbial metabolism has been obtained with bacterial strains isolated by traditional culture-dependent approaches. This is a severe drawback to understanding microbial degradation of chloroaromatics, as only a very minor proportion of microbes inhabiting the different ecosystems can be isolated by standard procedures and only a few reports are available on microbial associations (consortia) degrading this kind of compounds. Although it is possible that current knowledge covers a significant proportion of microbial strategies to degrade chloroaromatics, it would also be possible that new, unexpected ways to metabolize these compounds remain still unknown in the significant fraction of yet-uncultured microbes. Functional metagenomics approaches are required to address this point. Since natural haloaromatic formation is increasingly evident, studies including diverse ecosystems, not necessarily exposed to chloroaromatic pollutants, will also provide a better idea on the diversity of metabolic pathways for aerobic degradation of chloroaromatics.

24

D.H. Pieper et al.

Several cases of adaptation or recruitment of pathways involved in the metabolism of natural aromatics for metabolism of the corresponding chloroaromatic counterparts are already reported. A further exploration of these possibilities would provide new insights into these phenomena. For instance, degradation of aromatic compounds through CoA derivatives seems to play a more significant role in aerobic degradation than initially expected. CoA derivatives have been reported as intermediates of 4-chlorobenzoate degradation, but additional routes are expected to be discovered. Available are only a few studies of the degradation of mixed substituted chloroaromatic compounds, i.e., carrying additional amino/nitro/sulfonic substituents or heterocyclic aromatics, which without doubt, may give interesting new insights into aerobic microbial degradation of pollutants. Even for the pathways already reported, there are several unclarified questions that deserve attention. For example, by which mechanism is the third or fourth chlorosubstituent eliminated during trichloro- or tetrachlorocatechol metabolism? What are the biochemical grounds for the degradation of 4-chlorocatechol via the meta-cleavage? By which mechanism is 3,6-dichlorogentisate mineralized in dicamba degrading microorganisms?

References Arensdorf JJ, Focht DD (1995) A meta cleavage pathway for 4-chlorobenzoate, an intermediate in the metabolism of 4-chlorobiphenyl by Pseudomonas cepacia P166. Appl Environ Microbiol 61:443–447 Babbitt PC, Kenyon GL, Martin BM, Charest H, Sylvestre M, Scholten JD, Chang K-H, Liang P-H, Dunaway-Mariano D (1992) Ancestry of the 4-chlorobenzoate dehalogense: analysis of amino acid sequence identities among families of acyl: adenyl ligases, enoyl-CoA hydratases/isomerases, and acyl-CoA thiosterases. Biochemistry 31:5594–5604 Bartels I, Knackmuss H-J, Reineke W (1984) Suicide inactivation of catechol 2,3-dioxygenase from Pseudomonas putida mt-2 by 3-halocatechols. Appl Environ Microbiol 47:500–505 Beadle CA, Smith ARW (1982) The purification and properties of 2,4-dichlorophenol hydroxylase from a strain of Acinetobacter species. Eur J Biochem 123:323–332 Beil S, Mason JR, Timmis KN, Pieper DH (1998) Identification of chlorobenzene dioxygenase sequence elements involved in dechlorination of 1,2,4,5-tetrachlorobenzene. J Bacteriol 180:5520–5528 Belchik SM, Xun L (2008) Functions of flavin reductase and quinone reductase in 2,4,6trichlorophenol degradation by Cupriavidus necator JMP134. J Bacteriol 190:1615–1619 Benning MM, Wesenberg G, Liu RQ, Taylor KL, Dunaway-Mariano D, Holden HM (1998) The three-dimensional structure of 4-hydroxybenzoyl-CoA thioesterase from Pseudomonas sp. strain CBS-3. J Biol Chem 273:33572–33579 Blasco R, Wittich R-M, Mallavarapu M, Timmis KN, Pieper DH (1995) From xenobiotic to antibiotic. Formation of protoanemonin from 4-chlorocatechol by enzymes of the 3-oxoadipate pathway. J Biol Chem 270:29229–29235 Bosch R, Moore ERB, GarciaValdes E, Pieper DH (1999) NahW, a novel, inducible salicylate hydroxylase involved in mineralization of naphthalene by Pseudomonas stutzeri AN10. J Bacteriol 181:2315–2322

Aerobic Degradation of Chloroaromatics

25

Brinkmann U, Reineke W (1992) Degradation of chlorotoluenes by in vivo constructed hybrid strains: problems of enzyme specificity, induction and prevention of meta-pathway. FEMS Microbiol Lett 96:81–88 Cafaro V, Izzo V, Scognamiglio R, Notomista E, Capasso P, Casbarra A, Pucci P, Di Donato A (2004) Phenol hydroxylase and toluene/o-xylene monooxygenase from Pseudomonas stutzeri OX1: interplay between two enzymes. Appl Environ Microbiol 70:2211–2219 Cámara B, Bielecki P, Kaminski F, dos Santos VM, Plumeier I, Nikodem P, Pieper DH (2007) A gene cluster involved in degradation of substituted salicylates via ortho cleavage in Pseudomonas sp. strain MT1 encodes enzymes specifically adapted for transformation of 4-methylcatechol and 3-methylmuconate. J Bacteriol 189:1664–1674 Crawford RL, Jung CM, Strap JL (2007) The recent evolution of pentachlorophenol (PCP)-4monooxygenase (PcpB) and associated pathways for bacterial degradation of PCP. Biodegradation 18:525–539 Dai MH, Rogers JB, Warner JR, Copley SD (2003) A previously unrecognized step in pentachlorophenol degradation in Sphingobium chlorophenolicum is catalyzed by tetrachlorobenzoquinone reductase (PcpD). J Bacteriol 185:302–310 Danganan CE, Shankar S, Ye RW, Chakrabarty AM (1995) Substrate diversity and expression of the 2,4,5-trichlorophenoxyacetic acid oxygenase from Burkholderia cepacia AC1100. Appl Environ Microbiol 61:4500–4504 Daubaras DL, Saido K, Chakrabarty AM (1996) Purification of hydroxyquinol 1,2-dioxygenase and maleylacetate reductase: the lower pathway of 2,4,5-trichlorophenoxyacetic acid metabolism by Burkholderia cepacia AC1100. Appl Environ Microbiol 62:4276–4279 Dillon SC, Bateman A (2004) The Hotdog fold: wrapping up a superfamily of thioesterases and dehydratases. BMC Bioinform 12:109 Dogra C, Raina V, Pal R, Suar M, Lal S, Gartemann KH, Holliger C, van der Meer JR, Lal R (2004) Organization of lin genes and IS6100 among different strains of hexachlorocyclohexanedegrading Sphingomonas paucimobilis: evidence for horizontal gene transfer. J Bacteriol 186:2225–2235 Dong J, Carey PR, Wei YS, Luo LS, Lu XF, Liu RQ, Dunaway-Mariano D (2002) Raman evidence for Meisenheimer complex formation in the hydrolysis reactions of 4-fluorobenzoyl- and 4-nitrobenzoyl-coenzyme a catalyzed by 4-chlorobenzoyl-coenzyme A dehalogenase. Biochemistry 41:7453–7463 Dorn E, Knackmuss H-J (1978) Chemical structure and biodegradability of halogenated aromatic compounds. Substituent effects on 1,2-dioxygenation of catechol. Biochem J 174:85–94 Duarte M, Jauregui R, Vilchez-Vargas R, Junca H, Pieper DH (2014) AromaDeg, a novel database for phylogenomics of aerobic degradation of aromatics. Database 2014; bau118 Eltis LD, Bolin JT (1996) Evolutionary relationships among extradiol dioxygenases. J Bacteriol 178:5930–5937 Endo R, Kamakura M, Miyauchi K, Fukuda M, Ohtsubo Y, Tsuda M, Nagata Y (2005) Identification and characterization of genes involved in the downstream degradation pathway of gammahexachlorocyclohexane in Sphingomonas paucimobilis UT26. J Bacteriol 187:847–853 Ferraroni M, Seifert J, Travkin VM, Thiel M, Kaschabek S, Scozzafava A, Golovleva L, Schlömann M, Briganti F (2005) Crystal structure of the hydroxyquinol 1,2-dioxygenase from Nocardioides simplex 3E, a key enzyme involved in polychlorinated aromatics biodegradation. J Biol Chem 280:21144–21154 Fetzner S, Müller R, Lingens F (1992) Purification and some properties of 2-halobenzoate 1,2-dioxygenase, a two component enzyme system from Pseudomonas cepacia 2CBS. J Bacteriol 174:279–290 Fishman A, Tao Y, Rui L, Wood TK (2005) Controlling the regiospecific oxidation of aromatics via active site engineering of toluene para-monooxygenase of Ralstonia pickettii PKO1. J Biol Chem 280:506–514 Fukumori F, Hausinger RP (1993) Alcaligenes eutrophus JMP134 “2,4-dichlorophenoxyacetate monooxygenase” is an a-ketoglutarate-dependent dioxygenase. J Bacteriol 175:2083–2086

26

D.H. Pieper et al.

Furukawa K, Suenaga H, Goto M (2004) Biphenyl dioxygenases: functional versatilities and directed evolution. J Bacteriol 186:5189–5196 Gibson DT, Parales RE (2000) Aromatic hydrocarbon dioxygenases in environmental biotechnology. Curr Opin Biotechnol 11:236–243 Gisi MR, Xun L (2003) Characterization of chlorophenol 4-monooxygenase (TftD) and NADH: flavin adenine dinucleotide oxidoreductase (TftC) of Burkholderia cepacia AC1100. J Bacteriol 185:2786–2792 Göbel M, Kranz OH, Kaschabek SR, Schmidt E, Pieper DH, Reineke W (2004) Microorganisms degrading chlorobenzene via a meta-cleavage pathway harbor highly similar chlorocatechol 2,3-dioxygenase-encoding gene clusters. Arch Microbiol 182:147–156 Greated A, Lambertsen L, Williams PA, Thomas CM (2002) Complete sequence of the IncP-9 TOL plasmid pWW0 from Pseudomonas putida. Environ Microbiol 4:856–871 Hübner A, Danganan CE, Xun LY, Chakrabarty AM, Hendrickson W (1998) Genes for 2,4,5trichlorophenoxyacetic acid metabolism in Burkholderia cepacia AC1100: characterization of the tftC and tftD genes and locations of the tft operons on multiple replicons. Appl Environ Microbiol 64:2086–2093 Hugo N, Meyer C, Armengaud J, Gaillard J, Timmis KN, Jouanneau Y (2000) Characterization of three XylT-like [2Fe-2S] ferredoxins associated with catabolism of cresols or naphthalene: evidence for their involvement in catechol dioxygenase reactivation. J Bacteriol 182: 5580–5585 Huong NL, Itoh K, Miyamoto M, Suyama K, Yamamoto H (2007) Chlorophenol hydroxylase activity encoded by TfdB from 2,4-dichlorophenoxyacetic acid (2,4-D)-degrading Bradyrhizobium sp. strain RD5-C2. Biosci Biotechnol Biochem 71:1691–1696 Itoh K, Kanda R, Sumita Y, Kim H, Kamagata Y, Suyama K, Yamamoto H, Hausinger RP, Tiedje JM (2002) tfdA-like genes in 2,4-dichlorophenoxyacetic acid-degrading bacteria belonging to the Bradyrhizobium-Agromonas-Nitrobacter-Afipia cluster in alpha-proteobacteria. Appl Environ Microbiol 68:3449–3454 Itoh K, Tashiro Y, Uobe K, Kamagata Y, Suyama K, Yamamoto H (2004) Root nodule Bradyrhizobium spp. harbor tfdAalpha and cadA, homologous with genes encoding 2,4-dichlorophenoxyacetic acid-degrading proteins. Appl Environ Microbiol 70:2110–2118 Kahng HY, Malinverni JC, Majko MM, Kukor JJ (2001) Genetic and functional analysis of the tbc operons for catabolism of alkyl- and chloroaromatic compounds in Burkholderia sp. strain JS150. Appl Environ Microbiol 67:4805–4816 Kaschabek SR, Reineke W (1992) Maleylacetate reductase of Pseudomonas sp. strain B13: dechlorination of chloromaleylacetates, metabolites in the degradation of chloroaromatic compounds. Arch Microbiol 158:412–417 Kaschabek SR, Kasberg T, Müller D, Mars AE, Janssen DB, Reineke W (1998) Degradation of chloroaromatics: purification and characterization of a novel type of chlorocatechol 2,3-dioxygenase of Pseudomonas putida GJ31. J Bacteriol 180:296–302 Kaulmann U, Kaschabek SR, Schlömann M (2001) Mechanism of chloride elimination from 3-chloro- and 2,4-dichloro- cis, cis-muconate: new insight obtained from analysis of muconate cycloisomerase variant CatB-K169A. J Bacteriol 183:4551–4561 Kersten P, Chapman PJ, Dagley S (1985) Enzymatic release of halogens or methanol from some substituted protocatechuic acids. J Bacteriol 162:693–697 Kitagawa W, Takami S, Miyauchi K, Masai E, Kamagata Y, Tiedje JM, Fukuda M (2002) Novel 2,4-dichlorophenoxyacetic acid degradation genes from oligotrophic Bradyrhizobium sp. strain HW13 isolated from a pristine environment. J Bacteriol 184:509–518 Kitayama A, Suzuki E, Kawakami Y, Nagamune T (1996) Gene organization and low regioselectivity in aromatic-ring hydroxylation of a benzene monooxygenase of Pseudomonas aeruginosa JI104. J Ferment Bioeng 82:421–425 Leahy JG, Batchelor PJ, Morcomb SM (2003) Evolution of the soluble diiron monooxygenases. FEMS Microbiol Rev 27:449–479

Aerobic Degradation of Chloroaromatics

27

Ledger T, Pieper DH, Gonzalez B (2006) Chlorophenol hydroxylases encoded by plasmid pJP4 differentially contribute to chlorophenoxyacetic acid degradation. Appl Environ Microbiol 72:2783–2792 Lehrbach PR, Zeyer J, Reineke W, Knackmuss HJ, Timmis KN (1984) Enzyme recruitment in vitro: use of cloned genes to extend the range of haloaromatics degraded by Pseudomonas sp. strain B13. J Bacteriol 158:1025–1032 Liang P-H, Yang G, Dunaway-Mariano D (1993) Specificity of 4-chlorobenzoyl coenzyme A dehalogenase catalyzed dehalogenation of a halogenated aromatics. Biochemistry 32:12245–12250 Liu S, Ogawa N, Senda T, Hasebe A, Miyashita K (2005) Amino acids in positions 48, 52, and 73 differentiate the substrate specificities of the highly homologous chlorocatechol 1,2-dioxygenases CbnA and TcbC. J Bacteriol 187:5427–5436 Louie TM, Webster CM, Xun LY (2002) Genetic and biochemical characterization of a 2,4,6trichlorophenol degradation pathway in Ralstonia eutropha JMP134. J Bacteriol 184:3492–3500 Makdessi K, Lechner U (1997) Purification and characterization of 2,4-dichlorophenol hydroxylase isolated from a bacterium of the alpha-2 subgroup of the Proteobacteria. FEMS Microbiol Lett 157:95–101 Maltseva OV, Solyanikova IP, Golovleva LA (1994a) Chlorocatechol 1,2-dioxygenase from Rhodococcus erythropolis 1CP. Kinetic and immunochemical comparison with analogous enzymes from Gram-negative strains. Eur J Biochem 226:1053–1061 Maltseva OV, Solyanikova IP, Golovleva LA, Schlömann M, Knackmuss H-J (1994b) Dienelactone hydrolase from Rhodococcus erythropolis 1CP: purification and properties. Arch Microbiol 162:368–374 Mars AE, Kasberg T, Kaschabek SR, van Agteren MH, Janssen DB, Reineke W (1997) Microbial degradation of chloroaromatics: use of the meta-cleavage pathway for mineralization of chlorobenzene. J Bacteriol 179:4530–4537 Matsumura E, Ooi S, Murakami S, Takenaka S, Aoki K (2004) Constitutive synthesis, purification, and characterization of catechol 1,2-dioxygenase from the aniline-assimilating bacterium Rhodococcus sp AN-22. J Biosci Bioeng 98:71–76 Matus V, Sánchez MA, Martínez M, González B (2003) Efficient degradation of 2,4,6trichlorophenol requires a set of catabolic genes related to tcp genes from Ralstonia eutropha JMP134 (pJP4). Appl Environ Microbiol 69:7108–7115 McGowan C, Fulthorpe RR, Wright A, Tiedje JM (1998) Evidence for interspecies gene transfer in the evolution of 2,4-dichlorophenoxyacetic acid degraders. Appl Environ Microbiol 64:4089–4092 Mitchell KH, Studts JM, Fox BG (2002) Combined participation of hydroxylase active site residues and effector protein binding in a para to ortho modulation of toluene 4-monooxygenase regiospecificity. Biochemistry 41:3176–3188 Miyauchi K, Suh SK, Nagata Y, Takagi M (1998) Cloning and sequencing of a 2,5-dichlorohydroquinone reductive dehalogenase gene whose product is involved in degradation of gamma-hexachlorocyclohexane by Sphingomonas paucimobilis. J Bacteriol 180:1354–1359 Miyauchi K, Adachi Y, Nagata Y, Takagi M (1999) Cloning and sequencing of a novel metacleavage dioxygenase gene whose product is involved in degradation of gammahexachlorocyclohexane in Sphingomonas paucimobilis. J Bacteriol 181:6712–6719 Moiseeva OV, Solyanikova IP, Kaschabek SR, Groning J, Thiel M, Golovleva LA, Schlömann M (2002) A new modified ortho cleavage pathway of 3-chlorocatechol degradation by Rhodococcus opacus 1CP: genetic and biochemical evidence. J Bacteriol 184:5282–5292 Müller TA, Byrde SM, Werlen C, van der Meer JR, Kohler HP (2004) Genetic analysis of phenoxyalkanoic acid degradation in Sphingomonas herbicidovorans MH. Appl Environ Microbiol 70:6066–6075

28

D.H. Pieper et al.

Müller TA, Fleischmann T, van der Meer JR, Kohler HP (2006) Purification and characterization of two enantioselective alpha-ketoglutarate-dependent dioxygenases, RdpA and SdpA, from Sphingomonas herbicidovorans MH. Appl Environ Microbiol 72:4853–4861 Murray K, Duggleby CJ, Sala-Trepat JM, Williams PA (1972) The metabolism of benzoate and methylbenzoates via the meta-cleavage by Pseudomonas arvilla mt-2. Eur J Biochem 28:301–310 Nagata Y, Ohtomo R, Miyauchi K, Fukuda M, Yano K, Takagi M (1994) Cloning and sequencing of a 2,5-dichloro-2,5-cyclohexadiene-1,4-diol dehydrogenase gene involved in the degradation of gamma-hexachlorocyclohexane on Pseudomonas paucimobilis. J Bacteriol 176:3117–3125 Nagata Y, Endo R, Ito M, Ohtsubo Y, Tsuda M (2007) Aerobic degradation of lindane (gammahexachlorocyclohexane) in bacteria and its biochemical and molecular basis. Appl Microbiol Biotechnol 76:741–752 Nakatsu CH, Wyndham RC (1993) Cloning and expression of the transposable chlorobenzoate-3, 4-dioxygenase genes of Alcaligenes sp. strain BR60. Appl Environ Microbiol 59:3625–3633 Nakatsu CH, Providenti M, Wyndham RC (1997) The cis-diol dehydrogenase cbaC gene of Tn5271 is required for growth on 3-chlorobenzoate but not 3,4-dichlorobenzoate. Gene 196:209–218 Nikodem P, Hecht V, Schlömann M, Pieper DH (2003) New bacterial pathway for 4- and 5-chlorosalicylate degradation via 4-chlorocatechol and maleylacetate in Pseudomonas sp. strain MT1. J Bacteriol 185:6790–6800 Nordin K, Unell M, Jansson JK (2005) Novel 4-chlorophenol degradation gene cluster and degradation route via hydroxyquinol in Arthrobacter chlorophenolicus A6. Appl Environ Microbiol 71:6538–6544 Nurk A, Kasak L, Kivisaar M (1991) Sequence of the gene (pheA) encoding phenol monooxygenae from Pseudomonas sp. EST1001: expression in Escherichia coli and Pseudomonas putida. Gene 102:13–18 Ollis DL, Nai KL (1985) Crystallization and preliminary x-ray crystallographic data of dienelactone hydrolase from Pseudomonas sp. B13. J Biol Chem 260:9818–9819 Oltmanns RH, Rast HG, Reineke W (1988) Degradation of 1,4-dichlorobenzene by enriched and constructed bacteria. Appl Microbiol Biotechnol 28:609–616 Pieper DH (2005) Aerobic degradation of polychlorinated biphenyls. Appl Microbiol Biotechnol 67:170–191 Pieper DH, Kuhm AE, Stadler-Fritzsche K, Fischer P, Knackmuss H-J (1991) Metabolization of 3,5-dichlorocatechol by Alcaligenes eutrophus JMP134. Arch Microbiol 156:218–222 Prucha M, Peterseim A, Timmis KN, Pieper DH (1996) Muconolactone isomerase of the 3-oxoadipate pathway catalyzes dechlorination of 5-chloro-substituted muconolactones. Eur J Biochem 237:350–356 Reineke W (2001) Aerobic and anaerobic biodegradation potentials of microorganisms. In: Hutzinger O (ed) The handbook of environmental chemistry. Springer, Berlin, pp 1–161 Romanov V, Hausinger RP (1994) Pseudomonas aeruginosa 142 uses a three-component orthohalobenzoate 1,2-dioxygenase for the metabolism of 2,4-dichloro- and 2-chlorobenzoate. J Bacteriol 176:3368–3374 Savard P, Charest H, Sylvestre M, Shareck F, Scholten JD, Dunaway Mariano D (1992) Expression of the 4-chlorobenzoate dehalogenase genes from Pseudomonas sp. CBS3 in Escherichia coli and identification of the gene translation products. Can J Microbiol 38:1074–1083 Schell U, Helin S, Kajander T, Schlömann M, Goldman A (1999) Structural basis for the activity of two muconate cycloisomerase variants toward substituted muconates. Proteins Struct Funct Genet 34:125–136 Schlömann M (1994) Evolution of chlorocatechol catabolic pathways. Conclusions to be drawn from comparisons of lactone hydrolases. Biodegradation 5:301–321 Schlömann M, Schmidt E, Knackmuss H-J (1990) Different types of dienelactone hydrolase in 4-fluorobenzoate-utilizing bacteria. J Bacteriol 172:5112–5118

Aerobic Degradation of Chloroaromatics

29

Schmidt E, Knackmuss H-J (1980) Chemical structure and biodegradability of halogenated aromatic compounds. Conversion of chlorinated muconic acids into maleoylacetic acid. Biochem J 192:339–347 Schmitz A, Gartemann K-H, Fiedler J, Grund E, Eichenlaub R (1992) Cloning and sequence analysis of genes for dehalogenation of 4-chlorobenzoate from Arthrobacter sp. strain SU. Appl Environ Microbiol 58:4068–4071 Schofield CJ, Zhang Z (1999) Structural and mechanistic studies on 2-oxoglutarate-dependent oxygenases and related enzymes. Curr Opin Struct Biol 9:722–731 Seeger M, Timmis KN, Hofer B (1995) Degradation of chlorobiphenyls catalyzed by the bph-encoded biphenyl 2,3 dioxygenase and biphenyls 2,3-dihydrodiol 2,3-dehydrogenase of Pseudomonas sp. LB400. FEMS Microbiol Lett 133:259–264 Shingler V, Powlowski J, Marklund U (1992) Nucleotide sequence and functional analysis of the complete phenol/3,4-dimethylphenol catabolic pathway of Pseudomonas sp. strain CF600. J Bacteriol 174:711–724 Skiba A, Hecht V, Pieper DH (2002) Formation of protoanemonin from 2-chloro- cis, cis-muconate by the combined action of muconate cycloisomerase and muconolactone isomerase. J Bacteriol 184:5402–5409 Solyanikova IP, Malteva OV, Vollmer MD, Golovleva LA, Schlömann M (1995) Characterization of muconate and chloromuconate cycloisomerase from Rhodococcus erythropolis 1CP: indications for functionally convergent evolution among bacterial cycloisomerases. J Bacteriol 177:2821–2826 Teramoto M, Futamata H, Harayama S, Watanabe K (1999) Characterization of a high-affinity phenol hydroxylase from Comamonas testosteroni R5 by gene cloning, and expression in Pseudomonas aeruginosa PAO1c. Mol Gen Genet 262:552–558 Tropel D, Meyer C, Armengaud J, Jouanneau Y (2002) Ferredoxin-mediated reactivation of the chlorocatechol 2,3-dioxygenase from Pseudomonas putida GJ31. Arch Microbiol 177:345–351 Vaillancourt FH, Labbe G, Drouin NM, Fortin PD, Eltis LD (2002) The mechanism-based inactivation of 2,3-dihydroxybiphenyl 1,2-dioxygenase by catecholic substrates. J Biol Chem 277:2019–2027 Vallaeys T, Courde L, McGowan C, Wright AD, Fulthorpe RR (1999) Phylogenetic analyses indicate independent recruitment of diverse gene cassettes during assemblage of the 2,4-D catabolic pathway. FEMS Microbiol Ecol 28:373–382 van Berkel WJ, Kamerbeek NM, Fraaije MW (2006) Flavoprotein monooxygenases, a diverse class of oxidative biocatalysts. J Biotechnol 124:670–689 van der Meer JR, van Neerven ARW, de Vries EJ, de Vos WM, Zehnder AJB (1991) Cloning and characterization of plasmid-encoded genes for the degradation of 1,2-dichloro-, 1,4-dichloro-, and 1,2,4-trichlorobenzene of Pseudomonas sp. strain P51. J Bacteriol 173:6–15 Vollmer MD, Schlömann M (1995) Conversion of 2-chloro- cis, cis-muconate and its metabolites 2-chloro- and 5-chloromuconolactone by chloromuconate cycloisomerase of pJP4 and pAC27. J Bacteriol 177:2938–2941 Vollmer MK, Fischer P, Knackmuss H-J, Schlömann M (1994) Inability of muconate cycloisomerases to cause dehalogenation during conversion of 2-chloro- cis, cis-muconate. J Bacteriol 176:4366–4375 Westendorf A, Müller RH, Babel W (2003) Purification and characterisation of the enantiospecific dioxygenases from Delftia acidovorans MCI initiating the degradation of phenoxypropionate and phenoxyacetate herbicides. Acta Biotechnol 23:3–17 Worsey M, Williams PA (1975) Metabolism of toluene and xylenes by Pseudomonas putida (arvilla) mt-2: evidence for a new function of the TOL plasmid. J Bacteriol 124:7–13 Xu L, Resing K, Lawson SL, Babbitt PC, Copley SD (1999) Evidence that pcpA encodes 2,6-dichlorohydroquinone dioxygenase, the ring cleavage enzyme required for pentachlorophenol degradation in Sphingomonas chlorophenolica strain ATCC 39723. Biochemistry 38:7659–7669

30

D.H. Pieper et al.

Xun L, Webster C (2004) A monooxygenase catalyzes sequential dechlorinations of 2,4,6trichlorophenol by oxidative and hydrolytic reactions. J Biol Chem 279:6696–6700 Xun L, Topp E, Orser CS (1992a) Diverse substrate range of a Flavobacterium pentachlorophenol hydroxylase and reaction stoichiometries. J Bacteriol 174:2898–2902 Xun L, Topp E, Orser CS (1992b) Purification and characterization of a tetrachloro-p-hydroquinone reductive dehalogenase from a Flavobacterium sp. J Bacteriol 174:8003–8007 Zhuang ZH, Song F, Zhang WH, Taylor K, Archambault A, Dunaway-Mariano D, Dong J, Carey PR (2002) Kinetic, Raman, NMR, and site-directed mutagenesis studies of the Pseudomonas sp. strain CBS3 4-hydroxybenzoyl-CoA thioesterase active site. Biochemistry 41:11152–11160

Structure-Function Relationships and Engineering of Haloalkane Dehalogenases Piia Kokkonen, Tana Koudelakova, Radka Chaloupkova, Lukas Daniel, Zbynek Prokop, and Jiri Damborsky

Abstract

The structure-function relationships for haloalkane dehalogenases, one of the best characterized enzyme families involved in degradation of halogenated compounds, are described. A substantial amount of mechanistic and structural information is currently available on haloalkane dehalogenases, providing good theoretical framework for their modification by protein engineering. Examples of constructed mutants include variants with modified (i) activity and specificity, (ii) stability, and (ii) enantioselectivity. Many variants carried mutations in the tunnels connecting the buried active site with surrounding solvent, rather than in the active site itself. Mutagenesis of residues lining the protein tunnels represents attractive and a viable approach of protein engineering.

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Structure of HLDs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Catalytic Residues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Active Site and Access Tunnels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Function of HLDs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Catalytic Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Substrate Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Engineering of HLDs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Mutants with Modified Activity and Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 3 3 6 7 7 9 9 9

P. Kokkonen (*) • T. Koudelakova • R. Chaloupkova • L. Daniel • Z. Prokop • J. Damborsky Loschmidt Laboratories, Department of Experimental Biology and Research Centre for Toxic Compounds in the Environment RECETOX, Faculty of Science, Masaryk University, Brno, Czech Republic International Centre for Clinical Research, St. Anne’s University Hospital, Brno, Czech Republic e-mail: [email protected]; [email protected]; [email protected]; [email protected]; [email protected]; [email protected] # Springer International Publishing AG 2017 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils and Lipids,Handbook of Hydrocarbon and Lipid Microbiology , DOI 10.1007/978-3-319-39782-5_15-1

1

2

P. Kokkonen et al.

4.2 Mutants with Modified Stability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Mutants with Modified Enantioselectivity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

11 13 14 15

Introduction

Haloalkane dehalogenases (HLDs, EC 3.8.1.5) are enzymes cleaving a carbonhalogen bond in halogenated hydrocarbons (Scheme 1). The very first HLD was isolated from Xanthobacter autotrophicus GJ10 in 1985 (Keuning et al. 1985) and served as a model for carbon-halogen bond cleavage in halogenated aliphatic hydrocarbons. Since then, a number of newly isolated and biochemically characterized HLDs have grown to 24 enzymes (Koudelakova et al. 2013a; Fortova et al. 2013; Li and Shao 2014; Novak et al. 2014; Fung et al. 2015; Carlucci et al. 2016). HLDs have been isolated from bacteria colonizing contaminated environments (Keuning et al. 1985; Scholtz et al. 1987; Yokota et al. 1987; Janssen et al. 1989; Sallis et al. 1990; Nagata et al. 1997; Poelarends et al. 1998, 1999; Kumari et al. 2002; Sfetsas et al. 2009; Fung et al. 2015), but interestingly also from pathogenic organisms (Jesenska et al. 2000, 2002, 2005), an extremophile (Drienovska et al. 2012), and a eukaryote (Fortova et al. 2013). Phylogenetic analysis revealed that the HLD family can be divided into three subfamilies denoted HLD-I, HLD-II, and HLD-III, of which HLD-I and HLD-III are predicted to be the sister groups (Chovancova et al. 2007). A substantial amount of mechanistic and structural information is currently available on HLDs. The unique tertiary structures were determined by protein crystallography for DhlA, isolated from X. autotrophicus GJ10 (Franken et al. 1991), DhaA from Rhodococcus sp. TDTM0003 (Newman et al. 1999), LinB from Sphingobium japonicum UT26 (formerly Sphingomonas paucimobilis UT26) (Marek et al. 2000), DmbA from Mycobacterium tuberculosis H37Rv (Mazumdar et al. 2008), DbjA from Bradyrhizobium diazoefficiens USDA110 (formerly Bradyrhizobium japonicum USDA110) (Prokop et al. 2010), DppA from Plesiocystis pacifica SIR-1 (Hesseler et al. 2011), DmmA from the metagenomic DNA of a marine microbial consortium (Gehret et al. 2012), DatA from Agrobacterium fabrum C58 (formerly Agrobacterium tumefaciens C58) (Guan et al. 2014), DbeA from Bradyrhizobium

Scheme 1 General scheme of the reaction mechanism of HLDs. Alkyl-enzyme intermediate is formed in the first reaction step by nucleophilic attack of carboxylate oxygen of an aspartate group on the carbon atom of the substrate. This intermediate is in the second reaction step hydrolyzed by an activated water molecule, yielding a halide ion, a proton, and an alcohol as the products. Enz – enzyme

Structure-Function Relationships and Engineering of Haloalkane. . .

3

elkanii USDA94 (Chaloupkova et al. 2014), HanR from a Rhodobacteraceae bacterium (Novak et al. 2014), DmrA from Mycobacterium rhodesiae JS60 (Fung et al. 2015), and DccA from Caulobacter crescentus (Carlucci et al. 2016). Moreover, a number of crystal structures of HLDs with ligands and of different HLDs variants are available in the Protein Data Bank. The structure and reaction mechanism of HLDs has been studied in detail by protein crystallography, site-directed mutagenesis, enzyme kinetics, and molecular modeling (Fig. 1). The number of practical applications employing HLDs is increasing with growing knowledge of their properties and structure-function relationships. HLDs can find their use in the bioremediation of environmental pollutants (Stucki and Thuer 1995), biosensing of toxic chemicals (Campbell et al. 2006; Bidmanova et al. 2016), industrial biocatalysis (Swanson 1999; Janssen 2007; Prokop et al. 2009, 2010; Westerbeek et al. 2011a, b; van Leeuwen et al. 2012), decontamination of warfare agents (Prokop et al. in press, 2006, 2011), as well as cell imaging, protein analysis, and directed evolution using phage display (Los et al. 2008; Ohana et al. 2009; Koudelakova et al. 2013a; Delespaul et al. 2015).

2

Structure of HLDs

HLDs structurally belong to the α/β-hydrolase superfamily (Ollis et al. 1992; Nardini and Dijkstra 1999). The proteins in this superfamily do not possess obvious sequence similarity, even though they have diverged from a common ancestor. The three-dimensional structure of HLDs is composed of two domains: (i) the strictly conserved α/β-hydrolase main domain and (ii) the helical cap domain, variable in terms of number and the arrangement of secondary elements (Fig. 2). The α/β-hydrolase fold is mostly made up of an eight-stranded parallel β-sheet which is flanked by α-helices (Verschueren et al. 1993c). The cap domain consists of several helices connected by loops and is inserted to the main domain after the β-strand 6. Most structural differences between HLDs are found in the cap domain.

2.1

Catalytic Residues

HLDs possess an essential catalytic pentad composed of a catalytic triad and a pair of halide-stabilizing residues. The catalytic triad comprises a nucleophile, a catalytic base, and a catalytic acid (Fig. 2). The composition of the catalytic pentad varies among subfamilies: Asp-His-Asp + Trp-Trp in subfamily HLD-I, Asp-His-Glu + AsnTrp/Tyr in subfamily HLD-II, and Asp-His-Asp + Asn-Trp in subfamily HLD-III (Chovancova et al. 2007; Hasan et al. 2011; Guan et al. 2014). The nucleophile is located on a very sharp turn, known as the nucleophile elbow, where it can be easily reached by the substrate and the catalytic water molecule. The geometry of the nucleophile elbow also contributes the formation of the oxyanion-binding site, which is needed to stabilize the negatively charged transition state that occurs during hydrolysis (Verschueren et al. 1993c). Catalysis proceeds by the nucleophilic attack of

Fig. 1 Properties of characterized HLDs. Summary originates from a number of characterization studies (Keuning et al. 1985; Yokota et al. 1987; Janssen et al. 1988; Sallis et al. 1990; Pieters et al. 2001; Jesenska et al. 2002, 2005, 2009; Sato et al. 2005; Pavlova et al. 2007; Prokop et al. 2010; Hasan et al. 2011; Koudelakova et al. 2011; Westerbeek et al. 2011b; Hesseler et al. 2011; Gehret et al. 2012; Drienovska et al. 2012; Li and Shao 2014; Novak et al. 2014; Chaloupkova et al. 2014; Fung et al. 2015; Carlucci et al. 2016). The isoelectric point (pI) and molecular weight (MW) were predicted by Expasy server (Gasteiger et al. 2003). The detail description of thermal denaturation circular dichroism measurements and used recalculation is given in the review by Koudelakova et al. (2013a). Symbols: qualitative measures of activity/enantioselectivity:
– not active/selective, * – active/selective with a few substrates, ** – active/selective with several substrates, and *** – active/selective with a number of substrates; 1 – polymer, a – recalculated, b – dependent on conditions, c – predicted. Overall activity was estimated based on a multivariate analysis of specific activities towards 30 selected substrates (Koudelakova et al. 2011). The structure and reaction mechanism of HLDs (Scheme 1) has been studied by using protein crystallography (Verschueren et al. 1993a, b, c; Ridder et al. 1999; Newman et al. 1999; Marek et al. 2000; Oakley et al. 2002, 2004; Streltsov et al. 2003; Liu et al. 2007; Mazumdar et al. 2008; Hesseler et al. 2011; Guan et al. 2014; Carlucci et al. 2016; Fung et al. 2015; Novak et al. 2014; Chaloupkova et al. 2014; Prokop et al. 2010), site-directed mutagenesis (Pries et al. 1995a; Schanstra et al. 1997; Krooshof et al. 1997; Holloway et al. 1998; Hynkova et al. 1999; Schindler et al. 1999; Bohac et al. 2002; Chaloupkova et al. 2003, 2014; Pavlova et al. 2007; Prokop et al. 2010; Hasan et al. 2013;

4 P. Kokkonen et al.

Structure-Function Relationships and Engineering of Haloalkane. . .

5

Fig. 2 Molecular topology (a) and tertiary structure (b) of HLDs. α/β-hydrolase fold domain (white) and the specificity-determining cap domain (black) are distinguished. A nucleophile, a base and the first halide-stabilizing residue are conserved ( filled symbols), whereas the catalytic acid and the second halide-stabilizing residue are variable among HLDs (empty symbols)

the carboxylate oxygen of the nucleophile on the carbon atom of the substrate, yielding displacement of a halide, and continues with the formation of a covalent alkyl-enzyme intermediate (Scheme 1). The alkyl-enzyme intermediate is subsequently hydrolyzed by a water molecule that is activated by the catalytic base. A catalytic acid stabilizes the charge developed on the imidazole ring of the catalytic base during the hydrolytic part of the reaction.

2.2

Active Site and Access Tunnels

The active site of HLDs is located at the interface of the main domain and the cap domain (Fig. 2). The only polar groups localized in the active sites of HLDs are the residues of the catalytic triad. The active sites of HLDs differ in their size and accessibility to the solvent (Fig. 3). The mostly hydrophobic active sites are connected

ä Fig. 1 (continued) Brezovsky et al. 2016), enzyme kinetics (Schanstra and Janssen 1996; Schanstra et al. 1996a; Bosma et al. 2003; Prokop et al. 2003; Kaushik et al. 2016), and molecular modeling (Damborsky et al. 1997, 1998, 2003; Maulitz et al. 1997; Lightstone et al. 1998; Lau et al. 2000; Kmunicek et al. 2001, 2003, 2005; Shurki et al. 2002; Otyepka and Damborsky 2002; Bohac et al. 2002; Devi-Kesavan and Gao 2003; Hur et al. 2003; Kahn and Bruice 2003; Silberstein et al. 2003; Soriano et al. 2003, 2005; Nam et al. 2004; Olsson and Warshel 2004; Banas et al. 2006; Negri et al. 2007; Otyepka et al. 2008; Daniel et al. 2015)

6

P. Kokkonen et al.

Fig. 3 Anatomy of the active sites and tunnels in HLDs. The position of buried active site and two tunnels connecting the active site with surrounding environment is schematized in (a), with denoted (1) an active site, (2) a main tunnel, and (3) a slot. A surface of the active site and the tunnels is represented by wire in DhlA (b), and DbjA (c)

to the surrounding solvent by permanent or ligand-induced access tunnels which control the access of substrates and water to the active site and the egress of products from the active site (Marek et al. 2000; Otyepka and Damborsky 2002; Negri et al. 2007; Brezovsky et al. 2016). The size, shape, physicochemical properties, and dynamics of the tunnels vary between different HLDs. For example, the active site of the crystal structure of DhlA displays narrow tunnels, whereas the tunnels in the crystal structure of DbjA are larger and solvated (Fig. 3) (Verschueren et al. 1993c; Prokop et al. 2010). However, these static structures offer only a glimpse to the dynamic nature of the active site and the tunnels of HLDs. During molecular dynamics simulations the tunnels change their shape and occasionally it is possible to observe tunnels which are not visible in the crystal structures (Chovancova et al. 2012). By modifying the existing tunnels or by inducing a de novo transport tunnel it is possible

Structure-Function Relationships and Engineering of Haloalkane. . .

7

to affect the activity and substrate specificity of HLDs (Pavlova et al. 2009; Koudelakova et al. 2013a; Nagata et al. 2015; Brezovsky et al. 2016).

3

Function of HLDs

3.1

Catalytic Activity

HLDs exhibit catalytic efficiencies ranging from 104 to 105 M1 s1 with their best substrates (Koudelakova et al. 2013a). The kinetic mechanism of the catalyzed reaction, studied for three HLDs (Table 1), consists of four basic steps: (i) binding of the substrate, (ii) cleavage of the carbon-halogen bond, (iii) hydrolysis of the alkyl-enzyme intermediate, and (iv) the release of the products (Scheme 2) (Schanstra and Janssen 1996; Bosma et al. 2003; Prokop et al. 2003). The binding of the substrate and the cleavage of the carbon-halogen bond are fast steps for all studied enzymes, while the rate-limiting step differs in all cases. Hydrolysis of the alkyl-enzyme intermediate is limiting for LinB, the release of halide and alcohol limits reaction of DhlA and DhaA, respectively (Table 1). Moreover, a single HLD can show different rate-limiting steps for individual substrates (Pavlova et al. 2009). The varying rate-limiting step for close relative members of same enzyme family and even for different enzyme-substrate pairs of the same enzyme demonstrates that Table 1 Steady-state and transient kinetic parameters of selected HLDs

Enzyme DhlA wta DhlA wta DhlA V226Ab DhlA V226Ab DhlA F172Wc DhlA F172Wc DhlA D260N + N148Ed DhaA wte LinB wtf LinB wtf LinB wtf

Substrate DBE DCE DBE DCE DBE DCE DBE

Steady-state kinetics kcat Km (s1) (mM) 3 0.01 3 0.5 8 0.03 4 2 6 0.03 3 5 0.4 0.4

Transient kinetics Ks (k1/k1) k2 (mM) (s1) >0.03 >100 2 50 0,1 60 6 10 0,1 30 10 5 0,7 1

k3 (s1) 10 10 10 9 9 10 0.8

k4 (s1) 4 8 40 50 80 >80 >10

DBP CH BCH CCH

3 3 2 0.1

500 > 500

20 3 3 0.1

4 >1000 >1000 >1000

0.01 0.02 0.02 0.2

300 120 >200 >40

determined at pH 8.2 and 30  C (Schanstra et al. 1996a); bdetermined at pH 8.2 and 30  C (Schanstra et al. 1997); cdetermined at pH 8.2 and 30  C (Schanstra et al. 1996b); ddetermined at pH 8.2 and 30  C (Krooshof et al. 1998); edetermined at pH 8.2 and 30  C (Krooshof et al. 1997); f determined at pH 9.4 and 30  C (Bosma et al. 2003); Abreviations: 1,2-dichloroethane (DCE), 1,2-dibromoethane (DBE), 1,3-dibromopropane (DBP), 1-chlorohexane (CH), bromocyclohexane (BCH), chlorocyclohexane (CCH). The rate-determining step is shadowed a

8

E + RX

P. Kokkonen et al. k1 k -1

E.RX

k2 k-2

E-R. X

k3 k -3

E.X-.ROH

k4 k -4

E + X-+ ROH

Scheme 2 Kinetic mechanism of HLDs. E – enzyme, RX – substrate (halogenated alkane), E.RX – enzyme-substrate complex, E-R.X – alkyl-enzyme intermediate, E.X.ROH – enzyme-product complex, X – halide product, ROH – alcohol product. kx – kinetic constant of an individual catalytic step

extrapolation of this important catalytic property from one reaction to another can be misleading.

3.2

Substrate Specificity

HLDs are broad specificity enzymes, which can convert more than a hundred chlorinated, brominated, and iodinated chemicals: haloalkanes, haloalkenes, haloalcohols, halohydrins, haloethers, haloesters, haloacetamides, haloacetonitriles, and cyclohaloalkanes (Damborsky et al. 2001). Principal component analysis of substrate specificity profiles with 30 halogenated substrates revealed the presence of different substrate specificity groups: a group of catalytically robust enzymes which can convert recalcitrant chlorinated substrates, a group of enzymes which prefer brominated and iodinated substrates, and two groups which consist of a single HLD with a unique substrate specificity profile (Koudelakova et al. 2011). The following compounds were recognized as “universal” substrates: 1-bromobutane, 1-iodopropane, 1-iodobutane, 1,2-dibromoethane, and 4-bromobutanenitrile. The identified substrate specificity groups are not in agreement with the phylogenetic classification of HLDs, so the substrate specificity cannot be predicted only from the sequence (Chovancova et al. 2007; Koudelakova et al. 2011). The substrate specificity of HLDs arises from the structure and properties of the cap domain, the active site, and the access tunnels (Koudelakova et al. 2011). These structural features of the cap domain can be used in computational studies to find new substrates for HLDs (Daniel et al. 2015).

4

Engineering of HLDs

The initial protein engineering studies on HLDs were focused on site-directed mutagenesis of their potential catalytic residues selected rationally based on X-ray structures or homology models (Pries et al. 1994a, 1995b; Kennes et al. 1995; Krooshof et al. 1997; Hynkova et al. 1999; Nagata et al. 1999; Newman et al. 1999; Bohac et al. 2002; Sato et al. 2005; Pavlova et al. 2007). Follow-up protein engineering studies examined the possibility to improve the catalytic efficiency or stability of HLDs.

Structure-Function Relationships and Engineering of Haloalkane. . .

9

Fig. 4 Engineering of activity and specificity of DhaA by focused directed evolution (Pavlova et al. 2009). Positions with introduced mutations are indicated as the white spheres and the side chains of mutated residues in sticks. Some of the mutations changed the shape of access tunnels, which are shown in surface representation. The location of the active site is indicated by a black star

4.1

Mutants with Modified Activity and Specificity

Mutants with modified activity and specificity can be prepared by modification of the access tunnels (Chaloupkova et al. 2003; Banas et al. 2006) or by exploiting unique structural features of individual HLDs, such as the second halide-binding site of DbeA (Chaloupkova et al. 2014) or the different cap domain of DhlA (Pries et al. 1994b; Pikkemaat and Janssen 2002) (Fig. 4). DhlA from HLD-I subfamily is unique among known HLDs by its activity towards 1,2-dichloroethane (Koudelakova et al. 2011). To reveal determinants of such an activity, Pries et al. expressed DhlA in a strain of Pseudomonas that grows on long-chain (C6) alcohols and isolated 12 different mutants that utilized 1-chlorohexane after 4 weeks of a batch cultivation in a medium with different ratio of 1-chlorobutane and 1-chlorohexane (Pries et al. 1994b). Selected variants contained following mutations: ∇145–154, ∇152–153, Δ164–174, P168S, D170H, and ∇172–174, which were mostly located in the N-terminal part of the cap domain. Only the mutant Δ164–174 affected directly the active site by deleting its residues F164 and F172. The N-terminal part of the cap domain was further modified in vitro by randomly generated repeats and deletions in a follow-up study (Pikkemaat and Janssen 2002). Isolated mutants contained a repeat in the N-terminal part of the cap domain and showed up to threefold improved activity towards 1,2-dibromoethane. Construction of a dehalogenase enzyme with improved conversion of 1,2,3-trichloropropane (TCP) is an essential step towards engineering a TCP-degrading strain carrying a synthetic biodegradation pathway (Kurumbang et al. 2014). The first

10

P. Kokkonen et al.

improved variant of DhaA (C176F) was obtained by error prone PCR and showed a fourfold improvement in activity with TCP (Gray et al. 2001). The combination of DNA shuffling and error-prone PCR yielded a DhaA double-mutant (C176Y + Y273F), which was 3.5-fold more active towards TCP than the wild type enzyme (Bosma et al. 2002). Twenty-five unique DhaA variants with higher activities towards TCP were obtained by focused directed evolution of residues influencing the release of the product 2,3-dichloropropane-1-ol (Pavlova et al. 2009). Random acceleration molecular dynamics (Lüdemann et al. 2000), simulating the release of the product from the enzyme active site, was applied for the selection of the “hot spot” residues for saturation mutagenesis (Pavlova et al. 2009). The best mutant from the activity screening carried five mutations (I135F + C176Y + V245F + L246I + Y273F) and exhibited 26-fold improved catalytic efficiency towards TCP. Interestingly, described mutagenesis of DhaA targeted the access tunnels rather than the active site (Gray et al. 2001; Bosma et al. 2002; Pavlova et al. 2009). Improved variants had bulky aromatic residues introduced into their access tunnels leading to the occluded active site. The rate-limiting step of TCP conversion was shifted from carbon-halogen bond cleavage to the release of the reaction products in the best DhaA31 variant (Pavlova et al. 2009). The importance of transport tunnels for catalysis of HLDs was highlighted also in our recent study (Brezovsky et al. 2016). The main tunnel of LinB was closed by a disulfide bridge (D147C + L177C) or bulky substitution L177W, and a de novo transport tunnel was created based on computational design using focused directed evolution. The catalytic activity of mutants with blocked tunnels was reduced 17 times and showed large substrate inhibition. Opening of the novel access tunnel (W140A + F143L + L177W + I211L) resulted in the most proficient HLD reported to date.

4.2

Mutants with Modified Stability

Due to the limited knowledge of stability determinants of HLDs in 2001, Gray et al. attempted to improve stability of DhaA by a directed evolution technique called the gene site saturation mutagenesis (Gray et al. 2001; Kretz et al. 2004). The aim was to develop an efficient biocatalytic process for the conversion of the side-product TCP during production of epichlorohydrin (Swanson 1999; Gray et al. 2001). All possible single-site mutants of DhaA were screened using a high-throughput activity assay at elevated temperatures, and eight beneficial single-point mutations were discovered (Gray et al. 2001). A combination of these mutations yielded a DhaA variant (D78G + F80S + T148L + G171Q + I209L + N227T + W240Y + P291A) with a 30,000-fold longer half-life at 55  C, and increased melting temperature by 8  C. A systematic set of derived DhaA mutants showed that mutations targeting as few as four residues lining the access tunnel of DhaA (T148L + G171V + A172V + C176F) could extend the half-life 4000-fold in 40% (v/v) dimethyl sulfoxide at 37  C, and increase its melting temperature by 19  C (Koudelakova et al. 2013b). The poor

Structure-Function Relationships and Engineering of Haloalkane. . .

11

overall catalytic activity of the highly thermostable mutant in a buffer was improved by the substitution F176G (Liskova et al. 2015). Two successful computational designs of thermostable HLDs have been described (Floor et al. 2014; Bednar et al. 2015). Floor et al. used a fast in silico screening tool FRESCO for design of 136 single-point mutants and 13 new disulfide bonds in LinB (Floor et al. 2014). Based on results from functional characterization of all expressed mutants, ten stabilizing mutations and one disulfide bond were introduced into the combined LinB mutant (E15T + A53L A81K + D166K + F169V + E192K + A197P + G229Q + A247F + D255A; A5C + A185C) which exhibited 200-fold longer half-life at 60  C and increase in melting temperature by 23  C. The experimental effort can be further reduced by a recently developed computational method FireProt (Bednar et al. 2015). This method searches for stabilizing mutations via energy calculations and a phylogenetic back-to-consensus analysis. FireProt combines both calculations with smart filtering to design highly stabilized multiple-point mutants. Five multiple mutants of DhaA were designed, constructed, and characterized. The combined 11-fold mutant (E20S + F80R + C128F + T148L + A155P + A172I + C176F + D198W + V219W + C262L + D266F) showed a significantly improved half-life (>7 days) at 60  C and increased thermostability by 24  C. Studies described in previous paragraphs showed that structural changes leading to improved thermostability of an enzyme were also responsible for its improved tolerance to organic cosolvents. Both close and distant mutations to the active site could be beneficial (Fig. 5). Highly stabilizing residues close to the active site were found in the access tunnel (Koudelakova et al. 2013b). They narrowed the access tunnel, sealed the active site, and thus improved intramolecular hydrophobic packing of the enzyme. The thermostability-activity trade-off could be addressed by directed evolution of the enzyme of interest, i.e., by saturation mutagenesis of the access tunnel residues followed by activity screening (Liskova et al. 2015). Besides improving hydrophobic packing, the stabilizing mutations distant from the active site included formation of a disulfide bridge, formation of hydrogen bonds, removal and introduction of charges on the surface, replacing a glycine, and introduction of a proline in a loop (Gray et al. 2001; Floor et al. 2014; Bednar et al. 2015). Further engineering of stability of HLDs will be straightforward thanks to computational tools FireProt and HotSpot Wizard (Bednar et al. 2015; Bendl et al. 2016).

4.3

Mutants with Modified Enantioselectivity

Based on findings that DhlA and DhaA do not discriminate enantiomers of dihalogenated alkanes, terminally halogenated esters and prochiral halogenated propanes (including TCP), the development of enantioselective dehalogenases for the use in industrial biocatalysis has been defined as one of the major challenges in the engineering of HLDs (Pieters et al. 2001; Janssen 2004). Prokop et al. assayed DhaA, LinB, and DbjA for their enantioselective conversion of α-brominated esters and β-brominated alkanes to chiral alcohols, all three

12

P. Kokkonen et al.

Fig. 5 Engineering of stability of DhaA by directed evolution (Gray et al. 2001; Koudelakova et al. 2013b; Liskova et al. 2015). The representative structures of the main tunnels in the cap domains prepared by (a) mutagenesis of tunnel residues T148 L + G171Q+ A172V + C176F (Koudelakova et al. 2013b) and (b) saturation mutagenesis of tunnel residues A172 + C176 (Liskova et al. 2015). The introduced residues are shown as sticks. The tunnels in gray spheres (left) and the tunnel profiles (in right) are calculated by CAVER 3.01 (Chovancova et al. 2012)

proteins possessed high enantioselectivity (E-value > 200) with α -brominated esters (Prokop et al. 2010). DbjA additionally showed high enantioselectivity towards 2-bromopentane (E-value 132 at 37  C). Structural analysis revealed that DbjA has a unique surface loop at the interface of the main and the cap domain. The deletion of this loop (140HTEVAEE146) significantly lowered enantioselectivity of the enzyme towards 2-bromopentane (E-value 33 at 37  C). The conformation of nearby His139 changed in the DbjA mutant and thus altered the shape and size of the active site. Additional substitution H139A in this conformation-changing histidine made the enzyme enantioselective again (E-value 100). The follow-up study with DhaA implied that structural features distant from the active site can affect the dynamics and hydration of the active site and control the enantioselectivity (Sykora et al. 2014). The introduction of the unique surface loop and the active site of DbjA into the scaffold of DhaA did not improve

Structure-Function Relationships and Engineering of Haloalkane. . .

13

Fig. 6 Engineering of enantioselectivity of DbjA by rational design (Prokop et al. 2010). Wild type DbjA (a), and DbjA Δ140–146 + H139A (b) are shown. The region carrying deletion in the surface loop is shown in ribbon. The “gate-keeping” residue His/Ala139 and the catalytic triad are shown in sticks. The mutants were designed based on sequence/structure comparisons and constructed by site-directed mutagenesis

enantioselectivity of the constructed DhaA variant towards 2-bromopentane. On the other hand, pair-wise site-saturation mutagenesis of noncatalytic first-shell active site residues of DhaA yielded enantiocomplementary mutants, which convert prochiral TCP into highly enantioenriched (R)- or (S)-2,3-dichloropropan-1-ol (van Leeuwen et al. 2012). Variants containing 13 and 17 mutations enabled preparation of (R)-epichlorohydrin (90% ee) and (S)-epichlorohydrin (97% ee), respectively. These studies showed that it is challenging to engineer enantioselectivity rationally (Fig. 6). Factors such as protein dynamics and hydration could be better modified by focused directed evolution approach, because the computational methods cannot predict the effects of mutations on catalysis. The growing number of known enantioselective HLDs will help to understand the structural basis of enantioselectivity in these enzymes (Hasan et al. 2011; Drienovska et al. 2012).

5

Research Needs

As HLDs have a simple reaction mechanism, broad range of substrate specificities, and are relatively stable and easy to handle, they have become one of the best characterized enzyme families. Characterization of new family members and variants has led to new knowledge about structure-function relationships and the evolution of HLDs. The newly isolated enzymes, native or engineered, display great potential for practical applications which often require optimized properties: (i) high enantioselectivity with substrates that can be converted to valuable products by

14

P. Kokkonen et al.

biocatalysis, (ii) enhanced resistance to organic solvents for decontamination purposes, (iii) elevated activity or modified substrates specificity for bioremediation, (iv) broadened pH range for biosensing, and (v) increased thermostability and longterm stability for various applications. The prediction of the effect of mutations on the function and stability of the enzymes remains to be challenging despite recent improvements in computational methods (Kuipers et al. 2010; Wijma et al. 2014; Bednar et al. 2015; Bendl et al. 2016; Goldenzweig et al. 2016). Developing more precise methods would simultaneously reduce the experimental burden and pave the way for made-to-order enzyme variants for industrial needs. Some of the applications like neutralization of sulfur mustard, enantioselective conversion of halogenated alkanes, or degradation of toxic environmental pollutant 1,2,3-trichloropropane are clearly limited by poor catalytic properties of native enzymes or insufficient stabilities in organic cosolvents. Development of broadly applicable biosensors would benefit from HLDs with altered pH-range. Continuation of protein engineering efforts and isolation of new wild type HLDs from extremophiles should provide catalysts fitting the practical needs. Continuous collection of detailed kinetic and mechanistic information provides a critical knowledge base for rational protein engineering strategies. Transient kinetics and complete kinetic characterization would be beneficial for additional natural variants since three up-to-date fully characterized dehalogenases all provided unique behavior and difference in their rate-limiting steps. We also suggest to apply transient kinetics systematically for characterization of engineered variants to (i) dissect and quantify an impact of mutations on the individual catalytic steps, (ii) identify changes of rate limitation upon mutagenesis, and (iii) exactly specify directions for rational design. Beyond the basic kinetic pathway described in Scheme 2, additional kinetic phenomena, e.g., substrate or product inhibition, cooperation, and dynamic conformational changes, were identified to be critical for catalytic performance of various HLDs. It would be valuable to provide better understanding of these widespread phenomena by using transient kinetics and/or single molecule techniques for advanced rational engineering of these catalysts. The greatest challenge in the research of HLDs is the identification of their biological role. The genes coding for HLDs are widely distributed among various species, including strictly pathogenic microorganisms, such as Mycobacterium tuberculosis and Mycobacterium bovis. It is currently unclear, which type of chemical reaction and which substrates are converted by these isolated and biochemically characterized dehalogenases. At the same time, dehalogenating activity has not yet been confirmed experimentally for putative HLDs, while for many biochemically characterized HLDs endogenous substrates are not known. For proteins from subfamily HLD-III, even the tertiary structure is not available due to poor crystallizability of large protein multimers. More than 30 years after description of the first HLD, this enzyme family offers some hidden secrets. Acknowledgments Financial support is gratefully acknowledged from the Ministry of Education, Youth, and Sports of the Czech Republic (LO1214, LM2015051, LM2015047, LM2015055, LQ1605), The Czech Grant Agency (GA16-06096S, GA16-07965S, GA16-24223S), and the

Structure-Function Relationships and Engineering of Haloalkane. . .

15

European Union H2020 EXCELERATE (676559). Access to the METACentrum supercomputing facilities is highly appreciated (LM2015042 and LM2015085).

References Banas P, Otyepka M, Jerabek P, Petrek M, Damborsky J (2006) Mechanism of enhanced conversion of 1,2,3-trichloropropane by mutant haloalkane dehalogenase revealed by molecular modeling. J Comput Aided Mol Des 20:375–383 Bednar D, Beerens K, Sebestova E, Bendl J, Khare S, Chaloupkova R, Prokop Z, Brezovsky J, Baker D, Damborsky J (2015) FireProt: energy- and evolution-based computational design of thermostable multiple-point mutants. PLoS Comput Biol 11:e1004556 Bendl J, Stourac J, Sebestova E, Vavra O, Musil M, Brezovsky J, Damborsky J (2016) HotSpot wizard 2.0: automated design of site-specific mutations and smart libraries in protein engineering. Nucleic Acids Res 44:W479–W487 Bidmanova S, Steiner M-S, Stepan M, Vymazalova K, Gruber MA, Duerkop A, Damborsky J, Prokop Z, Wolfbeis OS (2016) Enzyme-based test strips for visual or photographic detection and quantitation of gaseous sulfur mustard. Anal Chem 88:6044–6049 Bohac M, Nagata Y, Prokop Z, Prokop M, Monincova M, Tsuda M, Koca J, Damborsky J (2002) Halide-stabilizing residues of haloalkane dehalogenases studied by quantum mechanic calculations and site-directed mutagenesis. Biochemistry 41:14272–14280 Bosma T, Damborsky J, Stucki G, Janssen D (2002) Biodegradation of 1,2,3-trichloropropane through directed evolution and heterologous expression of a haloalkane dehalogenase gene. Appl Environ Microbiol 68:3582–3587 Bosma T, Pikkemaat MG, Kingma J, Dijk J, Janssen DB (2003) Steady-state and pre-steady-state kinetic analysis of halopropane conversion by a rhodococcus haloalkane dehalogenase. Biochemistry 42:8047–8053 Brezovsky J, Babkova P, Degtjarik O, Fortova A, Gora A, Iermak I, Rezacova P, Dvorak P, Smatanova IK, Prokop Z, Chaloupkova R, Damborsky J (2016) Engineering a de novo transport tunnel. ACS Catal 6:7597–7610 Campbell D, Muller C, Reardon K (2006) Development of a fiber optic enzymatic biosensor for 1,2-dichloroethane. Biotechnol Lett 28:883–887 Carlucci L, Zhou E, Malashkevich VN, Almo SC, Mundorff EC (2016) Biochemical characterization of two haloalkane dehalogenases: DccA from Caulobacter crescentus and DsaA from Saccharomonospora azurea. Protein Sci 25:877–886 Chaloupkova R, Sykorova J, Prokop Z, Jesenska A, Monincova M, Pavlova M, Tsuda M, Nagata Y, Damborsky J (2003) Modification of activity and specificity of haloalkane dehalogenase from Sphingomonas paucimobilis UT26 by engineering of its entrance tunnel. J Biol Chem 278:52622–52628 Chaloupkova R, Prudnikova T, Rezacova P, Prokop Z, Koudelakova T, Daniel L, Brezovsky J, Ikeda-Ohtsubo W, Sato Y, Kuty M, Nagata Y, Smatanova IK, Damborsky J (2014) Structural and functional analysis of a novel haloalkane dehalogenase with two halide-binding sites. Acta Crystallogr D Biol Crystallogr 70:1884–1897 Chovancova E, Kosinski J, Bujnicki JM, Damborsky J (2007) Phylogenetic analysis of haloalkane dehalogenases. Proteins 67:305–316 Chovancova E, Pavelka A, Benes P, Strnad O, Brezovsky J, Kozlikova B, Gora A, Sustr V, Klvana M, Medek P, Biedermannova L, Sochor J, Damborsky J (2012) CAVER 3.0: a tool for the analysis of transport pathways in dynamic protein structures. PLoS Comput Biol 8: e1002708 Damborsky J, Kuty M, Nemec M, Koca J (1997) A molecular modeling study of the catalytic mechanism of haloalkane dehalogenase: 1. Quantum chemical study of the first reaction step. J Chem Inf Comput Sci 37:562–568

16

P. Kokkonen et al.

Damborsky J, Bohac M, Prokop M, Kuty M, Koca J (1998) Computational site-directed mutagenesis of haloalkane dehalogenase in position 172. Protein Eng 11:901–907 Damborsky J, Rorije E, Jesenska A, Nagata Y, Klopman G, Peijnenburg WJ (2001) Structurespecificity relationships for haloalkane dehalogenases. Environ Toxicol Chem SETAC 20:2681–2689 Damborsky J, Kmunicek J, Jedlicka T, Luengo S, Gago F, Ortiz AR, Wade RC (2003) Rational re-design of haloalkane dehalogenases guided by comparative binding energy analysis. In: Svendsen A (ed) Enzyme functionality: design, engineering and screening. Marcel Dekker, New York, pp 79–96 Daniel L, Buryska T, Prokop Z, Damborsky J, Brezovsky J (2015) Mechanism-based discovery of novel substrates of haloalkane dehalogenases using in silico screening. J Chem Inf Model 55:54–62 Delespaul W, Peeters Y, Herdewijn P, Robben J (2015) A novel helper phage for HaloTag-mediated co-display of enzyme and substrate on phage. Biochem Biophys Res Commun 460:245–249 Devi-Kesavan LS, Gao J (2003) Combined QM/MM study of the mechanism and kinetic isotope effect of the nucleophilic substitution reaction in haloalkane dehalogenase. J Am Chem Soc 125:1532–1540 Drienovska I, Chovancova E, Koudelakova T, Damborsky J, Chaloupkova R (2012) Biochemical characterization of a novel haloalkane dehalogenase from a cold-adapted bacterium. Appl Environ Microbiol 78:4995–4998 Floor RJ, Wijma HJ, Colpa DI, Ramos-Silva A, Jekel PA, Szymanski W, Feringa BL, Marrink SJ, Janssen DB (2014) Computational library design for increasing haloalkane dehalogenase stability. Chembiochem 15:1659–1671 Fortova A, Sebestova E, Stepankova V, Koudelakova T, Palkova L, Damborsky J, Chaloupkova R (2013) DspA from Strongylocentrotus purpuratus: the first biochemically characterized haloalkane dehalogenase of non-microbial origin. Biochimie 95:2091–2096 Franken SM, Rozeboom HJ, Kalk KH, Dijkstra BW (1991) Crystal structure of haloalkane dehalogenase: an enzyme to detoxify halogenated alkanes. EMBO J 10:1297–1302 Fung HKH, Gadd MS, Drury TA, Cheung S, Guss JM, Coleman NV, Matthews JM (2015) Biochemical and biophysical characterisation of haloalkane dehalogenases DmrA and DmrB in Mycobacterium strain JS60 and their role in growth on haloalkanes. Mol Microbiol 97:439–453 Gasteiger E, Gattiker A, Hoogland C, Ivanyi I, Appel RD, Bairoch A (2003) ExPASy: the proteomics server for in-depth protein knowledge and analysis. Nucleic Acids Res 31:3784–3788 Gehret JJ, Gu L, Geders TW, Brown WC, Gerwick L, Gerwick WH, Sherman DH, Smith JL (2012) Structure and activity of DmmA, a marine haloalkane dehalogenase. Protein Sci 21:239–248 Goldenzweig A, Goldsmith M, Hill SE, Gertman O, Laurino P, Ashani Y, Dym O, Unger T, Albeck S, Prilusky J, Lieberman RL, Aharoni A, Silman I, Sussman JL, Tawfik DS, Fleishman SJ (2016) Automated structure- and sequence-based design of proteins for high bacterial expression and stability. Mol Cell 63:337–346 Gray K, Richardson T, Kretz K, Short J, Bartnek F, Knowles R, Kan L, Swanson P, Robertson D (2001) Rapid evolution of reversible denaturation and elevated melting temperature in a microbial haloalkane dehalogenase. Adv Synth Catal 343:607–617 Guan L, Yabuki H, Okai M, Ohtsuka J, Tanokura M (2014) Crystal structure of the novel haloalkane dehalogenase DatA from Agrobacterium tumefaciens C58 reveals a special halide-stabilizing pair and enantioselectivity mechanism. Appl Microbiol Biotechnol 98:8573–8582 Hasan K, Fortova A, Koudelakova T, Chaloupkova R, Ishitsuka M, Nagata Y, Damborsky J, Prokop Z (2011) Biochemical characteristics of the novel haloalkane dehalogenase DatA, isolated from the plant pathogen Agrobacterium tumefaciens C58. Appl Environ Microbiol 77:1881–1884 Hasan K, Gora A, Brezovsky J, Chaloupkova R, Moskalikova H, Fortova A, Nagata Y, Damborsky J, Prokop Z (2013) The effect of a unique halide-stabilizing residue on the catalytic properties of haloalkane dehalogenase DatA from Agrobacterium tumefaciens C58. FEBS J 280:3149–3159

Structure-Function Relationships and Engineering of Haloalkane. . .

17

Hesseler M, Bogdanović X, Hidalgo A, Berenguer J, Palm GJ, Hinrichs W, Bornscheuer UT (2011) Cloning, functional expression, biochemical characterization, and structural analysis of a haloalkane dehalogenase from Plesiocystis pacifica SIR-1. Appl Microbiol Biotechnol 91:1049–1060 Holloway P, Knoke KL, Trevors JT, Lee H (1998) Alteration of the substrate range of haloalkane dehalogenase by site-directed mutagenesis. Biotechnol Bioeng 59:520–523 Hur S, Kahn K, Bruice TC (2003) Comparison of formation of reactive conformers for the SN2 displacements by CH3CO2- in water and by Asp124-CO2- in a haloalkane dehalogenase. Proc Natl Acad Sci USA 100:2215–2219 Hynkova K, Nagata Y, Takagi M, Damborsky J (1999) Identification of the catalytic triad in the haloalkane dehalogenase from Sphingomonas paucimobilis UT26. FEBS Lett 446:177–181 Janssen DB (2004) Evolving haloalkane dehalogenases. Curr Opin Chem Biol 8:150–159 Janssen D (2007) Biocatalysis by dehalogenating enzymes. Adv Appl Microbiol 61:233–252 Janssen DB, Gerritse J, Brackman J, Kalk C, Jager D, Witholt B (1988) Purification and characterization of a bacterial dehalogenase with activity toward halogenated alkanes, alcohols and ethers. Eur J Biochem FEBS 171:67–72 Janssen DB, Pries F, van der Ploeg J, Kazemier B, Terpstra P, Witholt B (1989) Cloning of 1,2-dichloroethane degradation genes of Xanthobacter autotrophicus GJ10 and expression and sequencing of the dhlA gene. J Bacteriol 171:6791–6799 Jesenska A, Sedlacek I, Damborsky J (2000) Dehalogenation of haloalkanes by Mycobacterium tuberculosis H37Rv and other mycobacteria. Appl Environ Microbiol 66:219–222 Jesenska A, Bartos M, Czernekova V, Rychlik I, Pavlik I, Damborsky J (2002) Cloning and expression of the haloalkane dehalogenase gene dhmA from Mycobacterium avium N85 and preliminary characterization of DhmA. Appl Environ Microbiol 68:3724–3730 Jesenska A, Pavlova M, Strouhal M, Chaloupkova R, Tesinska I, Monincova M, Prokop Z, Bartos M, Pavlik I, Rychlik I, Möbius P, Nagata Y, Damborsky J (2005) Cloning, biochemical properties, and distribution of mycobacterial haloalkane dehalogenases. Appl Environ Microbiol 71:6736–6745 Jesenska A, Monincova M, Koudelakova T, Hasan K, Chaloupkova R, Prokop Z, Geerlof A, Damborsky J (2009) Biochemical characterization of haloalkane dehalogenases DrbA and DmbC, representatives of a novel subfamily. Appl Environ Microbiol 75:5157–5160 Kahn K, Bruice TC (2003) Comparison of reaction energetics and leaving group interactions during the enzyme-catalyzed and uncatalyzed displacement of chloride from haloalkanes. J Phys Chem 107:6876–6885 Kaushik S, Prokop Z, Damborsky J, Chaloupkova R (2016) Kinetics of binding of fluorescent ligands to enzymes with engineered access tunnels. FEBS J 284(1):134–148 Kennes C, Pries F, Krooshof GH, Bokma E, Kingma J, Janssen DB (1995) Replacement of tryptophan residues in haloalkane dehalogenase reduces halide binding and catalytic activity. Eur J Biochem FEBS 228:403–407 Keuning S, Janssen DB, Witholt B (1985) Purification and characterization of hydrolytic haloalkane dehalogenase from Xanthobacter autotrophicus GJ10. J Bacteriol 163:635–639 Kmunicek J, Luengo S, Gago F, Ortiz AR, Wade RC, Damborsky J (2001) Comparative binding energy analysis of the substrate specificity of haloalkane dehalogenase from Xanthobacter autotrophicus GJ10. Biochemistry 40:8905–8917 Kmunicek J, Bohac M, Luengo S, Gago F, Wade RC, Damborsky J (2003) Comparative binding energy analysis of haloalkane dehalogenase substrates: modelling of enzyme-substrate complexes by molecular docking and quantum mechanical calculations. J Comput Aided Mol Des 17:299–311 Kmunicek J, Hynkova K, Jedlicka T, Nagata Y, Negri A, Gago F, Wade RC, Damborsky J (2005) Quantitative analysis of substrate specificity of haloalkane dehalogenase LinB from Sphingomonas paucimobilis UT26. Biochemistry 44:3390–3401 Koudelakova T, Chovancova E, Brezovsky J, Monincova M, Fortova A, Jarkovsky J, Damborsky J (2011) Substrate specificity of haloalkane dehalogenases. Biochem J 435:345–354

18

P. Kokkonen et al.

Koudelakova T, Bidmanova S, Dvorak P, Pavelka A, Chaloupkova R, Prokop Z, Damborsky J (2013a) Haloalkane dehalogenases: biotechnological applications. Biotechnol J 8:32–45 Koudelakova T, Chaloupkova R, Brezovsky J, Prokop Z, Sebestova E, Hesseler M, Khabiri M, Plevaka M, Kulik D, Smatanova IK, Rezacova P, Ettrich R, Bornscheuer UT, Damborsky J (2013b) Engineering enzyme stability and resistance to an organic cosolvent by modification of residues in the access tunnel. Angew Chem Int Ed Engl 52:1959–1963 Kretz KA, Richardson TH, Gray KA, Robertson DE, Tan X, Short JM (2004) Gene site saturation mutagenesis: a comprehensive mutagenesis approach. Methods Enzymol 388:3–11 Krooshof GH, Kwant EM, Damborsky J, Koca J, Janssen DB (1997) Repositioning the catalytic triad aspartic acid of haloalkane dehalogenase: effects on stability, kinetics, and structure. Biochemistry 36:9571–9580 Krooshof GJ, Ridder IS, Tepper AWJW, Vos GJ, Rozeboom HJ, Kalk KH, Dijkstra BW, Janssen DB (1998) Kinetic analysis and X-ray structure of haloalkane dehalogenase with a modified halide-binding site. Biochemistry 37:15013–15023 Kuipers RK, Joosten H-J, van Berkel WJH, Leferink NGH, Rooijen E, Ittmann E, van Zimmeren F, Jochens H, Bornscheuer U, Vriend G, dos Santos VAPM, Schaap PJ (2010) 3DM: systematic analysis of heterogeneous superfamily data to discover protein functionalities. Proteins 78:2101–2113 Kumari R, Subudhi S, Suar M, Dhingra G, Raina V, Dogra C, Lal S, van der Meer JR, Holliger C, Lal R (2002) Cloning and characterization of lin genes responsible for the degradation of hexachlorocyclohexane isomers by Sphingomonas paucimobilis strain B90. Appl Environ Microbiol 68:6021–6028 Kurumbang NP, Dvorak P, Bendl J, Brezovsky J, Prokop Z, Damborsky J (2014) Computer-assisted engineering of the synthetic pathway for biodegradation of a toxic persistent pollutant. ACS Synth Biol 3:172–181 Lau EY, Kahn K, Bash P, Bruice TC (2000) The importance of reactant positioning in enzyme catalysis: a hybrid quantum mechanics/molecular mechanics study of a haloalkane dehalogenase. Proc Natl Acad Sci USA 97:9937–9942 Li A, Shao Z (2014) Biochemical characterization of a haloalkane dehalogenase DadB from Alcanivorax dieselolei B-5. PLoS One 9:e89144 Lightstone FC, Zheng YJ, Bruice TC (1998) Molecular dynamics simulations of ground and transition states for the S(N)2 displacement of Cl- from 1,2-dichloroethane at the active site of Xanthobacter autotrophicus haloalkane dehalogenase. J Am Chem Soc 120:5611–5621 Liskova V, Bednar D, Prudnikova T, Rezacova P, Koudelakova T, Sebestova E, Smatanova IK, Brezovsky J, Chaloupkova R, Damborsky J (2015) Balancing the stability-activity trade-off by fine-tuning dehalogenase access tunnels. ChemCatChem 7:648–659 Liu X, Hanson BL, Langan P, Viola RE (2007) The effect of deuteration on protein structure: a highresolution comparison of hydrogenous and perdeuterated haloalkane dehalogenase. Acta Crystallogr D Biol Crystallogr 63:1000–1008 Los G, Encell L, McDougall M, Hartzell D, Karassina N, Zimprich C, Wood M, Learish R, Ohane R, Urh M, Simpson D, Mendez J, Zimmerman K, Otto P, Vidugiris G, Zhu J, Darzins A, Klaubert D, Bulleit R, Wood K (2008) HaloTag: a novel protein labeling technology for cell imaging and protein analysis. ACS Chem Biol 3:373–382 Lüdemann SK, Lounnas V, Wade RC (2000) How do substrates enter and products exit the buried active site of cytochrome P450cam? 1. Random expulsion molecular dynamics investigation of ligand access channels and mechanisms. J Mol Biol 303:797–811 Marek J, Vevodova J, Smatanova IK, Nagata Y, Svensson LA, Newman J, Takagi M, Damborsky J (2000) Crystal structure of the haloalkane dehalogenase from Sphingomonas paucimobilis UT26. Biochemistry 39:14082–14086 Maulitz AH, Lightstone FC, Zheng YJ, Bruice TC (1997) Nonenzymatic and enzymatic hydrolysis of alkyl halides: a theoretical study of the S(N)2 reactions of acetate and hydroxide ions with alkyl chlorides. Proc Natl Acad Sci USA 94:6591–6595

Structure-Function Relationships and Engineering of Haloalkane. . .

19

Mazumdar PA, Hulecki JC, Cherney MM, Garen CR, James MNG (2008) X-ray crystal structure of Mycobacterium tuberculosis haloalkane dehalogenase Rv2579. Biochim Biophys Acta 1784:351–362 Nagata Y, Miyauchi K, Damborsky J, Manova K, Ansorgova A, Takagi M (1997) Purification and characterization of a haloalkane dehalogenase of a new substrate class from a gammahexachlorocyclohexane-degrading bacterium, Sphingomonas paucimobilis UT26. Appl Environ Microbiol 63:3707–3710 Nagata Y, Hynkova K, Damborsky J, Takagi M (1999) Construction and characterization of histidine-tagged haloalkane dehalogenase (LinB) of a new substrate class from a gammahexachlorocyclohexane-degrading bacterium, Sphingomonas paucimobilis UT26. Protein Expr Purif 17:299–304 Nagata Y, Ohtsubo Y, Tsuda M (2015) Properties and biotechnological applications of natural and engineered haloalkane dehalogenases. Appl Microbiol Biotechnol 99:9865–9881 Nam K, Prat-Resina X, Garcia-Viloca M, Devi-Kesavan LS, Gao J (2004) Dynamics of an enzymatic substitution reaction in haloalkane dehalogenase. J Am Chem Soc 126:1369–1376 Nardini M, Dijkstra BW (1999) Alpha/beta hydrolase fold enzymes: the family keeps growing. Curr Opin Struct Biol 9:732–737 Negri A, Marco E, Damborsky J, Gago F (2007) Stepwise dissection and visualization of the catalytic mechanism of haloalkane dehalogenase LinB using molecular dynamics simulations and computer graphics. J Mol Graph Model 26:643–651 Newman J, Peat TS, Richard R, Kan L, Swanson PE, Affholter JA, Holmes IH, Schindler JF, Unkefer CJ, Terwilliger TC (1999) Haloalkane dehalogenases: structure of a rhodococcus enzyme. Biochemistry 38:16105–16114 Novak HR, Sayer C, Isupov MN, Gotz D, Spragg AM, Littlechild JA (2014) Biochemical and structural characterisation of a haloalkane dehalogenase from a marine Rhodobacteraceae. FEBS Lett 588:1616–1622 Oakley AJ, Prokop Z, Bohác M, Kmunícek J, Jedlicka T, Monincová M, Kutá-Smatanová I, Nagata Y, Damborský J, Wilce MCJ (2002) Exploring the structure and activity of haloalkane dehalogenase from Sphingomonas paucimobilis UT26: evidence for product- and watermediated inhibition. Biochemistry 41:4847–4855 Oakley AJ, Klvana M, Otyepka M, Nagata Y, Wilce MCJ, Damborský J (2004) Crystal structure of haloalkane dehalogenase LinB from Sphingomonas paucimobilis UT26 at 0.95 a resolution: dynamics of catalytic residues. Biochemistry 43:870–878 Ohana RF, Encell LP, Zhao K, Simpson D, Slater MR, Urh M, Wood KV (2009) HaloTag7: a genetically engineered tag that enhances bacterial expression of soluble proteins and improves protein purification. Protein Expr Purif 68:110–120 Ollis DL, Cheah E, Cygler M, Dijkstra B, Frolow F, Franken SM, Harel M, Remington SJ, Silman I, Schrag J (1992) The alpha/beta hydrolase fold. Protein Eng 5:197–211 Olsson MH, Warshel A (2004) Solute solvent dynamics and energetics in enzyme catalysis: the S (N)2 reaction of dehalogenase as a general benchmark. J Am Chem Soc 126:15167–15179 Otyepka M, Damborsky J (2002) Functionally relevant motions of haloalkane dehalogenases occur in the specificity-modulating cap domains. Protein Sci 11:1206–1217 Otyepka M, Banas P, Magistrato A, Carloni P, Damborsky J (2008) Second step of hydrolytic dehalogenation in haloalkane dehalogenase investigated by QM/MM methods. Proteins 70:707–717 Pavlova M, Klvana M, Jesenska A, Prokop Z, Konecna H, Sato T, Tsuda M, Nagata Y, Damborsky J (2007) The identification of catalytic pentad in the haloalkane dehalogenase DhmA from Mycobacterium avium N85: reaction mechanism and molecular evolution. J Struct Biol 157:384–392 Pavlova M, Klvana M, Prokop Z, Chaloupkova R, Banas P, Otyepka M, Wade RC, Tsuda M, Nagata Y, Damborsky J (2009) Redesigning dehalogenase access tunnels as a strategy for degrading an anthropogenic substrate. Nat Chem Biol 5:727–733

20

P. Kokkonen et al.

Pieters RJ, Lutje Spelberg JH, Kellogg RM, Janssen DB (2001) The enantioselectivity of haloalkane dehalogenases. Tetrahedron Lett 42:469–471 Pikkemaat MG, Janssen DB (2002) Generating segmental mutations in haloalkane dehalogenase: a novel part in the directed evolution toolbox. Nucleic Acids Res 30:E35 Poelarends GJ, Wilkens M, Larkin MJ, van Elsas JD, Janssen DB (1998) Degradation of 1,3-dichloropropene by Pseudomonas cichorii 170. Appl Environ Microbiol 64:2931–2936 Poelarends GJ, van Hylckama Vlieg JE, Marchesi JR, Freitas Dos Santos LM, Janssen DB (1999) Degradation of 1,2-dibromoethane by Mycobacterium sp. strain GP1. J Bacteriol 181:2050–2058 Pries F, Kingma J, Pentenga M, van Pouderoyen G, Jeronimus-Stratingh CM, Bruins AP, Janssen DB (1994a) Site-directed mutagenesis and oxygen isotope incorporation studies of the nucleophilic aspartate of haloalkane dehalogenase. Biochemistry 33:1242–1247 Pries F, van den Wijngaard AJ, Bos R, Pentenga M, Janssen DB (1994b) The role of spontaneous cap domain mutations in haloalkane dehalogenase specificity and evolution. J Biol Chem 269:17490–17494 Pries F, Kingma J, Janssen DB (1995a) Activation of an asp-124->Asn mutant of haloalkane dehalogenase by hydrolytic deamidation of asparagine. FEBS Lett 358:171–174 Pries F, Kingma J, Krooshof GH, Jeronimus-Stratingh CM, Bruins AP, Janssen DB (1995b) Histidine 289 is essential for hydrolysis of the alkyl-enzyme intermediate of haloalkane dehalogenase. J Biol Chem 270:10405–10411 Prokop Z, Monincova M, Chaloupkova R, Klvana M, Nagata Y, Janssen DB, Damborsky J (2003) Catalytic mechanism of the haloalkane dehalogenase LinB from Sphingomonas paucimobilis UT26. J Biol Chem 278:45094–45100 Prokop Z, Oplustil F, DeFrank J, Damborsky J (2006) Enzymes fight chemical weapons. Biotechnol J 1:1370–1380 Prokop Z, Damborsky J, Janssen DB, Nagata Y (2009) Method of production of optically active halohydrocarbons and alcohols using hydrolytic dehalogenation catalysed by haloalkanedehalogenases. Masaryk University, Brno, Patent US 7632666 B2 Prokop Z, Sato Y, Brezovsky J, Mozga T, Chaloupkova R, Koudelakova T, Jerabek P, Stepankova V, Natsume R, van Leeuwen JGE, Janssen DB, Florian J, Nagata Y, Senda T, Damborsky J (2010) Enantioselectivity of haloalkane dehalogenases and its modulation by surface loop engineering. Angew Chem Int Ed Engl 49:6111–6115 Prokop Z, Damborsky J, Oplustil F, Jesenska A, Nagata Y (2011) Method of detoxication of yperite by using haloalkane dehalogenases. Masaryk University, Brno, Patent WO 2006/128390 A1 Prokop Z, Koudelakova T, Bidmanova S, Damborsky J (in press) Applying enzymes in degradation processes: decontamination of chemical weapons. In: Williams G, Hall M (eds) Modern biocatalysis: advances towards synthetic biological systems. The Royal Society of Chemistry, UK Ridder IS, Rozeboom HJ, Dijkstra BW (1999) Haloalkane dehalogenase from Xanthobacter autotrophicus GJ10 refined at 1.15 a resolution. Acta Crystallogr 55:1273–1290 Sallis PJ, Armfield SJ, Bull AT, Hardman DJ (1990) Isolation and characterization of a haloalkane halidohydrolase from Rhodococcus erythropolis Y2. J Gen Microbiol 136:115–120 Sato Y, Monincova M, Chaloupkova R, Prokop Z, Ohtsubo Y, Minamisawa K, Tsuda M, Damborsky J, Nagata Y (2005) Two rhizobial strains, Mesorhizobium loti MAFF303099 and Bradyrhizobium japonicum USDA110, encode haloalkane dehalogenases with novel structures and substrate specificities. Appl Environ Microbiol 71:4372–4379 Schanstra JP, Janssen DB (1996) Kinetics of halide release of haloalkane dehalogenase: evidence for a slow conformational change. Biochemistry 35:5624–5632 Schanstra JP, Kingma J, Janssen DB (1996a) Specificity and kinetics of haloalkane dehalogenase. J Biol Chem 271:14747–14753 Schanstra JP, Ridder A, Kingma J, Janssen DB (1997) Influence of mutations of Val226 on the catalytic rate of haloalkane dehalogenase. Protein Eng 10:53–61

Structure-Function Relationships and Engineering of Haloalkane. . .

21

Schanstra JP, Ridder IS, Heimeriks GJ, Rink R, Poelarends GJ, Kalk KH, Dijkstra BW, Janssen DB (1996b) Kinetic characterization and X-ray structure of a mutant of haloalkane dehalogenase with higher catalytic activity and modified substrate range. Biochemistry 35:13186–13195 Schindler JF, Naranjo PA, Honaberger DA, Chang CH, Brainard JR, Vanderberg LA, Unkefer CJ (1999) Haloalkane dehalogenases: steady-state kinetics and halide inhibition. Biochemistry 38:5772–5778 Scholtz R, Schmuckle A, Cook A, Leisinger T (1987) Degradation of 1-monohaloalkanes by Arthrobacter sp. strain-HA1. J Gen Microbiol 133:267–274 Sfetsas CC, Milios L, Skopelitou K, Venieraki A, Todou R, Flemetakis E, Katinakis P, Labrou NE (2009) Characterization of 1,2-dibromoethane-degrading haloalkane dehalogenase from Bradyrhizobium japonicum USDA110. Enzym Microb Technol 45:397–404 Shurki A, Strajbl M, Villa J, Warshel A (2002) How much do enzymes really gain by restraining their reacting fragments? J Am Chem Soc 124:4097–4107 Silberstein M, Dennis S, Brown L, Kortvelyesi T, Clodfelter K, Vajda S (2003) Identification of substrate binding sites in enzymes by computational solvent mapping. J Mol Biol 332:1095–1113 Soriano A, Silla E, Tunon I (2003) Internal rotation of 1,2-dichloroethane in haloalkane dehalogenase. A test case for analyzing electrostatic effects in enzymes. J Phys Chem 107:6234–6238 Soriano A, Silla E, Tunon I, Ruiz-Lopez MF (2005) Dynamic and electrostatic effects in enzymatic processes. An analysis of the nucleophilic substitution reaction in haloalkane dehalogenase. J Am Chem Soc 127:1946–1957 Streltsov VA, Prokop Z, Damborský J, Nagata Y, Oakley A, Wilce MCJ (2003) Haloalkane dehalogenase LinB from Sphingomonas paucimobilis UT26: X-ray crystallographic studies of dehalogenation of brominated substrates. Biochemistry 42:10104–10112 Stucki G, Thuer M (1995) Experiences of a large-scale application of 1,2-dichloroethane degrading microorganisms for groundwater treatment. Environ Sci Technol 29:2339–2345 Swanson P (1999) Dehalogenases applied to industrial-scale biocatalysis. Curr Opin Biotechnol 10:365–369 Sykora J, Brezovsky J, Koudelakova T, Lahoda M, Fortova A, Chernovets T, Chaloupkova R, Stepankova V, Prokop Z, Smatanova IK, Hof M, Damborsky J (2014) Dynamics and hydration explain failed functional transformation in dehalogenase design. Nat Chem Biol 10:428–430 van Leeuwen JGE, Wijma HJ, Floor RJ, van der Laan J-M, Janssen DB (2012) Directed evolution strategies for enantiocomplementary haloalkane dehalogenases: from chemical waste to enantiopure building blocks. Chembiochem 13:137–148 Verschueren KH, Franken SM, Rozeboom HJ, Kalk KH, Dijkstra BW (1993a) Refined X-ray structures of haloalkane dehalogenase at pH 6.2 and pH 8.2 and implications for the reaction mechanism. J Mol Biol 232:856–872 Verschueren KH, Kingma J, Rozeboom HJ, Kalk KH, Janssen DB, Dijkstra BW (1993b) Crystallographic and fluorescence studies of the interaction of haloalkane dehalogenase with halide ions. Studies with halide compounds reveal a halide binding site in the active site. Biochemistry 32:9031–9037 Verschueren KH, Seljée F, Rozeboom HJ, Kalk KH, Dijkstra BW (1993c) Crystallographic analysis of the catalytic mechanism of haloalkane dehalogenase. Nature 363:693–698 Westerbeek A, Szymanski W, Feringa BL, Janssen DB (2011a) Dynamic kinetic resolution process employing haloalkane dehalogenase. ACS Catal 1:1654–1660 Westerbeek A, Szymanski W, Wijma HJ, Marrink SJ, Feringa BL, Janssen DB (2011b) Kinetic resolution of alpha-bromoamides: experimental and theoretical investigation of highly enantioselective reactions catalyzed by haloalkane dehalogenases. Adv Synth Catal 353:931–944 Wijma HJ, Floor RJ, Jekel PA, Baker D, Marrink SJ, Janssen DB (2014) Computationally designed libraries for rapid enzyme stabilization. Protein Eng Des Sel 27:49–58 Yokota T, Omori T, Kodama T (1987) Purification and properties of haloalkane dehalogenase from Corynebacterium sp. strain m15-3. J Bacteriol 169:4049–4054

Aerobic Degradation of Gasoline Ether Oxygenates Michael Hyman

Abstract

Ether oxygenates including methyl tertiary butyl ether (MTBE), ethyl tertiary butyl ether (ETBE), tertiary amyl ether (TAME), and diisopropyl ether (DIPE) have been, and continue to be, widely used components of gasoline. The ether bonds and branched hydrocarbon structures of these compounds make these challenging molecules for microbial biodegradation processes. The collective research over the last 20 years suggests that aerobic biodegradation of MTBE and other ether oxygenates by axenic cultures occurs through three physiologically distinct processes that can be differentiated by the fate of the tertiary alcohol intermediates such as tertiary butyl alcohol (TBA) and tertiary amyl alcohol (TAA) that are common to all of these processes. These biodegradation processes represent a continuum and, in order of increasing complexity, include co-oxidation, cometabolism, and growth-supporting metabolism. This review summarizes the main microorganisms, enzymes, and pathways involved in each of these processes and highlights research areas where there is both clear consensus and areas where results are more ambiguous and likely require further investigation. Where relevant, this review also aims to illustrate how basic microbiological research has implications for the remediation of ether oxygenate contamination by aerobic treatment processes.

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1 Important Characteristics of Gasoline Ether Oxygenates . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2 Range of Aerobic Ether Oxygenate Biodegradation Processes . . . . . . . . . . . . . . . . . . . . . . 1.3 Microbial Co-oxidation of MTBE and Other Ether Oxygenates . . . . . . . . . . . . . . . . . . . .

2 3 3 4

M. Hyman (*) Department of Plant and Microbial Biology, North Carolina State University, Raleigh, NC, USA e-mail: [email protected] # Springer International Publishing AG 2016 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-319-39782-5_16-1

1

2

M. Hyman 1.4 1.5 1.6 1.7 1.8 1.9 1.10 1.11 1.12 1.13

Co-oxidation of Ether Oxygenates by Pseudomonas Strains . . . . . . . . . . . . . . . . . . . . . . . . Co-oxidation of Ether Oxygenates by Pseudonocardia Strains . . . . . . . . . . . . . . . . . . . . . Co-oxidation of Ether Oxygenates by Rhodococcus Strains . . . . . . . . . . . . . . . . . . . . . . . . . Co-oxidation of Ether Oxygenates by Fungi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Syntrophic Processes Involving Ether Oxygenate Co-oxidation . . . . . . . . . . . . . . . . . . . . Microbial Cometabolism of MTBE and Other Ether Oxygenates . . . . . . . . . . . . . . . . . . . Microbial Metabolism of MTBE and Other Ether Oxygenates . . . . . . . . . . . . . . . . . . . . . . Growth of Hydrogenophaga flava ENV735 on MTBE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Growth of Methylibium petroleiphilum PM1 on MTBE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Growth of Aquincola tertiaricarbonis L108 on MTBE and Other Ether Oxygenates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.14 Cytochrome P450 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.15 Tertiary Alcohol Monooxygenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.16 2HIBA-CoA Mutase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.17 Compound Specific Isotope Analysis (CSIA) of Aerobic Ether Oxygenate Biodegradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.18 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

4 6 6 8 10 11 14 15 15 17 18 18 20 22 24 25

Introduction

Ether oxygenates have been a major component of the global gasoline supply for over 30 years. As their collective name implies, these chemicals all contain oxygen in the form of an ether bond (Fig. 1). This oxygen can improve the efficiency of gasoline combustion and decrease levels of air pollutants such as CO, NOx, and ozone-forming volatiles in automobile exhaust (ITRC 2005). Ether oxygenates also have high octane ratings and are often added to gasoline at high concentrations (15 vol %), (ITRC 2005). Consequently, they are also valuable blending components that can extend the overall supply of gasoline. However, these compounds have also become important environmental contaminants, frequently as the result of accidental spills of gasoline. Much of the research into biodegradation of these compounds aims to improve our understanding of the environmental fate of these compounds and mechanisms that can be used to remediate this contamination. This review focuses on the individual microorganisms, pathways, and enzymes involved in the aerobic biodegradation of the major ether oxygenates MTBE, ETBE, TAME, and, to a lesser extent, DIPE (Fig. 1). While the biochemical basis for several syntrophic processes for ether oxygenate mineralization is briefly addressed, the many reports of undefined aerobic microbial cultures that can degrade MTBE and other ether oxygenates are not discussed. Likewise, even though ethanol is also widely used as a gasoline oxygenate, biodegradation of alcohols is also not discussed in this review. The notable exceptions to this are tertiary butyl alcohol (TBA) and tertiary amyl alcohol (TAA) (Fig. 1). Although TBA has also been used itself as a gasoline oxygenate (ITRC 2005), the main interest in these tertiary alcohols in this review is that they are terminal or transient metabolites in all aerobic ether oxygenate biodegradation processes.

Aerobic Degradation of Gasoline Ether Oxygenates

3

Fig. 1 Structures of the four main ether oxygenates and their corresponding major metabolites

1.1

Important Characteristics of Gasoline Ether Oxygenates

Compared to other well-studied and more toxic components of gasoline such as benzene, the physical properties of ether oxygenates are notable for their generally high vapor pressures, high aqueous solubilities, and low Henry’s law constants. While these physical properties are important for understanding the environmental distribution of these compounds, the chemical structures of ether oxygenates most directly impact their biodegradation. Historically, ether oxygenates have often been reported as poorly biodegradable or as recalcitrant compounds, and the ether bond and branched hydrocarbon structures consistently found in these compounds are both individual features that have long been recognized as general impediments to microbial biodegradation (White et al. 1996; Alexander 1973). The ether linkage itself has been described as “. . .the single most common and unifying structural feature which confers to both biological and xenobiotic compounds a high degree of resistance to biological mineralization” (White et al. 1996). The significance of the branched structures of ether oxygenates can perhaps be judged from a survey of newly isolated ether-degrading bacteria (Kim et al. 2007). Out of 27 isolates, 17 isolates grew on n-butyl methyl ether or n-propyl ether, but none grew on their respective branched isomers, MTBE and DIPE.

1.2

Range of Aerobic Ether Oxygenate Biodegradation Processes

The majority of the research into aerobic biodegradation of ether oxygenates by axenic microbial cultures has focused on MTBE, and this emphasis is reflected in

4

M. Hyman

Fig. 2 The role of a hemiacetal intermediate (hydroxymethyl tertiary butyl ether) in the production of TBA from MTBE. Solid lines represent enzyme-catalyzed reactions. Dashed lines represent abiotic reactions. The type of enzyme involved in each reaction is shown in italics. Undetected intermediates are shown in brackets

this review. Collectively, research into aerobic ether oxygenate biodegradation suggests there are three physiologically distinct processes that can be distinguished by the fate of tertiary alcohols that are generated in each case. These processes lie on a physiological continuum ranging from single step co-oxidation reactions through to growth-related metabolism. Each type of biodegradation process is discussed separately in the following sections.

1.3

Microbial Co-oxidation of MTBE and Other Ether Oxygenates

The simplest aerobic biodegradation process for MTBE and other ether oxygenates is co-oxidation by microorganisms that do not grow on these compounds but can transform them after or during growth on other, often structurally unrelated, substrates. Microorganisms that co-oxidize MTBE include, among others, Pseudomonas, Pseudonocardia, and Rhodococcus strains, as well as some fungi. These microorganisms all oxidize MTBE using nonspecific monooxygenases. The sole multicarbon product of MTBE oxidation is TBA, which is typically not further oxidized at any significant rate and accumulates extracellularly. Production of TBA from MTBE by monooxygenases is presumed to involve an initial hydroxylation of the methoxy carbon leading to formation of an unstable hemiacetal, hydroxymethyl tertiary butyl ether. This hemiacetal then typically undergoes rapid abiotic dismutation to equimolar amounts of TBA and formaldehyde (Fig. 2). A similar process also occurs during ETBE and TAME oxidation, leading to mixtures of TBA and acetaldehyde and TAA and formaldehyde, respectively.

1.4

Co-oxidation of Ether Oxygenates by Pseudomonas Strains

Co-oxidation of MTBE and other ether oxygenates has been most extensively studied in Pseudomonas strains that grow on shorter chain (C10) in the presence of either MTBE or TBA. In contrast, no immediate MTBE- or TBA-oxidizing activity was detected in resting cell assays using cells previously grown on longer chain n-alkanes in the absence of MTBE or TBA. Two other strains closely related to M. vaccae JOB5, Mycobacterium austroafricanum strains IFP 2012 and 2015, have been described that have questionable abilities to grow on MTBE (François et al. 2002; Ferreira et al. 2006a). Even with high initial inoculum densities (0.1–0.4 OD600), complete consumption of modest concentrations of MTBE (70–140 mg MTBE L 1) by these strains requires several weeks and results in small changes (~0.1 OD600) in culture density. In contrast to MTBE, growth of strains IPF 2012 and 2015 on high concentrations of TBA is more robust. The results of physiological studies with these IFP strains are similar to those described for MTBE-degrading propanotrophs (Steffan et al. 1997; Smith et al. 2003a). For example, in both strains, the degradation of MTBE involves production of TBA and other intermediates including TBF, 2MPD, and 2HIBA (François et al. 2003). Acetylene also inhibits both MTBE and TBA oxidation, suggesting the same monooxygenase oxidizes both MTBE and TBA. While a Ks value for MTBE has not been determined, a Ks for TBA has been estimated at 1.1 mM (François et al. 2002). The TBA-oxidizing activity of both strains IFP 2012 and IFP 2015 is also promoted by growth on n-alkanes including propane, n-hexane, and n-hexadecane (Ferreira et al. 2007). Although a single AlkB-like alkane hydroxylase capable of oxidizing C3 to C16 n-alkanes has not been previously identified, it has been proposed that this type of enzyme is responsible for the TBA (and hence MTBE)oxidizing activity in M. vaccae JOB5 and both strains IFP 2012 and IFP 2015 after growth on these n-alkanes (Ferreira et al. 2007). At least in the case of M. vaccae JOB5, this is unlikely for several reasons. First, the Ks values previously determined for MTBE oxidation by the alkane hydroxylase in P. putida GPo1 is at least an order of magnitude higher than the Ks values for MTBE and TBA in strain JOB5, and this enzyme does not oxidize TBA. Second, M. vaccae JOB5 possesses two alkB genes, and RT-qPCR analyses have indicated that neither of these genes are expressed during growth on propane (Sharp et al. 2010). Lastly, it is clear that hydrocarbonoxidizing bacteria can express multiple monooxygenases in response to alkanes, and these enzymes may not always be related to the biodegradation activity under investigation. As mentioned earlier, the MTBE-degrading propanotroph

14

M. Hyman

Mycobacterium strain ENV421 expresses at least three different monooxygenases in response to propane, and at least two of these enzymes do not oxidize this gas (Masuda et al. 2012b). Two other genes have been more clearly implicated in the TBA-oxidizing activity of M. austroafricanum strain IFP 2012. These genes, mpbB and mpbC, encode a choline dehydrogenase-like flavoprotein and an aldehyde dehydrogenase-like enzyme, respectively (Ferreria et al. 2006b). These genes enable oxidation of 2MPD to 2HIBA when they are heterologously co-expressed. It was proposed that MpbB oxidizes 2MPD to 2-hydroxyisobutyraldehyde (2HIBAL) and that MpbC further oxidizes this aldehyde to 2HIBA. Although 2HIBAL has not been detected as an intermediate in any studies of MTBE oxidation to date, it has been suggested that MpbC rapidly oxidizes this aldehyde so that it does not accumulate. However, mpbB and mpbC were not individually cloned and expressed so the role of 2HIBAL as an intermediate in MTBE oxidation remains unresolved, especially as some cholineoxidizing enzymes can catalyze 4-electron oxidations of their alcohol substrates to acids without the release of an aldehyde intermediate (Fan et al. 2006). Genes similar to mpbB and mpbC have not been detected in the genomes of bacteria such as Methylibium petroleiphilum PM1 that grow more reliably on both MTBE and TBA (Kane et al. 2007). Little else is known about the intermediates that are generated after 2HIBA by the propanotrophic, MTBE-degrading actinomycetes discussed above. Taken together, the studies with these strains suggest that the various steps in MTBE catabolism are relatively slow fortuitous reactions catalyzed by enzymes whose activities are not coordinated to the level found in bacteria that grow more consistently on ether oxygenates. While growth of some Mycobacterium strains can be supported by high concentrations of TBA, the fact that MTBE- and TBA-derived metabolites typically accumulate extracellularly and then undergo subsequent degradation suggests that different enzyme systems may need to be sequentially induced to achieve significant catabolism of MTBE. It should also be noted that while there is clear evidence for an initial series of reactions converting MTBE to 2HIBA, these reactions do not provide either energy or carbon for these microorganisms (Fig. 4). It may be that the metabolism of the C1 products of MTBE oxidation play an important role in sustaining MTBE and TBA oxidation. Conversely, as trace levels of metabolites such as acetone have been reported during the degradation of MTBEderived metabolites (François et al. 2003), it may be that the consistent ability of these organisms to metabolize propane, which often involves acetone as an intermediate, also facilitates the later stages of MTBE catabolism.

1.10

Microbial Metabolism of MTBE and Other Ether Oxygenates

The final biodegradation process for MTBE and other ether oxygenates involves the unambiguous use of these compounds as sole sources of carbon and energy for growth. While several of the intermediates identified in the studies of MTBE cometabolism by the Mycobacterium strains described above are also involved in

Aerobic Degradation of Gasoline Ether Oxygenates

15

growth-supporting metabolism of MTBE, these intermediates do not typically accumulate and are efficiently consumed in a coherent and coordinated pathway. These organisms are further distinguished from the MTBE-cometabolizing actinomycetes discussed above by the fact that they are all Gram-negative bacteria.

1.11

Growth of Hydrogenophaga flava ENV735 on MTBE

The H2-utilizing bacterium Hydrogenophaga flava ENV735 was isolated from an MTBE-fed enrichment culture and grows slowly on both MTBE and TBA as sole sources of carbon and energy (Hatzinger et al. 2001). Growth on MTBE is enhanced by yeast extract but is unaffected by H2. The biomass yield on MTBE is 0.4 g g 1, and both physiological and inhibitor studies indicate that two different enzymes are responsible for oxidizing MTBE and TBA in this strain. The MTBE-oxidizing activity is constitutive and is partially inhibited by aminobenzotriazole and allylthiourea, but not by CO or acetylene. In contrast, TBA-oxidizing activity is inducible. In non-induced cells, TBA therefore temporarily accumulates during MTBE oxidation before it is subsequently consumed. The oxidation of TBA is only partially inhibited by acetylene and unaffected by the cytochrome P450 inhibitors, CO, and aminobenzotriazole. Low levels of 2HIBA are also be detected during MTBE oxidation, and this acid also supports growth of strain ENV735. Strain ENV735 also grows on formaldehyde, and MTBE oxidation in formaldehydegrown cells is transiently inhibited until formaldehyde is fully consumed. An adhesion-deficient mutant strain of ENV735 suitable for bioaugmentation has also been described (Streger et al. 2002), but no other studies describing the physiology, pathway, or enzymes involved in ether oxygenate biodegradation by this strain have been reported.

1.12

Growth of Methylibium petroleiphilum PM1 on MTBE

Methylibium petroleiphilum PM1 was originally isolated from an MTBE-degrading mixed culture growing in a peat moss biofilter (Hanson et al. 1999). Like H. flava ENV735, this bacterium grows slowly on MTBE, and it mineralizes or assimilates ~65% of 14C from [U-14C]-MTBE. A growth yield of 10-fold increase in both mdpE and mdpJ expression. Clearer evidence for roles of mdpA and mdpJ in MTBE-degrading activity were reported after it was recognized that ethanol-grown cells have significant levels of mdpA transcription (Joshi et al. 2016). For example, time course studies with pyruvate-grown cells have shown expression of mdpA and mdpJ increases by ~8- and ~27-fold, respectively, when cells are exposed to MTBE. Expression of these genes also increases by >80- and >60-fold, respectively, during exposure to TBA (Joshi et al. 2016). Another growth-supporting substrate, benzene, delays mdpA and mdpJ expression, while MTBE does not impact transcription of bmoA, which encodes a component of benzene monooxygenase. Ethylbenzene, which strongly inhibits MTBE oxidation by strain PM1 (Deeb et al. 2001), also strongly inhibits transcription of mdpA, mdpJ, and bmoA (Joshi et al. 2016). The protein encoded by mdpA is an AlkB-like alkane hydroxylase. Homologues of an alkG-like rubredoxin, an alkT-like rubredoxin reductase, and a putative alkSlike transcriptional regulator (mdpC) do not occur as a discrete operon but all reside within a 10 kb locus on the pPM1 plasmid (Schmidt et al. 2008). Methimazole (10 mM) was shown to inhibit both MTBE and TBA oxidation by MTBE-grown cells but only MTBE oxidation in TBA-grown cells. These effects were interpreted to suggest MdpA is a flavin-containing enzyme. However, methimazole is a reversible competitive inhibitor of flavin monooxygenases (FMOs) and is neither an inactivator nor a broad-spectrum inhibitor of all flavin-containing enzymes (Ziegler 2002; Tomasi et al. 1995). Despite this, when the wild-type copy of mdpA is replaced

Aerobic Degradation of Gasoline Ether Oxygenates

17

with a mutant version, only wild-type cells oxidize MTBE, while both the wild-type and mutant strains oxidize TBA at similar rates. Complementation experiments were unable to recover strains that could grow on MTBE although complemented cells grown on ethanol could oxidize low concentrations of MTBE, while similar but very low rates of TBA oxidation were observed in both ethanol-grown wild-type and complemented cells. A recent study has focused on the putative transcriptional activator, mdpC, and has demonstrated that the translation inhibitor chloramphenicol inhibits transcription of mdpA and mdpJ but not mdpC in pyruvate-grown cells exposed to MTBE (Joshi et al. 2015). In mdpC strains, transcription of both mdpA and mdpJ is strongly inhibited compared to wild-type cells during exposure to either MTBE or TBA. In contrast, rates of TBA oxidation are ~40% faster in mdpC cells compared to wild-type cells, while rates of MTBE oxidation are inhibited by ~70% in mdpC cells. While some uncertainty still surrounds the role of mdpA and its regulation in MTBE oxidation by strain PM1, another closely related strain, Methylibium R8, exhibits very similar 13/12C and 2/1H enrichments to strain PM1 in compoundspecific isotope analysis (CSIA) of MTBE oxidation. This suggests strains PM1 and R8 very likely have similar MTBE-oxidizing enzymes (Rosell et al. 2007), and strain R8 also contained an mdpA gene. A mixed MTBE-degrading culture (US3-M) also exhibits similar 13/12C and 2/1H fractionation to strains PM1 and R8 (Bastida et al. 2010). This study also used 13C-protein stable isotope probing and 2D-PAGE analysis to identify proteins produced by the [U-13C]-MTBE-utilizing members of the mixed culture. All 23 detected proteins were closely related to proteins encoded in the genome of strain PM1. These proteins included both MdpJ and MdpK, as well as a series of uncharacterized proteins (MpeB0532-MpeB0535) whose genes were previously shown to be strongly expressed in MTBE-grown cells of strain PM1 (Hristova et al. 2007). However, neither MdpA nor the AlkG-like rubredoxin and AlkT-like rubredoxin reductase proposed to be directly involved in MTBE oxidation by strain PM1 (Schmidt et al. 2008) were detected. While a failure to detect MdpA may reflect its membrane-bound nature and possible poor resolution in PAGE systems, the AlkT- and AlkG-like proteins proposed to be associated with MdpA are both soluble cytoplasmic proteins. In addition to MdpA, a variety of other enzymes associated with MTBE oxidation by strain PM1 have been characterized, but the research leading to their identification has been often been conducted in other MTBE-metabolizing strains such as Aquincola tertiaricarbonis L108. These studies are described in the following section.

1.13

Growth of Aquincola tertiaricarbonis L108 on MTBE and Other Ether Oxygenates

To date the most complete understanding of the enzymes and pathway of ether oxygenate metabolism has been obtained with A. tertiaricarbonis L108. Strain L108 was isolated on MTBE by enrichment cultures inoculated with groundwater from an

18

M. Hyman

MTBE-contaminated site in Germany. Strain L10 is a spontaneous mutant of strain L108 that does not grow on MTBE but can still grow on TBA. In the description of the new genus Aquincola, strains L108 and L10 and the TBA-utilizing strain Burkholderia CIP I-2052 (Piveteau et al. 2001) described earlier were all proposed as A. tertiaricarbonis species that are physiologically related by their unusual abilities to use compounds containing tertiary carbons such as TBF, TBA, TAA, and 2HIBA as growth-supporting substrates (Lechner et al. 2007). Strain L108 exhibits 95.6% 16S rRNA gene similarity to M. petroleiphilum PM1 and is versatile bacterium that can grow on MTBE, ETBE, TAME, TBA, TAA, and 2HIBA (Lechner et al. 2007; Müller et al. 2008). Among these substrates, MTBE supports the lowest maximal growth rate (0.045 h 1), while the fastest growth occurs with 2HIBA (0.17 h 1). Growth yields on MTBE are ~0.5 g g 1 and as high as 0.8 g g 1 with TAME and TAA (Müller et al. 2008). However, obtaining these yields requires high rates of substrate utilization to overcome the high minimum substrate concentration and maintenance energy barrier required for growth. The Ks values for MTBE and ETBE have been estimated at ~1.9 mM and ~0.1 mM, respectively (Müller et al. 2008), and the initial monooxygenase-catalyzed ether-cleaving step in ether oxygenate metabolism is viewed as a key rate-limiting step controlling the rate of assimilable carbon production (Müller et al. 2007). Three key enzymes involved in MTBE metabolism have been characterized in A. tertiaricarbonis species, including this monooxygenase. These enzymes are individually discussed in the following sections:

1.14

Cytochrome P450

The initial oxidation of ether oxygenate in strain L108 is catalyzed by the same ethgene-encoded cytochrome P450 originally identified in ETBE-degrading Rhodococcus strains (Chauvaux et al. 2001). However, unlike the ethRABCD operon in Rhodococcus ruber IFP 2001, the operon in strain L108 lacks the transcriptional regulator, ethR, and the entire eth operon is absent in strain L10. Consequently, strain L108 constitutively expresses the eth-gene-encoded cytochrome P450, while strain L10 does not grow on or oxidize any ether oxygenates (Schuster et al. 2013; Lechner et al. 2007). Like R. ruber IFP 2001, strain L108 can oxidize a wide range of other ethers, including DIPE. While acetone has been reported as the sole product of DIPE oxidation (Schuster et al. 2013), it seems likely that 2-propanol, the other predicted product of DIPE oxidation, is also further oxidized to acetone during DIPE oxidation (Schäfer et al. 2012).

1.15

Tertiary Alcohol Monooxygenase

The further oxidation of TBA to 2MPD by strain L108 is catalyzed by a phthalate dioxygenase-like enzyme. This enzyme was first identified in an analysis of proteins differentially produced by cells grown on lactate, 2HIBA, or TBA (Schäfer et al.

Aerobic Degradation of Gasoline Ether Oxygenates

19

2007). A 2D-PAGE analysis identified both the 55 kDa hydroxylase and 38 kDa reductase components of this enzyme, and these proteins were both shown to be closely related to proteins predicted from two adjacent plasmid-borne genes in the genome of M. petroleiphilum PM1. As discussed earlier, these genes, mdpJK, have been strongly implicated in MTBE oxidation through a variety of approaches. The enzyme encoded by these genes is thought, like other similar dioxygenases, to be able to act as a monooxygenase and is recognized as a tertiary alcohol monooxygenase. Studies of the activity of MdpJ in wild-type whole cells and in mdpJ knockout strains have demonstrated several interesting reactions involved in ether oxygenate catabolism. For instance, low concentrations of isobutylene (2-methylpropene) are produced by A. tertiaricarbonis L108, M. petroleiphilum PM1, and Methylibium strain R8 during MTBE, ETBE, or TBA oxidation, but not during oxidation of 2MPD or 2HIBA. Higher concentrations of isoamylene (2-methyl-2-butene) are produced during oxidation of TAME or TAA. Both of these alkene-forming reactions were initially attributed to the activity of MdpJ (Schäfer et al. 2011), but studies with mdpJ knockout strains have subsequently shown that these gases are more likely produced by a dehydratase activity of an isomerase enzyme (Schuster et al. 2012). The role of MdpJK in the metabolism of TAME and TAA has also been examined. Unlike the monooxygenase-catalyzed oxidation of TBA to 2MPD, oxidation of TAA can potentially generate multiple hydroxylated products. Based on the sequential conversion of TBA to 2MPD and 2HIBA involved in MTBE oxidation (Fig. 4), the analogous products of TAA oxidation might be expected to be a variety of diols and their corresponding hydroxy acids. However, oxidation of TAA by chloramphenicol-treated, TBA-grown cells of strain L108 results in accumulation of 2-methyl-3-buten-2-ol, prenol (3-methyl-2-buten-1-ol), prenal (3-methyl-2butenal), and 3-methylcrotonic acid (Schuster et al. 2012). In the absence of chloramphenicol, these metabolites transiently accumulate but are then consumed by TBA-grown cells. In similar experiments with TAA-grown cells, only low concentrations of 2-methyl-3-buten-2-ol accumulate. Collectively, these results suggest that the catabolic routes for TBA and TAA are different in strain L108. These results further suggest that the main metabolites produced from TAA by TBA-grown cells are intermediates in TAA oxidation, and the enzymes necessary for their catabolism are inducible in TBA-grown cells but are present at high levels in TAA-grown cells. A pathway for the catabolism of TAA by strain L108 has been proposed in which TAA is first converted to 2-methyl-3-buten-2-ol by a desaturase activity of MdpJ. This alcohol then undergoes isomerization to prenol followed by sequential dehydrogenase-catalyzed oxidations of prenol to prenal and 3-methylcrotonic acid. This metabolite then likely enters the well-established leucine degradation pathway before passing to central metabolic pathways. In support of this, TAA metabolism appears to require biotin, which is required for a carboxylation step involved in leucine catabolism. However, unlike growth on TBA, growth on TAA does not require cobalamin. None of the enzymes involved in this putative pathway other than MdpJK have been identified, but an initial isomerization step has been implicated in

20

M. Hyman

the catabolism of 2-methyl-3-buten-2-ol by a Pseudomonas putida strain (Malone et al. 1999). In addition to the reaction catalyzed with TAA, MdpJ also desaturates 2-butanol, 3-methyl-2-butanol, and 3-pentanol to 3-buten-2-ol, 3-methyl-3-buten-2-ol, and 1-penten-3-ol, respectively (Schäfer et al. 2012).

1.16

2HIBA-CoA Mutase

The third enzyme involved in MTBE oxidation that has been characterized in A. tertiaricarbonis strains isomerizes 2HIBA-CoA and 3-hydroxybutyrate-CoA. This isomerization converts the branched structure of 2HIBA to a more readily metabolizable linear structure. The activity of this enzyme, 2-hydroxybutyryl-CoA mutase (Hcm), was first characterized in growth studies with all three A. tertiaricarbonis strains. Rohwerder et al. (2006) demonstrated growth of strain L108 on MTBE, TBA, or 2HIBA, and growth of strains L10 and CIP I-2052 on TBA or 2HIBA is dependent on cobalt and is accelerated when cobalt is replaced with cobalamin. In contrast, growth of these strains on substrates lacking a tertiary butyl moiety is cobalt independent. 3-Hydroxybutyrate has also been shown to accumulate during 2HIBA degradation in whole-cell studies with strain L10 and during an ATPand CoA-dependent reaction in cell-free extracts of 2HIBA-grown cells of strain L108. Hcm was initially identified by 2D-PAGE analyses and mass spectral analysis of proteins selectively produced during growth of strain L108 on 2HIBA. Initially, a small (~15 kDa) protein was identified from the genome of M. petroleiphilum PM1 as a subunit of methyl-malonyl-CoA mutase. Subsequent analyses of the relevant genes in strains PM1 and L108 identified this protein as a cobalamin-binding subunit of isobutyryl-CoA mutase (ICM). Both strains L108 and PM1 contain near identical genes for an enzyme consisting of a large acyl-CoA substrate-binding subunit and a smaller cobalamin-binding subunit (Rohwerder et al. 2006). These subunits are now known as HcmA and HcmB, respectively. The genes encoding these proteins occur in an operon in which hcmA is separated from hcmB by genes encoding an acyl-CoA synthetase and a chaperone (Yaneva et al. 2012). Knockout strains lacking either hcmA or hcmB both still grow on TAA but are unable to grow on either TBA or 2MPD and excrete stoichiometric amounts of 2HIBA in resting cell assays containing TBA or 2MPD (Yaneva et al. 2012). Kinetic studies with heterologously expressed and affinity-purified HcmA and HcmB indicate that both subunits are required for mutase activity. The reconstituted enzyme is highly specific for the isomerization of 2HIBA-CoA and 3-hydroxybutyryl-CoA and exhibits kcat/Km values for these substrates that are several orders of magnitude greater than for reactions involving either butyryl- or isobutyryl-CoA catalyzed by other mutases. The enzyme also shows considerable stereospecificity and predominantly generates the (S)- rather than the (R)-enantiomer of 3-hydroxybutyryl-CoA (Yaneva et al. 2012). Several recent studies have focused on potential applications of Hcm as catalyst for generating 2HIBA from renewable sources of 3-hydroxybutyrate such as

Aerobic Degradation of Gasoline Ether Oxygenates

21

polyhydroxybutyrate (PHB). In PHB biosynthesis, acetyl-CoA is initially converted into (R)-3-hydroxybutyryl-CoA but not the (S)-enantiomer. Crystallographic studies have investigated the structural features of Hcm from A. tertiaricarbonis L108 that impact its stereospecificity (Kurteva-Yaneva et al. 2015), and variants of Hcm that isomerize 2HIBA-CoA and (R)-3-hydroxybutyryl-CoA have also now been described in a thermophilic H2-oxidizing bacterium, Kyrpidia tusciae (Weichler et al. 2015) and Bacillus massiliosenegalensis (Rohde et al. 2016). Production of 2HIBA by diverting metabolites from PHB synthesis through Hcm has also been demonstrated both in recombinant strains of Cuprividus necator H16 grown on H2 + CO2 (Przybylski et al. 2015) and methanol-grown Methylobacterium extorquens (Rohde et al. 2016). The key role of Hcm in MTBE-metabolizing bacteria is clearly illustrated in Fig. 4. Ignoring both the potential benefits and detriments provided by the production and further oxidation of the initial C1 products of MTBE oxidation, the steps preceding 2HIBA production involve two reductant-generating dehydrogenase-catalyzed reactions and two reductant-consuming monooxygenase-catalyzed reactions, including the initial oxidation of MTBE. The subsequent steps initiated by Hcm-dependent 2HIBA isomerization are therefore the only major reactions that can yield either carbon or energy for growth on MTBE and TBA. Given the importance of Hcm, it not surprising that it has been detected in other studies that have examined the degradation of MTBE and TBA by undefined mixed microbial communities. For example, hcmA was detected in 13C-metagenomic DNA obtained in an SIP study examining [U-13C] TBA degradation in a biological granular activated carbon (bio-GAC) reactor (Aslett et al. 2011). However, it is less clear why very similar genes are found in bacteria including Rhodobacter sphaeroides, Nocardioides JS614, and Xanthobacter autotrophicus Py2 that are not known to have either MTBE- or TBA-degrading activities (Rohwerder et al. 2006). As originally suggested by Rohwerder and Müller (2010), a Hcm-like reaction is likely involved in the cobalt-dependent growth of Mycobacterium sp. ELW1 on isobutylene. In strain ELW1, 2HIBA is a growth substrate and a detected intermediate in isobutylene catabolism (Kottegoda et al. 2015). Notably, like strain ELW1, Nocardioides JS614 and X. autotrophicus Py2 are also alkene-metabolizing bacteria. Although strains JS614 and Py2 can both co-oxidize isobutylene (Owens et al. 2009; Ensign 1996), neither strain has been reported to grow on this gas. One final important issue concerning Hcm is the significance of the cobalamin required by this enzyme. Cobalamin biosynthesis is energetically expensive, and microbial community structure can be strongly impacted by the presence or absence of cobalamin-synthesizing bacteria (Degnan et al. 2014). While organisms such as M. petroleiphilum PM1 have a complete cobalamin biosynthesis capability (Kane et al. 2007), it is not clear how widely this capability is distributed among other MTBE- and TBA-metabolizing microorganisms in the environment and whether ether oxygenate-degrading activity in the environment is impacted by cobalaminsynthesizing bacteria. The possibility clearly also exists that some attempts to isolate MTBE- and TBA-metabolizing bacteria in the past have not fully addressed the

22

M. Hyman

nutritional needs of these microorganisms by using media with insufficient cobalt or cobalamin.

1.17

Compound Specific Isotope Analysis (CSIA) of Aerobic Ether Oxygenate Biodegradation

Compound-specific isotope analysis (CSIA) is a useful tool for determining both the extent and mechanism of contaminant biodegradation. To facilitate the interpretation of CSIA data from field samples, laboratory studies are frequently conducted with mixed or pure microbial cultures where the enzymes, pathways, and microorganisms involved in the biodegradation process are well defined. In all cases it appears that monooxygenases are responsible for initiating the aerobic oxidation of MTBE, and this step is the primary cause of isotope fractionation observed during MTBE biodegradation. Consequently, CSIA studies with model aerobic MTBE-degrading strains have provided useful information about the types of monooxygenases involved in these processes. The first report of carbon fractionation during MTBE biodegradation examined microcosms containing aquifer materials, and similar final shifts in δ13C values (5.1–6.9‰) for MTBE were detected after >90% of the MTBE was consumed in microcosms incubated with and without 3-methylpentane as a stimulant for MTBE co-oxidation (Hunkeler et al. 2001). Plots of δ13C against ln of the fraction of MTBE remaining were linear and were used to determine carbon enrichment factors (εC) ranging from 1.52 to 1.97‰. Both the 13/12C and 2/1H fractionation associated with MTBE oxidation by pure cultures of M. petroleiphilum PM1 were subsequently determined, and εC and εH values were estimated at 2.0 to 2.4 ‰ and 33 to 37 ‰, respectively (Gray et al. 2002). Plots of Δδ2H versus Δδ13C were subsequently introduced (Kuder et al. 2005; Zwank et al. 2005), and at least initially, this two-dimensional approach suggested that aerobic and anaerobic MTBE degradation processes could be easily distinguished as anaerobic processes tended to exhibit small changes in δ2H and large changes in δ13C, whereas aerobic biodegradation processes tended to show the opposite effects. However, in a study of MTBE oxidation by A. tertiaricarbonis L108 and R. ruber IFP 2001, the εC value for MTBE (< 1 ‰) was shown to be much lower than for M. petroleiphilum PM1 and the closely related strain Methylibium R8, while the εH value was negligible and fell with the range observed for anaerobic MTBE degradation (Rosell et al. 2007). The low εH value for MTBE oxidation by strain L108 has been used to demonstrate the dominant activity of strain L108 and related strains in MTBE oxidation in either artificial mixed cultures of strain L018 and M. petroleiphilum PM1 (Rosell et al. 2010) or naturally occurring microbial communities supporting aerobic MTBE biodegradation (Jechalke et al. 2011). To date the most comprehensive analysis of εC and εH values for ether oxygenate degradation has demonstrated that CSIA of MTBE oxidation reinforces many of the physiological and molecular studies of bacteria to degrade MTBE (Rosell et al. 2012). While this and other CSIA studies have also addressed ETBE and TAME

2.50

AlkB-alkane hydroxylase

Unknown

Mycobacterium austroafricanum IFP 2012 Mycobacterium vaccae JOB5

ns not significant, CI confidence interval

2.64

MdpA-alkane hydroxylase

Methylibium sp. R8

1.4 2.0 to 2.4 2.3

Monooxygenase EthB cytochrome P450 EthB cytochrome P450 Tetrahydrofuran monooxygenase AlkB alkane hydroxylase MdpA-alkane hydroxylase

Strain Aquincola tertiaricarbonis L108 Rhodococcus ruber IFP 2001 Pseudonocardia tetrahydrofuranoxydans K1 Pseudomonas putida GPo1 Methylibium petroleiphilum PM1

εC (0/00) 0.48 0.28 2.3

0.04

0.08

0.1

0.1 0.1–0.3

95% CI (0/00) 0.05 0.06 0.2

4.2

ns

11 33 to 37 40

εH (0/00) ns ns 100

Table 1 Summary of carbon (EC) and hydrogen (EH) enrichment factors for CSIA studies of MTBE oxidation

0.9

+1

4

2 4–5

95% CI (0/00) 0.2 +5 10

Rosell et al. (2012)

Bastida et al. (2010) Rosell et al. (2012)

Reference Rosell et al. (2007) Rosell et al. (2007) McKelvie et al. (2009) Rosell et al. (2012) Gray et al. (2002)

Aerobic Degradation of Gasoline Ether Oxygenates 23

24

M. Hyman

biodegradation, only the data for MTBE are summarized in Table 1. This summary demonstrates that strains such as A. tertiaricarbonis L108 and Rhodococcus ruber IFP that utilize eth-gene-encoded cytochrome P450s to oxidize MTBE consistently exhibit both very low εC values (< 1 ‰) and nonsignificant εH values. In contrast, Pseudonocardia tetrahydrofuranoxidans K1, which utilizes tetrahydrofuran monooxygenase to initiate MTBE oxidation, exhibits both a high εC value ( 2.3‰) and the highest εH value so far detected for MTBE oxidation ( 100‰). The remaining organisms listed in Table 1 have all been suggested, with varying degrees of confidence, to utilize AlkB-like alkane hydroxylases to initiate MTBE oxidation. Despite this, there are also trends within this group. For example, MTBE oxidation by P. putida GPo1 exhibits the lowest εC value ( 1.4‰). In contrast, MTBE oxidation by both Methylibium strains PM1 and R8 consistently exhibits high εC values ( 2.0 to 2.4‰) and the highest εH values ( 33 to 40‰). Finally, MTBE oxidation by the propanotrophic Mycobacterium strains JOB5 and IFP 2012 consistently exhibits the highest εC values ( 2.5 to 2.64‰) and the lowest εH values (ns to 4.2‰). While these results clearly support the suggestion that physiologically and phylogenetically related organisms such as the two MTBE-metabolizing Methylibium strains or the two propanotrophic Mycobacterium strains employ similar enzymes to oxidize MTBE, there is currently insufficient data to determine whether these two seemingly different enzymes are AlkB-like monooxygenase, especially as the εC and εH values they exhibit are considerably different to these values exhibited by the well-characterized form of this enzyme in P. putida GPo1.

1.18

Research Needs

Like this review, the suggested areas for future research into aerobic ether oxygenate biodegradation focus on the physiology, enzymology, and pathways involved in these processes. In the case of MTBE co-oxidation, it seems likely that additional organisms with different monooxygenases to those already described still remain to be identified. As was illustrated by studies of MTBE oxidation by n-alkane-grown P. putida GPo1, negative results obtained using low concentrations of MTBE (Steffan et al. 1997) may be misleading due to the high Ks values for MTBE oxidation (Smith and Hyman 2004). While identifying additional enzymes with low affinities for MTBE may not expand our understanding of the fate of ether oxygenates in the environment where these compounds are typically present at low concentrations, studies with new organisms and enzymes could certainly be helpful for CSIA studies that have started to consider the different catalytic mechanisms of MTBE-oxidizing monooxygenases (Rosell et al. 2012). In the case of microbial cometabolism by propanotrophic actinomycetes, there are clearly two areas that require further study. First, additional research is required to define the final metabolic fate of 2HIBA and the role, if any, of 2HIBAL in MTBE oxidation. The role of 2HIBAL is of interest not only because it is a likely but as yet undetected intermediate but also because this compound is also potentially involved in MTBE metabolism by organisms such as A. tertiaricarbonis L108. As 2HIBA is

Aerobic Degradation of Gasoline Ether Oxygenates

25

also of great significance to the growth of strain L108 on MTBE, understanding the fate of this intermediate in propanotrophic actinomycetes may help explain why some of these strains grow on TBA while others do not. Second, the potential role of AlkB-like alkane hydroxylases in MTBE and TBA oxidation by these organisms also needs to be clarified. As alkane-oxidizing bacteria are well known to express multiple monooxygenases in response to alkane growth substrates, molecular approaches that focus on a single candidate enzyme or gene are unsatisfactory, and genome-based analyses should be used to answer this question. Likewise, genomeenabled approaches such as activity-based protein profiling (Bennett et al. 2016) that focus on catalytically active enzymes rather than transcriptional responses may also provide valuable insights into this issue. Finally, while the roles of many key enzymes involved in MTBE metabolism have been clearly established in A. tertiaricarbonis L108, this is less true for M. petroleiphilum PM1. In this organism the regulation and role of MdpA in MTBE oxidation need to be more clearly defined, as does the role of the protein products (MpeB0532-MpeB0535) of several currently unassigned genes which are highly upregulated in MTBE-grown cells (Hristova et al. 2007). Given the several unusual and interesting activities that have already been revealed during biochemical studies with enzymes from MTBE-metabolizing strains, these genes and their protein products may also be involved in equally unexpected and novel reactions.

References Alexander M (1973) Nonbiodegradable and other recalcitrant molecules. Biotechnol Bioeng 15:611–647 Aslett D, Haas J, Hyman M (2011) Identification of tertiary butyl alcohol (TBA)-utilizing organisms in BioGAC reactors using 13C-DNA stable isotope probing. Biodegradation 22:961–972 Auffret M, Labbé D, Thouand G, Greer CW, Fayolle-Guichard F (2009) Degradation of mixtures of hydrocarbons, gasoline and diesel oil additives by Rhodococcus aetherivorans and Rhodococcus wratislaviensis. Appl Environ Microbiol 75:7774–7782 Bastida F, Rosell M, Franchini AG, Seifert J, Finsterbusch S, Jehmlich N, Jechalke S, von Bergen M, Richnow HH (2010) Elucidating MTBE degradation in a mixed consortium using a multidisciplinary approach. FEMS Microbiol Ecol 73:370–384 Belhaj A, Desnoues N, Elmerich C (2002) Alkane biodegradation in Pseudomonas aeruginosa strains isolated from a polluted zone: Identification of alkB and alkB-related genes. Res Microbiol 153:339–344 Bennett K, Sadler NC, Wright AT, Yeager C, Hyman MR (2016) Activity-based protein profiling of ammonia monooxygenase in Nitrosomonas europaea. Appl Environ Microbiol 82:2270–2279 Béguin P, Chauvaux S, Miras I, François A, Fayolle F, Monot F (2003) Genes involved in the degradation of ether fuels by bacteria of the Mycobacterium/Rhodococcus group. Oil Gas Sci Technol 58:489–495 Bravo AL, Sigala JC, Borgne SL, Morales M (2015) Expression of an alkane hydroxylase (alkb) gene and methyl-tert-butyl ether co-metabolic oxidation in Pseudomonas citronellolis. Biotechnol Lett 37:807–814 Chauvaux S, Chevalier F, Le Dantec C, Fayolle F, Miras I, Kunst F, Béguin P (2001) Cloning of a genetically unstable cytochrome P-450 gene cluster involved in degradation of the pollutant ethyl tert-butyl ether by Rhodococcus ruber. J Bacteriol 183:6551–6557

26

M. Hyman

Curry S, Ciuffetti L, Hyman M (1996) Inhibition of growth of a Graphium sp. on gaseous n-alkanes by gaseous n-alkynes and n-alkenes. Appl Environ Microbiol 62:2198–2200 de Klerk H, van der Linden AS (1974) Bacterial degradation of cyclohexane. Participation of a co-oxidation reaction. Antonie Van Leeuwenhoek 40:7–15 Deeb RA, Hu H-Y, Hanson JR, Scow KM, Alvarez-Cohen L (2001) Substrate interactions in BTEX and MTBE mixtures by an MTBE-degrading isolate. Environ Sci Technol 35:312–317 Degnan PH, Taga ME, Goodman AL (2014) Vitamin B12 as a modulator of gut microbial ecology. Cell Metab 20:769–778 Ensign SA (1996) Aliphatic and chlorinated alkenes and epoxides as inducers of alkene monooxygenase and epoxidase activities in Xanthobacter strain Py2. Appl Environ Microbiol 62:61–66 Fan F, Germann MW, Gadda G (2006) Mechanistic studies of choline oxidase with betaine aldehyde and its isosteric analogue, 3-3dimethylbutyraldehyde. Biochemistry 45:1979–1986 Fayolle F, Hernandez G, Roux FL, Vandecasteele J-P (1998) Isolation of two aerobic bacterial strains that degrade efficiently ethyl t-butyl ether (ETBE). Biotechnol Lett 20:283–286 Ferreira NL, Maciel H, Mathis H, Monot F, Fayolle-Guichard F, Greer CW (2006a) Isolation and characterization of a new Mycobacterium austroafricanum strain, IFP 2015, growing on MTBE. Appl Microbiol Biotechnol 70:358–365 Ferreira NL, Labbé D, Monot F, Fayolle-Guichard F, Greer CW (2006b) Genes involved in the methyl tert-butyl ether (MTBE) metabolic pathway of Mycobacterium austroafricanum IFP 2012. Microbiology 152:1361–1374 Ferreira NL, Mathis H, Labbé D, Monot F, Greer CW, Fayolle-Guichard F (2007) N-alkane assimilation and tert-butyl alcohol (TBA) oxidation capacity in Mycobacterium austroafricanum strains. Appl Microbiol Biotechnol 75:909–919 Fournier D, Hawari J, Halasz A, Streger SH, McClay KR, Masuda H, Hatzinger PB (2009) Aerobic biodegradation of N-nitrosodimethylamine by the propanotroph Rhodococcus ruber ENV425. Appl Environ Microbiol 75:5088–5093 François A, Mathis H, Godefroy D, Piveteau P, Fayolle F, Monot F (2002) Biodegradation of methyl tert-butyl ether and other fuel oxygenates by a new strain, Mycobacterium austroafricanum IFP2012. Appl Environ Microbiol 68:2754–2762 François A, Garnier L, Mathis H, Fayolle F, Monot F (2003) Roles of tert-butyl formate, tert-butyl alcohol and acetone in the regulation of methyl tert-butyl ether degradation by Mycobacterium austroafricanum IFP 2012. Appl Microbiol Biotechnol 62:256–262 Garnier PM, Auria R, Augur C, Revah S (1999) Cometabolic biodegradation of methyl t-butyl ether by Pseudomonas aeruginosa grown on pentane. Appl Microbiol Biotechnol 51:498–503 Garnier PM, Auria R, Augur C, Revah S (2000) Cometabolic biodegradation of methyl tert-butyl ether by a soil consortium: effect of components present in gasoline. J Gen Appl Microbiol 46:79–84 Gedalanga PB, Pornwongthing P, Mora R, Chiang S-Y D, Baldwin B, Ogles D, Mahendra S (2014) Identification of biomarker genes to predict biodegradation of 1,4-dioxane. Appl Environ Microbiol 80:3209–3218 Goodfellow M, Jones AL, Maldonado LA, Salanitro J (2004) Rhodococcus aetherivorans sp. nov., A new species that contains methyl t-butyl ether-degrading actinomycetes. Syst Appl Microbiol 27:61–65 Gray JR, Lacrampe-Coulome G, Gandhi D, Scow KM, Wilson RD, Mackay DM, Lollar BS (2002) Carbon and hydrogen isotopic fractionation during biodegradation of methyl tert-butyl ether. Environ Sci Technol 36:1931–1938 Hanson JR, Ackerman CE, Scow KM (1999) Biodegradation of methyl tert-butyl ether by a bacterial pure culture. Appl Environ Microbiol 65:4788–4792 Hardison LK, Curry SS, Ciuffetti LM, Hyman MR (1997) Metabolism of diethyl ether and cometabolism of methyl tert-butyl ether by a filamentous fungus, a Graphium sp. Appl Environ Microbiol 63:3059–3067

Aerobic Degradation of Gasoline Ether Oxygenates

27

Hatzinger PB, McClay K, Vainberg S, Tugusheva M, Condee CW, Steffan RJ (2001) Biodegradation of methyl tert-butyl ether by a pure bacterial culture. Appl Environ Microbiol 67:5601–5607 Hernandez G, Francois A, Piveteau P, Fayolle F, Monot F (2007) Method for treating bacterial effluent containing at least Gordonia terrae CIP-I-2194 ether. US Patent 7,166,457 B1 Hernandez-Perez G, Fayolle F, Vandecasteele J-P (2001) Biodegradation of ethyl t-butyl ether (ETBE), methyl t-butyl ether (MTBE) and t-amyl ether (TAME) by Gordonia terrae. Appl Microbiol Biotechnol 55:117–121 Hofrichter M, Ullrich R (2014) Oxidations catalyzed by fungal peroxygenases. Curr Opin Chem Biol 19:116–125 House AJ, Hyman MR (2010) Effects of gasoline components on MTBE and TBA cometabolism by Mycobacterium austroafricanum JOB5. Biodegradation 21:525–541 Hristova KR, Schmidt R, Chakicherla AY, Legler TC, Wu J, Chain PS, Scow KM, Kane SR (2007) Comparative transcriptome analysis of Methylibium petroleiphilum PM1 exposed to the fuel oxygenates methyl tert-butyl ether and ethanol. Appl Environ Microbiol 73:7347–7357 Hunkeler D, Butler BJ, Aravena R, Barker JF (2001) Monitoring biodegradation of methyl tertbutyl ether (MTBE) using compound specific isotope analysis. Environ Sci Technol 35:676–681 Inoue D, Tsunoda T, Sawada K, Yamamoto N, Saito Y, Sei K, Ike M (2016) 1,4-dioxane degradation potential of members of the genera Pseudonocardia and Rhodococcus. Biodegradation 27:277–286 ITRC (2005) Overview of groundwater remediation technologies for MTBE and TBA. MTBE-1. Interstate Technology and Regulatory Council, MTBE and Other Fuel oxygenates Team. Available at http://www.itrcweb.org Jechalke S, Rosell M, Martínez-Lavanchy PM, Pérez-Leiva P, Rohwerder T, Vogt C, Richnow HH (2011) Linking low-level stable isotope fractionation to expression of the cytochrome P450 monooxygenase-encoding ethB gene for elucidation of methyl tert-butyl ether biodegradation in aerated treatment pond systems. Appl Environ Microbiol 77:1086–1096 Johnson EL, Hyman MR (2006) Propane and n-butane oxidation by Pseudomonas putida GPo1. Appl Environ Microbiol 72:950–952 Johnson EL, Smith CA, O’Reilly KT, Hyman MR (2004) Induction of methyl tertiary butyl ether (MTBE)-oxidizing activity in Mycobacterium vaccae JOB5 by MTBE. Appl Environ Microbiol 70:1023–1030 Joshi G, Schmidt R, Scow KM, Denison MS, Hristova KR (2015) gen mdpC plays a regulatory role in the methyl tert-butyl ether degradation pathway of Methylibium petroleiphilum strain PM1. FEMS Microbiol Lett 362:1–7 Joshi G, Schmidt R, Km S, Denison MS, Hristova KR (2016) Effect of benzene and ethylbenzene on the transcription of methyl tert-butyl ether degradation genes of Methylibium petroleiphilum PM1. Microbiology 162:1563–1571 Kane SR, Chakicherla AY, Chain PSG, Schmidt R, Shin MW, Legler TC, Scow KM, Larimer FW, Lucas SM, Richardson PM, Hristova KR (2007) Whole-genome analysis of the methyl-tertbutyl ether-degrading beta-proteobacterium Methylibium petroleiphilum PM1. J Bacteriol 189:1931–1945 Katapodis AG, Wimalesena K, Lee J, May SW (1984) Mechanistic studies on non-heme iron monooxygenase catalysis: Epoxidation, aldehyde formation, and demethylation by the ω-hydroxylation system of Pseudomonas oleovorans. J Am Chem Soc 110:7928–7935 Kharoune M, Kharoune L, Lebault JM, Pauss A (2001) Isolation and characterization of two aerobic bacterial strains that completely degrade ethyl tert-butyl ether (ETBE). Appl Microbiol Biotechnol 55:348–353 Kim Y-H, Engesser K-H, Kim S-J (2007) Physiological, numerical and molecular characterization of alkyl ether-utilizing rhodococci. Environ Microbiol 9:1497–1510 Kinne M, Poraj-Kobielska M, Ralph SA, Ullrich R, Hofrichter M, Hammel KE (2009) Oxidative cleavage of diverse ethers by an extracellular fungal peroxygenase. J Biol Chem 284:29343–29349

28

M. Hyman

Kohlweyer U, Thiemer B, Schräder T, Andreesen JR (2000) Tetrahydrofuran degradation by a newly isolated culture of Pseudonocardia sp. strain K1. FEMS Microbiol Lett 186:301–306 Kottegoda S, Waligora E, Hyman MR (2015) Metabolism of 2-methylpropane (isobutylene) by the aerobic bacterium Mycobacterium sp. strain ELW1. Appl Environ Microbiol 81:1966–1976 Kuder T, Wilson JT, Kaiser P, Kolhatkar R, Philp P, Allen J (2005) Enrichment of stable carbon and hydrogen isotopes during anaerobic biodegradation of MTBE: microcosm and field evidence. Environ Sci Technol 39:213–220 Kurteva-Yaneva N, Zahn M, Weichler M-T, Starke R, Harms H, Müller RH, Sträter N, Rohwerder T (2015) Structural basis of the stereospecificity of bacterial B12-dependent 2-hydroxyisobutyrylCoA mutase. J Biol Chem 290:9727–9737 Kusano M, Sakai Y, Kato N, Yoshomoto H, Sone H, Tani Y (1998) Hemiacetal dehydrogenase activity in alcohol dehydrogenases in Saccharomyces cerevisiae. Biosci Biotechnol Biochem 62:1956–1961 Le Digabel Y, Demanéche S, Benoit Y, Vogel TM, Fayolle-Guichard F (2013) Ethyl tert-butyl ether (ETBE) biodegradation by a syntrophic association of Rhodococcus sp. IFP 2042 and Bradyrhizobium sp. IFP 2049 isolated from a polluted aquifer. Appl Microbiol Biotechnol 97:10531–10539 Le Digabel Y, Demanéche S, Benoit Y, Fayolle-Guichard F, Vogel TM (2014) Ethyl tert-butyl ether (ETBE)-degrading microbial communities in enrichments from polluted environments. J Hazard Mater 279:502–510 Lechner U, Brodkorb D, Geyer R, Hause G, Härtig C, Auling G, Fayolle-Guichard F, Piveteau P, Müller RH, Rohwerder T (2007) Aquincola tertiaricarbonis gen. nov., sp. nov., a tertiary butyl moiety-degrading bacterium. Int J Syst Evol Microbiol 57:1295–1303 Liu CY, Speitel GE, Georgiou G (2001) Kinetics of methyl t-butyl ether cometabolism at low concentrations by pure cultures of butane-degrading bacteria. Appl Environ Microbiol 67:2197–2201 Magaña-Reyes M, Morales M, Revah S (2005) Methyl tert-butyl ether and tert-butyl alcohol degradation by Fusarium solani. Biotechnol Lett 27:1797–1801 Malandain C, Fayolle-Guichard F, Vogel TM (2010) Cytochrome P450-mediated degradation of fuel oxygenates by environmental isolates. FEMS Microbiol Ecol 72:289–296 Malone VF, Chastain AJ, Ohlsson JT, Poneleit LS, Nemecek-Marshall M, Fall R (1999) Characterization of a Pseudomonas putida allylic alcohol dehydrogenase induced by growth on 2-methyl-3-buten-2-ol. Appl Environ Microbiol 65:2622–2630 Masuda H, McClay K, Steffan RJ, Zylstra GJ (2012a) Biodegradation of tetrahydrofuran and 1,4-dioxane by soluble diiron monooxygenase in Pseudonocardia sp. strain ENV478. J Mol Microbiol Biotechnol 22:312–316 Masuda H, McClay K, Steffan RJ, Zylstra GJ (2012b) Characterization of three propane-inducible oxygenases in Mycobacterium sp. strain ENV421. Lett Appl Microbiol 55:175–181 McKelvie JR, Hyman MR, Elsner M, Smith C, Aslett DM, Lacrampe-Couloume G, SherwoodLolar (2009) Isotopic fractionation of methyl tert-butyl ether suggests different initial reactions mechanism during aerobic biodegradation. Environ Sci Technol 43:2793–2799 Mo K, Lora CO, Wanken AE, Javanmardian M, Yang M, Kulpa CF (1997) Biodegradation of methyl t-butyl ether pure bacterial cultures. Appl Microbiol Biotechnol 47:69–72 Morales M, Velázquez E, Jan J, Revah S, González U, Razo-Flores E (2004) Methyl tert-butyl ether biodegradation by microbial consortia obtained from soil samples of gasoline-polluted sites in Mexico. Biotechnol Lett 26:269–275 Morales M, Nava V, Velásquez E, Razo-Flores E, Revah S (2009) Mineralization of methyl tertbutyl ether and other gasoline oxygenates by Pseudomonads using short chain n-alkanes as growth source. Biodegradation 20:271–280 Müller RH, Rohwerder T, Harms H (2007) Carbon conversion efficiency and limits or productive degradation of methyl tert-butyl ether and related compounds. Appl Environ Microbiol 73:1783–1791

Aerobic Degradation of Gasoline Ether Oxygenates

29

Müller RH, Rohwerder T, Harms H (2008) Degradation of fuel oxygenates and their main intermediates by Aquincola tertiaricarbonis L108. Microbiology 154:1414–1421 Nagy I, Compernolle F, Ghys K, Vanderleyden J, De Mot R (1995a) A single cytochrome P-450 system is involved in degradation of the herbicides EPTC (S-ethyl dithiocarbamate) and atrazine by Rhodococcus sp. strain NI86/21. Appl Environ Microbiol 61:2056–2060 Nagy I, Schoofs G, Compernolle F, Proost P, Vanderleyden J, De Mot R (1995b) Degradation of the thiocarbamate herbicide EPTC (S-ethyl dipropylcarbamothioate) and biosafening by Rhodococcus sp. strain NI86/21 involve an inducible cytochrome P-450 system and aldehyde dehydrogenase. J Bacteriol 177:676–687 Nakatsu CH, Hristova K, Hanada S, Meng X-Y, Hanson JR, Scow KM, Kamagata Y (2006) Methylibium petroleiphilum gen nov., sp. nov., a novel methyl tert-butyl ether-degrading methylotroph of the Betaproteobacteria. Int J Syst Evol Microbiol 56:983–989 Owens CR, Karceski JK, Mattes TE (2009) Gaseous alkene biotransformation and enantioselective epoxyalkane formation by Nocardioides sp. strain JS614. Appl Microbiol Biotechnol 84:685–692 Peter S, Kinne M, Wang X, Ullrich R, Kayser G, Groves JT, Hofrichter M (2011) Selective hydroxylation of alkanes by an extracellular fungal peroxygenase. FEBS J 278:3567–3675 Piveteau P, Fayolle F, Vandecasteele J-P, Monot F (2001) Biodegradation of tert-butyl alcohol and related xenobiotics by a methylotrophic bacterial isolate. Appl Microbiol Biotechnol 55:369–373 Przybylski D, Rohwerder T, Dilbner C, Maskow T, Harms H, Müller RH (2015) Exploiting mixtures of H2, CO2, and O2 for improved production of methacrylate precursor 2-hydroxyisobutyric acid by engineered Cupriavidus necator strains. Appl Microbiol Biotechnol 99:2131–2145 Rohde M-T, Tischer S, Harms H, Rohwerder T (2016) Production of 2-hydroxyisobutyric acid from methanol by Methylobacterium extorquens AM1 expressing (R)-3-hydroxybutyryl-CoA isomerizing enzymes. Appl Environ Microbiol. doi:10.1128/AEM.02622-16 Rohwerder T, Müller RH (2010) Biosynthesis of 2-hydroxyisobutyric acid (2-HIBA) from renewable carbon. Microb Cell Factories 9:13 Rohwerder T, Breuer U, Benndorf D, Lechner U, Müller RH (2006) The alkyl tert-butyl ether intermediate 2-hydroxyisobutyrate is degraded by a novel cobalamin-dependent mutase pathway. Appl Environ Microbiol 72:4128–4135 Rosell M, Barceló D, Rohwerder T, Breuer U, Gehre M, Richnow HH (2007) Variations in 13C/12C and D/H enrichment factors of aerobic bacterial fuel oxygenate degradation. Environ Sci Technol 41:2036–2043 Rosell M, Finsterbusch S, Jechalke S, Hübschmann T, Vogt C, Richnow HH (2010) Evaluation of the effects of low oxygen concentration on stable isotope fractionation during aerobic MTBE biodegradation. Environ Sci Technol 44:319–315 Rosell M, Gonzales-Olmos R, Rohwerder T, Rusevova K, Georgi A, Kopinke F-D, Richnow HH (2012) Critical evaluation of the 2D-CSIA scheme for distinguishing fuel oxygenate degradation reaction mechanisms. Environ Sci Technol 46:4757–4766 Salanitro JP, Diaz LA, Williams MP, Wisniewski HL (1994) Isolation of a bacterial culture that degrades methyl t-butyl ether. Appl Environ Microbiol 60:2593–2596 Sales CM, Grostern A, Parales JV, Parales RE, Alvarez-Cohen L (2013) Oxidation of the cyclic ethers 1,4-dioxane and tetrahydrofuran by a monooxygenase in two Pseudonocardia species. Appl Environ Microbiol 79:7702–7708 Schäfer F, Breuer U, Benndorf D, von Bergen M, Harms H, Müller RH (2007) Growth of Aquincola tertiaricarbonis L108 on tert-butyl alcohol leads to the induction of a phthalate dioxygenaserelated protein and its associated oxidoreductase subunit. Eng Life Sci 7:512–519 Schäfer F, Muzica L, Schuster J, Treuter N, Rosell M, Harms H, Müller RH, Rohwerder T (2011) Formation of alkenes via degradation of tert-alkyl ethers and alcohols by Aquincola tertiaricarbonis L108 and Methylibium spp. Appl Environ Microbiol 77:5981–5987

30

M. Hyman

Schäfer F, Schuster J, Wuurz B, Härtig C, Harms H, Müller RH, Rohwerder T (2012) Synthesis of short-chain diols and unsaturated alcohols from secondary alcohol substrates by the Rieske nonheme mononuclear iron oxygenase MdpJ. Appl Environ Microbiol 78:6280–6284 Schmidt R, Battaglia V, Scow K, Kane S, Hristova KR (2008) Involvement of a novel enzyme, MdpA, in methyl-tert-butyl ether degradation in Methylibium petroleiphilum PM1. Appl Environ Microbiol 74:6631–6638 Schuster J, Schäfer F, Hübler N, Brandt A, Rosell M, Härtig C, Harms H, Müller RH, Rohwerder T (2012) Bacterial degradation of tert-amyl alcohol proceeds via hemiterpene 2-methyl-3-buten2-ol by employing the tertiary alcohol desaturase function of the Rieske nonheme mononuclear iron oxygenase MdpJ. J Bacteriol 194:972–981 Schuster J, Purswani J, Breuer U, Pozo C, Harms H, Müller RH, Rohwerder T (2013) Constitutive expression of the cytochrome P450 ethABCD monooxygenase system enables degradation of synthetic dialkyl ethers in Aquincola tertiaricarbonis L108. Appl Environ Microbiol 79:2321–2327 Sharp JO, Sales CM, Alvarez-Cohen L (2010) Functional characterization of propane-enhanced Nnitrosodimethylamine degradation by two actinomycetales. Biotechnol Bioeng 107:924–932 Skinner KM, Martinez-Prado A, Hyman MR, Williamson KJ, Ciuffetti LM (2008) Pathway, inhibition and regulation of methyl tertiary butyl ether oxidation in a filamentous fungus, Graphium sp. Appl Microbiol Biotechnol 77:1359–1365 Skinner K, Cuifetti L, Hyman M (2009) Metabolism and cometabolism of cyclic ethers by a filamentous fungus, a Graphium sp. Appl Environ Microbiol 75:5514–5522 Smith CA, Hyman MR (2004) Oxidation of methyl tert-butyl ether by alkane hydroxylase in dicyclopropylketone-induced and n-octane-grown Pseudomonas putida GPo1. Appl Environ Microbiol 70:4544–4550 Smith CA, Hyman MR (2010) Oxidation of gasoline oxygenates by closely related non-haem-iron alkane hydroxylases in Pseudomonas mendocina KR1 and other n-octane-utilizing Pseudomonas strains. Environ Microbiol Rep 2:426–432 Smith CA, O’Reilly KT, Hyman MR (2003a) Characterization of the initial reactions during the cometabolic oxidation of methyl tert-butyl ether by propane-grown Mycobacterium vaccae JOB5. Appl Environ Microbiol 69:796–804 Smith CA, O’Reilly KT, Hyman MR (2003b) Cometabolism of methyl tertiary butyl ether and gaseous n-alkanes by Pseudomonas mendocina KR-1 grown on C5-C8 n-alkanes. Appl Environ Microbiol 69:7385–7394 Smits THM, Witholt B, van Beilen JB (2003) Functional characterization of genes involved in alkane oxidation by Pseudomonas aeruginosa. Antonie Van Leeuwenhoek 84:193–200 Steffan RJ, McClay K, Vainberg S, Condee CW, Zhang D (1997) Biodegradation of the gasoline oxygenates methyl tert-butyl ether, ethyl tert-butyl ether, and tert-amyl methyl ether by propane-oxidizing bacteria. Appl Environ Microbiol 63:4216–4222 Streger SH, Vainberg S, Dong H, Hatzinger PB (2002) Enhancing transport of Hydrogenophaga flava ENV735 for bioaugmentation of aquifers contaminated with methyl tert-butyl ether. Appl Environ Microbiol 68:5571–5579 Szabó Z, Gyula P, Roboka H, Bató E, Gálik B, Pach P, Pekker P, Papp I, Bihari Z (2015) Draft genome sequence of Methylibium sp. strain T29, a novel fuel oxygenate-degrading bacterial isolate from Hungary. Stand Genomic Sci. doi:10.1186/s40793-015-0023-z Thiemer B, Andreesen JR, Schräder T (2003) Cloning and characterization of a gene cluster involved in tetrahydrofuran degradation in Pseudonocardia sp. strain K1. Arch Microbiol 179:266–277 Tomasi I, Artaud I, Bertheau Y, Mansuy D (1995) Metabolism of polychlorinated phenols by Pseudomonas cepacia AC1100: determination of the first two steps and specific inhibitory effect of methimazole. J Bacteriol 177:307–311 Trippe KM, Wolpert TJ, Hyman MR, Ciuffetti LM (2014) RNAi silencing of a cytochrome P450 monooxygenase disrupts the ability of the filamentous fungus, Graphium sp., to grow on shortchain gaseous alkanes and ethers. Biodegradation 25:137–151

Aerobic Degradation of Gasoline Ether Oxygenates

31

Vainberg S, McClay K, Masuda H, Root D, Condee C, Zylstra GJ, Steffan RJ (2006) Biodegradation of ether pollutants by Pseudonocardia sp. ENV478. Appl Environ Microbiol 72:5218–5224 van Beilen JB, Funhoff EG (2007) Alkane hydroxylases involved in microbial alkane oxidation. Appl Microbiol Biotechnol 74:13–21 van Beilen JB, Kingma J, Witholt B (1994) Substrate specificity of the alkane hydroxylase system in Pseudomonas oleovorans GPo1. Enzym Microb Technol 16:904–911 van Beilen JB, Li Z, Duetz WA, Smits THM, Witholt B (2003) Diversity of alkane hydroxylase systems in the environment. Oil Gas Sci Technol 58:427–440 van Beilen JB, Smits THM, Roos FF, Brunner T, Balada SB, Röthlisberger M, Witholt B (2005) Identification of an amino acid position that determines the substrate range of integral membrane alkane hydroxylase. J Bacteriol 187:85–91 Vomberg A, Klinner U (2000) Distribution of alkB genes within n-alkane-degrading bacteria. J Appl Microbiol 89:339–348 Wang F, Zhou J, Li Z, Dong W, Hou Y, Cui Z (2015) Involvement of the cytochrome P450 system EthBAD in the N-deethoxymethylation of acetochlor by Rhodococcus sp. strain T3-1. Appl Environ Microbiol 81:2182–2188 Weichler M-T, Kurteva-Yaneva N, Przybylski D, Schuster J, Müller RH, Rohwerder T (2015) Thermophilic coenzyme B12-dependent acyl coenzyme A (CoA) mutase from Kyrpidia tusciae DSM 2912 preferentially catalyzes isomerization of (R)-3-hydroxybutyryl-CoA and 2-hydroxyisobutyryl-CoA. Appl Environ Microbiol 81:4564–4572 White GF, Russell NJ, Tidswell EC (1996) Bacterial scission of ether bonds. Microbiol Rev 60:216–232 Yamashita M, Tani A, Kawai F (2005) A new ether bond-splitting enzyme found in gram-positive polyethylene glycol 6000-utilizing bacterium, Pseudonocardia sp. strain K1. Appl Microbiol Biotechnol 66:174–179 Yaneva N, Schuster J, Schäfer F, Lede V, Przybylski D, Paproth T, Harms H, Müller RH, Rohwerder T (2012) Bacterial acyl-CoA mutase specifically catalyzes coenzyme B12-dependent isomerization of 2-hydroxyisobutyryl-CoA and (S)-3-hydroxybutyryl-CoA. J Biol Chem 287:15502–15511 Zaitsev GM, Uotila JS, Häggblom MM (2007) Biodegradation of methyl tert-butyl ether by coldadapted mixed and pure cultures. Appl Microbiol Biotechnol 74:1092–1102 Ziegler DM (2002) An overview of the mechanism, substrate specificities, and structure of FMOs. Drug Metab Rev 34:503–511 Zwank L, Berg M, Elsner M, Schmidt T, Schwarzenbach RP, Haderlein SB (2005) New evaluation scheme for two-dimensional isotope analysis to decipher biodegradation processes: application to groundwater contamination by MTBE. Environ Sci Technol 39:1018–1029

Genetic Features and Regulation of nAlkane Metabolism in Yeasts Ryouichi Fukuda and Akinori Ohta

Abstract

The yeasts Candida tropicalis, Candida maltosa, and Yarrowia lipolytica have an excellent ability to use n-alkanes as the sole carbon and energy source. Here, we summarize the current knowledge of the genetic features and regulation of n-alkane metabolism in these yeasts. The transcription of genes encoding the CYP52-family cytochromes P450 that catalyze the initial hydroxylation of n-alkanes has been shown to be activated when these yeasts are cultured in the presence of n-alkanes. In Y. lipolytica, the transcription of ALK1, the gene encoding P450, is activated by a complex composed of two basic helix-loophelix transcription activators Yas1p and Yas2p through a promoter element ARE1. This transcription is regulated by an Opi1-family transcriptional repressor Yas3p. In the absence of n-alkanes, Yas3p binds to Yas2p in the nucleus thereby repressing the transcription of ALK1. However, in the presence of n-alkanes, Yas3p is sequestered to the endoplasmic reticulum to derepress the transcription of the gene.

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Response to n-Alkanes in n-Alkane-Assimilating Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Mechanism of Transcriptional Activation of Genes Responsible for n-Alkane Degradation in Y. lipolytica . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1 Promoter Elements of the CYP52-Family P450 Gene Involved in n-Alkane Response

2 2 4 4

R. Fukuda (*) Department of Biotechnology, The University of Tokyo, Tokyo, Japan e-mail: [email protected] A. Ohta Department of Biological Chemistry, College of Bioscience and Biotechnology, Chubu University, Kasugai, Aichi, Japan e-mail: [email protected] # Springer International Publishing AG 2017 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-319-39782-5_24-1

1

2

R. Fukuda and A. Ohta

3.2 Transcription Activators Involved in n-Alkane Response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 3.3 Regulation of n-Alkane Metabolic Genes by the Opi1-Family Transcription Factor 5 3.4 Role of the Opi1-Family Proteins in Other Yeasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 4 Repression of Transcription of n-Alkane Metabolic Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

1

Introduction

A variety of microorganisms, including certain species of yeasts, has developed metabolic systems to assimilate n-alkanes containing 10–18 carbons as the sole carbon and energy source. The degradation pathway of n-alkanes in yeasts and the enzymes involved in it have been extensively studied in Candida tropicalis (Tanaka and Fukui 1989), Candida maltosa (Mauersberger et al. 1996), and Yarrowia lipolytica (Barth and Gaillardin 1996, 1997; Fickers et al. 2005; Fukuda 2013; Fukuda and Ohta 2013; Nicaud 2012) as described in chapter ▶ Enzymes for Aerobic Degradation of Alkanes in Yeasts' by Fukuda and Ohta. In these yeasts, n-alkanes are first hydroxylated to fatty alcohols in the endoplasmic reticulum (ER) by cytochromes P450 belonging to the CYP52 family (Figs. 2 and 4 of the chapter ▶ Enzymes for Aerobic Degradation of Alkanes in Yeasts by Fukuda and Ohta). Fatty alcohols are then oxidized to fatty aldehydes by fatty alcohol dehydrogenase (FADH) in the ER or by fatty alcohol oxidase (FAO) in the peroxisome. Fatty aldehydes are further oxidized to fatty acids by fatty aldehyde dehydrogenase (FALDH) in the ER or the peroxisome, and then finally activated to acyl-CoAs by acyl-CoA synthetase (ACS). These activated fatty acids are then utilized for lipid synthesis, or degraded in the peroxisome via β-oxidation. It is crucial for an organism to respond to and adapt rapidly to environmental changes for survival. In C. tropicalis, C. maltosa, and Y. lipolytica, the metabolism of n-alkanes was found to be regulated at the transcriptional level. The transcription of the genes encoding enzymes involved in the metabolism of n-alkanes was found to be activated in the presence of n-alkanes. The mechanism of the transcription regulation in these yeasts is a very interesting subject, particularly because the regulation of transcription by hydrophobic compounds in lower eukaryotes remains largely elusive. This chapter will focus on the molecular mechanisms of the transcriptional regulation of genes involved in the degradation of n-alkanes in these yeasts.

2

Response to n-Alkanes in n-Alkane-Assimilating Yeasts

Experiments conducted in the 1970s showed that the production of cytochromes P450 was induced when C. tropicalis was cultured in a medium containing n-tetradecane (Lebeault et al. 1971). Cloning and subsequent characterization of the genes in C. tropicalis revealed that this yeast has at least eight genes encoding the

Genetic Features and Regulation of n-Alkane Metabolism in Yeasts Fig. 1 Response to n-alkanes in n-alkane-assimilating yeasts. When n-alkaneassimilating yeasts are cultured in the medium containing n-alkane, the transcriptional induction of genes involved in the degradation of n-alkanes and the proliferation of the ER and the peroxisome, in which nalkanes are degraded, are observed

3

n-Alkane

Organelle proliferation

Induction of n-alkane metabolic genes

CH2OH CHO COOH

Nucleus

ER

Peroxisome

CYP52-family P450s, (ALK1–ALK7 and CYP52D2) and that the transcription of four of them (ALK1–ALK3 and ALK5) is activated by n-alkanes (Fig. 1) (Craft et al. 2003; Nelson 2009; Sanglard et al. 1987;Seghezzi et al. 1991, 1992). In C. maltosa, of the eight genes (ALK1–ALK8) encoding the CYP52-family P450s, the transcription of all except ALK4 was induced by n-alkanes (Ohkuma et al. 1991, 1995a). Y. lipolytica has 12 genes (ALK1–ALK12) encoding the CYP52-family P450s, transcription of nine of which (ALK1–ALK6, ALK9, ALK11, and ALK12) was induced by n-alkanes (Hirakawa et al. 2009; Iida et al. 1998, 2000; Iwama et al. 2016; Takai et al. 2012). The transcriptional induction of the P450 genes by n-alkanes was also observed in sophorolipid-producing yeast Candida bombicola (Van Bogaert et al. 2009). The transcription of genes involved in the degradation of n-alkane metabolites was also found to be induced in the presence of n-alkanes in Y. lipolytica (Fig. 4 of the chapter ▶ Enzymes for Aerobic Degradation of Alkanes in Yeasts by Fukuda and Ohta). In Y. lipolytica, the transcription of ADH1 and ADH3, encoding alcohol dehydrogenases, and FAO1, encoding a fatty alcohol oxidase, is upregulated in the presence of n-alkanes (Iwama et al. 2015). In addition, transcription of three (HFD1–HFD3) of the four genes (HFD1–HFD4) encoding fatty aldehyde dehydrogenases and FAA1 and FAT1 encoding ACSs was increased in the presence of nalkanes (Iwama et al. 2014; Tenagy et al. 2015). The transcription of PAT1 encoding a peroxisomal acetoacetyl-CoA thiolase involved in β-oxidation was also induced by n-alkanes (Yamagami et al. 2001). These results confirm that the transcription of genes important for the n-alkane degradation is activated in response to n-alkanes. Our transcriptome analysis in Y. lipolytica cells cultured in medium containing either glucose or n-decane suggested that the transcripts of approximately 500 genes were increased more than twofold in response to n-decane (our unpublished results). n-Alkanes have also been reported to induce the proliferation of the ER and peroxisome in the n-alkane-assimilating yeasts (Fig. 1) (Mauersberger et al. 1987; Osumi et al. 1974; Vogel et al. 1992). Proliferation of the peroxisome by fatty acids

4

R. Fukuda and A. Ohta

has been widely observed in various organisms, including C. tropicalis, Y. lipolytica, and Saccharomyces cerevisiae, and its mechanism has been extensively studied (Gurvitz and Rottensteiner 2006). In contrast, the mechanism underlying the proliferation of the ER by n-alkanes in n-alkane-assimilating yeasts remains poorly understood. Interestingly, overproduction of the CYP52-family P450 induced the proliferation of the ER in C. maltosa and S. cerevisiae (Ohkuma et al. 1995b; Schunck et al. 1991). Proliferation of the ER due to overproduction of various membrane proteins has been reported (Federovitch et al. 2005) and may be one of the quality control mechanisms of the ER to avoid over accumulation of proteins in the ER membrane.

3

Mechanism of Transcriptional Activation of Genes Responsible for n-Alkane Degradation in Y. lipolytica

The transcriptional induction of the P450 genes in response to n-alkane was first identified in yeasts of the Candida genus as described above. However, the molecular mechanism of its regulation remains unclear. This is largely due to the difficulty in obtaining mutant strains defective for transcriptional activation in the presence of n-alkanes, since C. tropicalis and C. maltosa are diploid or partial diploid yeasts, in which teleomorphs have not been found. Furthermore, the CUG codon has been shown to code for serine instead of leucine in C. tropicalis and C. maltosa, as well as in other n-alkane-assimilating yeasts phylogenetically close to them, including Candida albicans, Candida dubliniensis, Candida parapsilosis, Debaryomyces hansenii, Lodderomyces elongisporus, and Meyerozyma guilliermondii (Massey et al. 2003; Sugiyama et al. 1995; Ueda et al. 1994), and this poses an obstacle to the analysis of DNA-protein and/or protein-protein interaction using S. cerevisiae system. Y. lipolytica, on the other hand, has a teleomorph and a stable haploid and diploid life cycle, and genetic methods that permit isolation and characterization of mutants as well as molecular biology methods are well established in it (Barth and Gaillardin 1996, 1997). Among the yeasts that can assimilate n-alkanes, the genome sequences were determined initially in Y. lipolytica, D. hansenii (Dujon et al. 2004), and C. albicans (Jones et al. 2004), followed by C. tropicalis and others (Butler et al. 2009). As a result of these advantages, the mechanism of the transcriptional regulation by n-alkane has been elucidated in Y. lipolytica.

3.1

Promoter Elements of the CYP52-Family P450 Gene Involved in n-Alkane Response

The transcriptional regulation in response to n-alkanes has been investigated by studying ALK1 encoding the primary CYP52-family P450 in the assimilation of n-alkanes in Y. lipolytica (Iida et al. 1998, 2000; Iwama et al. 2016; Takai et al. 2012). The transcription of ALK1 was highly activated in the presence of n-alkanes

Genetic Features and Regulation of n-Alkane Metabolism in Yeasts

5

and the transcript level was found to be highest among the 12 ALK genes (Hirakawa et al. 2009). Promoter analysis of ALK1 led to the identification of a sequence named ARR1 (alkane responsive region 1), which is involved in the transcription activation (Sumita et al. 2002). ARR1 was found to contain two elements, ARE1 (alkane responsive element 1) and ARE2. In electrophoretic mobility shift assay, specific shift bands corresponding to both ARE1 and ARE2 could be identified with cellular extracts of Y. lipolytica cells cultured in n-alkane-containing medium. ARE1 contains a sequence similar to the E-box motif, the consensus sequence to which basic helix-loop-helix (bHLH) transcription factors interact (Murre et al. 1994), and was found to play a critical role in the transcriptional activation process (see below). Interestingly, ARE1-like sequences were found in the promoter regions of various genes involved in n-alkane metabolism (Yamagami et al. 2004). In contrast, the role of ARE2 is still unknown.

3.2

Transcription Activators Involved in n-Alkane Response

A gene YAS1 encoding a transcription factor that activates the ARE1-mediated transcription was identified by the analysis of a mutant defective in the transcriptional activation through ARE1 by n-alkanes and in the growth on n-alkanes (Yamagami et al. 2004). YAS1 encodes a bHLH transcription factor of 137 amino acids (Fig. 2). A deletion mutant of YAS1 showed defects in the transcription activation of ALK1 by n-alkanes. However, Yas1p did not bind to ARE1 in vitro. bHLH transcription factors generally form homo- or heterodimers through their HLH regions and interact with the E-box motif through the basic regions (Murre et al. 1994). Indeed, a gene named YAS2 encoding a 700-amino acid protein that contains a bHLH motif similar to that of Yas1 was identified from the genome database. This gene was found to be involved in the ARE1-mediated transcriptional activation (Fig. 2) (Endoh-Yamagami et al. 2007). The deletion mutant of YAS2 was also defective in the transcription induction of ALK1 by n-alkanes. Yas1p and Yas2p formed a complex in vitro and bound to ARE1 only when both proteins existed. Yas1p and Yas2p constitutively localized in the nucleus (Hirakawa et al. 2009; Yamagami et al. 2004). These results suggest that the complex of Yas1p and Yas2p binds to ARE1 and activates the transcription of ALK1 in response to n-alkane. Deletion mutants of YAS1 or YAS2 did not have the ability to grow on n-alkanes, indicating the importance of Yas1p-Yas2p complex in the ARE1mediated transcriptional activation.

3.3

Regulation of n-Alkane Metabolic Genes by the Opi1-Family Transcription Factor

The bHLH motifs of Yas1p and Yas2p show sequence similarities to Ino4p and Ino2p, respectively, of S. cerevisiae. In S. cerevisiae, these function as transcription activators regulating genes involved in phospholipid synthesis. In this yeast,

6

R. Fukuda and A. Ohta 1

304

Ino2p RID 1

bHLH 700

Yas2p

1

151

Ino4p bHLH 1

137

Yas1p PA binding domain 1

Opi1p

l-Yas3p

s-Yas3p

FFAT motif 404

Leu zipper

1

1

AID

727

422

Fig. 2 Schematic diagrams of bHLH transcription activators and Opi1-family proteins in S. cerevisiae and Y. lipolytica. The bHLH motifs, repressor interaction domains (RID), PA-binding domains, FFAT motifs, leucine zipper motifs, and activator interaction domains (AID) are indicated by grey boxes

transcription of phospholipid synthetic genes, including INO1 encoding inositol-3phosphate synthase, is activated in the absence of myo-inositol and repressed in the presence of it (Henry et al. 2012). A heterodimer of Ino2p and Ino4p constitutively binds to an upstream activating element, UASINO/ICRE (inositol choline responsive element), in the promoter regions of the target genes. The transcription repressor Opi1p binds through its activator interaction domain (AID) to the repressor interaction domain (RID) of Ino2p (Heyken et al. 2005). Loewen et al. proposed a model in which Opi1p is retained to the ER by binding to phosphatidic acid (PA) using its PA-binding domain and to an ER membrane-spanning protein Scs2p through its FFAT (two phenylalanine in an acidic tract) motif in the absence of myo-inositol. This leads to the activation of transcription of the phospholipid synthesis genes by the Ino2-Ino4 complex (Loewen et al. 2004). However, in the presence of myo-inositol, PA is utilized for the synthesis of phosphatidylinositol (PI). Opi1p is then released from the ER and transported to the nucleus where it binds to Ino2p to repress the transcription. Therefore, Opi1p is the key regulator controlling transcription of genes responsible for phospholipid biosynthesis in response to myo-inositol.

Genetic Features and Regulation of n-Alkane Metabolism in Yeasts

7

In the Y. lipolytica genome database, an ortholog of OPI1 was identified and named as YAS3 (Hirakawa et al. 2009). Two different transcripts with different transcription initiation sites were obtained from YAS3. These transcripts were predicted to encode a long form of Yas3p (l-Yas3p of 727 amino acids) and a short form of Yas3p (s-Yas3p of 422 amino acids) (Fig. 2). The AID, PA-binding domain, and leucine zipper domain found in Opi1p were conserved in both forms of Yas3p, but the FFAT motif was absent in Yas3p (Fig. 2). Whether these two forms of Yas3p have different functions is still not clear, but it has been shown that the s-Yas3p is sufficient to regulate the transcription of ALK1. The deletion mutant of YAS3 accumulated much more transcripts of ALK1 than the wild-type strain, even when cells were cultured in the medium containing glucose or glycerol. Yas3p interacted with Yas2p, but not with Yas1p. In line with this observation, a RID-like sequence motif was found in Yas2p (Fig. 2). In addition, while Yas3p was localized in the nucleus when cultured in the medium containing glucose, it was sequestered to the ER in the presence of n-alkanes. Among the 12 ALK genes, the transcription of ALK1, ALK2, ALK4, ALK6, ALK9, and ALK11 appeared to be regulated by the Yas1p-Yas2p-Yas3p system. In contrast, Yas3p was not involved in the transcriptional regulation of INO1 by myo-inositol. A question remained as to why Yas3p remained localized to the ER in the presence of n-alkanes. Yas3p was found to bind to PA and phosphoinositides (PIPs), particularly to phosphatidylinositol 4-phosphate (PI(4)P), in vitro, but not to n-alkanes (Kobayashi et al. 2013). In addition, the ARE1-mediated transcription was upregulated in mutants defective for an ortholog of S. cerevisiae PAH1, encoding PA phosphatase and an ortholog of SAC1, encoding PIP phosphatase in the ER. These results suggest that Yas3p is localized to the ER by binding to PA and/or PIP in the ER membrane. In contrast to S. cerevisiae, an ortholog of SCS2 or its paralog, SCS22, was not required for the transcriptional activation of ALK1 by n-alkanes in line with the absence of FFAT motif-like sequence in Yas3p, although the deletion mutant of SCS2 exhibited a growth defect when cultured on n-decane (Kobayashi et al. 2008). Based on these results, a model of the transcriptional regulation of n-alkane metabolic genes was proposed (Fig. 3). A heterocomplex of the bHLH transcription activators, Yas1p and Yas2p, constitutively localizes in the nucleus and binds to ARE1 in the promoter regions of the genes involved in the n-alkane metabolism. In the absence of n-alkanes, Yas3p is transported to the nucleus and binds to Yas2p of the Yas1p-Yas2p complex, resulting in repression of the ARE1-dependent transcription. When the medium is supplemented with nalkanes, Yas3p is retained to the ER by binding to PA and/or PIP, and the transcription is activated by Yas1p-Yas2p complex. It remains to be determined whether the amounts of PA and PIPs in the ER membrane increase in response to n-alkanes. Scs2p is not involved in this process, but it is possible that other ER-resident protein is involved in the localization of Yas3p to the ER. This is suggested by the fact that the C-terminal region of Yas3p was found to be localized to the ER in a PA- and PI (4)P-independent manner (Kobayashi et al. 2015).

8

R. Fukuda and A. Ohta

Fig. 3 Model of n-alkaneresponsive transcriptional regulation by the transcription factors Yas1p, Yas2p, and Yas3p in Y. lipolytica. In the absence of n-alkanes (upper), Opi1-family transcription repressor Yas3p binds to the complex composed of bHLH transcription activators Yas1p and Yas2p in the nucleus, and ARE1-dependent transcription is repressed. In the presence of n-alkanes (lower), Yas3p is sequestered to the ER membrane through the interaction with PA and/or PIP, and ARE1-dependent transcription is activated by Yas1p-Yas2p complex

− n-Alkane

Yas3p Yas2p Yas1p

Genes for n-alkane assimilation

ARE1

ALK1 etc.

nucleus ER

+ n-Alkane

n-Alkane

Yas3p

PA PIP

Yas2p Yas1p ARE1

Genes for n-alkane assimilation ALK1 etc.

nucleus ER

3.4

Role of the Opi1-Family Proteins in Other Yeasts

Orthologs of OPI1 exist in the genomes of a variety of yeasts (Fig. 4), although their functions are largely unknown. In Candida glabrata, a yeast closely related to S. cerevisiae phylogenetically, a homolog of Opi1p is involved in the transcriptional regulation of an ortholog of INO1 by myo-inositol (Bethea et al. 2010). In contrast, Opi1p homolog does not regulate INO1 expression in C. albicans, but it controls the expression of SAP2 encoding the secreted aspartyl protease and is involved in the filamentous growth and virulence (Chen et al. 2015). Therefore, Opi1-family proteins possibly regulate processes other than phospholipid synthesis. n-Alkane assimilating yeasts C. tropicalis, C. maltosa, C. dubliniensis, C. parapsilosis, D. hansenii, L. elongisporus, and M. guilliermondii all have Opi1-family proteins, and it would be of great interest to examine whether these orthologs are involved in the transcriptional regulation of n-alkane metabolism in these yeasts.

Genetic Features and Regulation of n-Alkane Metabolism in Yeasts

Saccharomyces cerevisiae Candida glabrata

1000 1000

Ashbya gossypii

999

644

9

Kluyveromyces lactis Komagataella pastoris

1000

Ogataea polymorpha 685

742

Candida maltosa

1000 1000

Candida tropicalis

1000

Candida albicans Candida dubliniensis

Candida parapsilosis

1000

863

Lodderomyces elongisporus Debaryomyces hansenii

978 1000

Meyerozyma guilliermondii Yarrowia lipolytica Rhodosporidium toruloides Ustilago maydis 0.1

Fig. 4 Phylogenetic tree of the Opi1-family proteins in yeasts. Phylogenetic tree of the Opi1family proteins of yeasts was constructed using ClustalW (DDBJ, v2.1) and drawn using Njplot. The scale bar indicates 0.1 substitutions per site. The bootstrap values by 1000 repetitions are indicated. The accession numbers of sequences from UniProtKB are as follows: Ashbya gossypii (Q75DH7), Candida albicans (Q5ALN4), Candida dubliniensis (B9WAR2), Candida glabrata (Q6FN27), C. maltosa (M3JFB2), Candida parapsilosis (G8BGL1), C. tropicalis (C5M6C2), Debaryomyces hansenii (Q6BJD0), Kluyveromyces lactis (Q6CIM8), Komagataella pastoris (A0A1B2J6H0), Lodderomyces elongisporus (A5DT94), Meyerozyma guilliermondii (A5DPS9), Ogataea polymorpha (A0A1B7SCJ0), Rhodosporidium toruloides (M7XL84), S. cerevisiae (P21957), Ustilago maydis (A0A0D1CM93), and Y. lipolytica (B9X0I4)

4

Repression of Transcription of n-Alkane Metabolic Genes

Glucose is the primary source of carbon and energy for most organisms, and the transcription of genes involved in other carbon and energy source metabolism remains repressed in the presence of glucose. Transcriptional repression of genes involved in n-alkane metabolism by glucose is observed in n-alkane-assimilating yeasts. In C. tropicalis and C. bombicola, expression of a subset of the CYP52family P450 genes induced by n-alkanes is repressed by glucose (Seghezzi et al. 1992; Van Bogaert et al. 2009). In C. maltosa, transcription of most of the ALK genes is severely repressed by glucose, but not by glycerol (Ohkuma et al. 1995a). The carbon catabolite repression by glucose has been well documented in S. cerevisiae (Conrad et al. 2014; Kayikci and Nielsen 2015), but it remains to be elucidated

10

R. Fukuda and A. Ohta

whether the transcription of the P450 genes is repressed by a similar mechanism in these n-alkane-assimilating yeasts. In marked contrast to these yeasts, in Y. lipolytica, the transcription of genes involved in n-alkane metabolism is strictly repressed by glycerol, but not so much by glucose (Iida et al. 1998, 2000; Mori et al. 2013). In line with this observation, glycerol is a preferred carbon and energy source for this organism, and it shows better growth on glycerol than on glucose (Mori et al. 2013). The transcriptional repression by glycerol was also observed in Kluyveromyces lactis, in which the transcription of KlICL1 encoding isocitrate lyase was repressed by glycerol (Rodicio et al. 2008). The molecular mechanisms underlying the transcriptional repression by glycerol is unclear, but it was shown that phosphorylation of glycerol is required for the glycerol repression in both these yeasts (Mori et al. 2013; Rodicio et al. 2008).

5

Research Needs

It has been revealed that, in Y. lipolytica, the Opi1-family protein Yas3p plays a pivotal role in the transcriptional regulation of genes involved in n-alkane metabolism by n-alkanes. However, it remains to be clarified how n-alkanes are recognized and how these signals are transduced retaining Yas3p to the ER. Transcriptional activation of the ARE1-containing promoter by n-alkane was also observed in the deletion mutant of the 12 ALK genes, which could not utilize n-alkanes owing to a defect in the hydroxylation of n-alkanes (Takai et al. 2012). This suggests that n-alkanes and not their metabolites activate the ARE1-mediated transcription. It is possible that there are proteins that sense n-alkanes and trigger the transcriptional response to n-alkanes. Alternatively, since n-alkanes are supposed to accumulate in the membranes, alterations in the membrane conditions may be perceived by a sensor protein. n-Alkane-assimilating yeasts have been shown to have great potential for production of single-cell protein (SCP) as well as various useful compounds, including long-chain dicarboxylic acids, by metabolizing n-alkanes (Barth and Gaillardin 1996, 1997; Fickers et al. 2005; Mauersberger et al. 1996; Tanaka and Fukui 1989). Elucidation of the mechanisms underlying the regulation of n-alkane metabolism will contribute to the construction of efficient bioconversion systems using these n-alkane-assimilating yeasts.

References Barth G, Gaillardin C (1996) Yarrowia lipolytica. In: Wolf K (ed) Non-conventional yeast in biotechnology. A handbook. Springer, Berlin, pp 313–388 Barth G, Gaillardin C (1997) Physiology and genetics of the dimorphic fungus Yarrowia lipolytica. FEMS Microbiol Rev 19:219–237 Bethea EK, Carver BJ, Montedonico AE, Reynolds TB (2010) The inositol regulon controls viability in Candida glabrata. Microbiology 156:452–462

Genetic Features and Regulation of n-Alkane Metabolism in Yeasts

11

Butler G, Rasmussen MD, Lin MF, Santos MA, Sakthikumar S et al (2009) Evolution of pathogenicity and sexual reproduction in eight Candida genomes. Nature 459:657–662 Chen YL, de Bernardis F, Yu SJ, Sandini S, Kauffman S et al (2015) Candida albicans OPI1 regulates filamentous growth and virulence in vaginal infections, but not inositol biosynthesis. PLoS One 10:e0116974 Conrad M, Schothorst J, Kankipati HN, Van Zeebroeck G, Rubio-Texeira M, Thevelein JM (2014) Nutrient sensing and signaling in the yeast Saccharomyces cerevisiae. FEMS Microbiol Rev 38:254–299 Craft DL, Madduri KM, Eshoo M, Wilson CR (2003) Identification and characterization of the CYP52 family of Candida tropicalis ATCC 20336, important for the conversion of fatty acids and alkanes to α,ω-dicarboxylic acids. Appl Environ Microbiol 69:5983–5991 Dujon B, Sherman D, Fischer G, Durrens P, Casaregola S et al (2004) Genome evolution in yeasts. Nature 430:35–44 Endoh-Yamagami S, Hirakawa K, Morioka D, Fukuda R, Ohta A (2007) Basic helix-loop-helix transcription factor heterocomplex of Yas1p and Yas2p regulates cytochrome P450 expression in response to alkanes in the yeast Yarrowia lipolytica. Eukaryot Cell 6:734–743 Federovitch CM, Ron D, Hampton RY (2005) The dynamic ER: experimental approaches and current questions. Curr Opin Cell Biol 17:409–414 Fickers P, Benetti PH, Waché Y, Marty A, Mauersberger S et al (2005) Hydrophobic substrate utilisation by the yeast Yarrowia lipolytica, and its potential applications. FEMS Yeast Res 5:527–543 Fukuda R (2013) Metabolism of hydrophobic carbon sources and regulation of it in n-alkaneassimilating yeast Yarrowia lipolytica. Biosci Biotechnol Biochem 77:1149–1154 Fukuda R, Ohta A (2013) Utilization of hydrophobic substrate by Yarrowia lipolytica. In: Barth G (ed) Yarrowia lipolytica genetics, genomics, and physiology. Springer, Heidelberg, pp 111–119 Gurvitz A, Rottensteiner H (2006) The biochemistry of oleate induction: transcriptional upregulation and peroxisome proliferation. Biochim Biophys Acta 1763:1392–1402 Henry SA, Kohlwein SD, Carman GM (2012) Metabolism and regulation of glycerolipids in the yeast Saccharomyces cerevisiae. Genetics 190:317–349 Heyken WT, Repenning A, Kumme J, Schuller HJ (2005) Constitutive expression of yeast phospholipid biosynthetic genes by variants of Ino2 activator defective for interaction with Opi1 repressor. Mol Microbiol 56:696–707 Hirakawa K, Kobayashi S, Inoue T, Endoh-Yamagami S, Fukuda R, Ohta A (2009) Yas3p, an Opi1 family transcription factor, regulates cytochrome P450 expression in response to n-alkanes in Yarrowia lipolytica. J Biol Chem 284:7126–7137 Iida T, Ohta A, Takagi M (1998) Cloning and characterization of an n-alkane-inducible cytochrome P450 gene essential for n-decane assimilation by Yarrowia lipolytica. Yeast 14:1387–1397 Iida T, Sumita T, Ohta A, Takagi M (2000) The cytochrome P450ALK multigene family of an n-alkane-assimilating yeast, Yarrowia lipolytica: cloning and characterization of genes coding for new CYP52 family members. Yeast 16:1077–1087 Iwama R, Kobayashi S, Ohta A, Horiuchi H, Fukuda R (2014) Fatty aldehyde dehydrogenase multigene family involved in the assimilation of n-alkanes in Yarrowia lipolytica. J Biol Chem 289:33275–33286 Iwama R, Kobayashi S, Ohta A, Horiuchi H, Fukuda R (2015) Alcohol dehydrogenases and an alcohol oxidase involved in the assimilation of exogenous fatty alcohols in Yarrowia lipolytica. FEMS Yeast Res 15:fov014 Iwama R, Kobayashi S, Ishimaru C, Ohta A, Horiuchi H, Fukuda R (2016) Functional roles and substrate specificities of twelve cytochromes P450 belonging to CYP52 family in n-alkane assimilating yeast Yarrowia lipolytica. Fungal Genet Biol 91:43–54 Jones T, Federspiel NA, Chibana H, Dungan J, Kalman S et al (2004) The diploid genome sequence of Candida albicans. Proc Natl Acad Sci U S A 101:7329–7334 Kayikci Ö, Nielsen J (2015) Glucose repression in Saccharomyces cerevisiae. FEMS Yeast Res 15: fov068

12

R. Fukuda and A. Ohta

Kobayashi S, Hirakawa K, Fukuda R, Ohta A (2008) Disruption of the SCS2 ortholog in the alkaneassimilating yeast Yarrowia lipolytica impairs its growth on n-decane, but does not impair inositol prototrophy. Biosci Biotechnol Biochem 72:2219–2223 Kobayashi S, Hirakawa K, Horiuchi H, Fukuda R, Ohta A (2013) Phosphatidic acid and phosphoinositides facilitate liposome association of Yas3p and potentiate derepression of ARE1 (alkaneresponsive element one)-mediated transcription control. Fungal Genet Biol 61:100–110 Kobayashi S, Tezaki S, Horiuchi H, Fukuda R, Ohta A (2015) Acidic phospholipid-independent interaction of Yas3p, an Opi1-family transcriptional repressor of Yarrowia lipolytica, with the endoplasmic reticulum. Yeast 32:691–701 Lebeault JM, Lode ET, Coon MJ (1971) Fatty acid and hydrocarbon hydroxylation in yeast: role of cytochrome P-450 in Candida tropicalis. Biochem Biophys Res Commun 42:413–419 Loewen CJ, Gaspar ML, Jesch SA, Delon C, Ktistakis NT et al (2004) Phospholipid metabolism regulated by a transcription factor sensing phosphatidic acid. Science 304:1644–1647 Massey SE, Moura G, Beltrão P, Almeida R, Garey JR et al (2003) Comparative evolutionary genomics unveils the molecular mechanism of reassignment of the CTG codon in Candida spp. Genome Res 13:544–557 Mauersberger S, Kärgel E, Matyashova RNM, Mueller HG (1987) Subcellular organization of alkane oxidation in the yeast Candida maltosa. J Basic Microbiol 27:565–582 Mauersberger S, Ohkuma M, Schunck WH, Takagi M (1996) Candida maltosa. In: Wolf K (ed) Nonconventional yeasts in biotechnology. Springer, Berlin, pp 411–580 Mori K, Iwama R, Kobayashi S, Horiuchi H, Fukuda R, Ohta A (2013) Transcriptional repression by glycerol of genes involved in the assimilation of n-alkanes and fatty acids in yeast Yarrowia lipolytica. FEMS Yeast Res 13:233–240 Murre C, Bain G, van Dijk MA, Engel I, Furnari BA et al (1994) Structure and function of helixloop-helix proteins. Biochim Biophys Acta 1218:129–135 Nelson DR (2009) The cytochrome p450 homepage. Hum Genomics 4:59–65 Nicaud JM (2012) Yarrowia lipolytica. Yeast 29:409–418 Ohkuma M, Tanimoto T, Yano K, Takagi M (1991) CYP52 (cytochrome P450alk) multigene family in Candida maltosa: molecular cloning and nucleotide sequence of the two tandemly arranged genes. DNA Cell Biol 10:271–282 Ohkuma M, Muraoka S, Tanimoto T, Fujii M, Ohta A, Takagi M (1995a) CYP52 (cytochrome P450alk) multigene family in Candida maltosa: identification and characterization of eight members. DNA Cell Biol 14:163–173 Ohkuma M, Park SM, Zimmer T, Menzel R, Vogel F et al (1995b) Proliferation of intracellular membrane structures upon homologous overproduction of cytochrome P-450 in Candida maltosa. Biochim Biophys Acta 1236:163–169 Osumi M, Miwa N, Teranishi Y, Tanaka A, Fukui S (1974) Ultrastructure of Candida yeasts grown on n-alkanes. Appearance of microbodies and its relationship to high catalase activity. Arch Microbiol 99:181–201 Rodicio R, López ML, Cuadrado S, Cid AF, Redruello B et al (2008) Differential control of isocitrate lyase gene transcription by non-fermentable carbon sources in the milk yeast Kluyveromyces lactis. FEBS Lett 582:549–557 Sanglard D, Chen C, Loper JC (1987) Isolation of the alkane inducible cytochrome P450 (P450alk) gene from the yeast Candida tropicalis. Biochem Biophys Res Commun 144:251–257 Schunck WH, Vogel F, Gross B, Kärgel E, Mauersberger S et al (1991) Comparison of two cytochromes P-450 from Candida maltosa: primary structures, substrate specificities and effects of their expression in Saccharomyces cerevisiae on the proliferation of the endoplasmic reticulum. Eur J Cell Biol 55:336–345 Seghezzi W, Sanglard D, Fiechter A (1991) Characterization of a second alkane-inducible cytochrome P450-encoding gene, CYP52A2, from Candida tropicalis. Gene 106:51–60 Seghezzi W, Meili C, Ruffiner R, Kuenzi R, Sanglard D, Fiechter A (1992) Identification and characterization of additional members of the cytochrome P450 multigene family CYP52 of Candida tropicalis. DNA Cell Biol 11:767–780

Genetic Features and Regulation of n-Alkane Metabolism in Yeasts

13

Sugiyama H, Ohkuma M, Masuda Y, Park SM, Ohta A, Takagi M (1995) In vivo evidence for non-universal usage of the codon CUG in Candida maltosa. Yeast 11:43–52 Sumita T, Iida T, Yamagami S, Horiuchi H, Takagi M, Ohta A (2002) YlALK1 encoding the cytochrome P450ALK1 in Yarrowia lipolytica is transcriptionally induced by n-alkane through two distinct cis-elements on its promoter. Biochem Biophys Res Commun 294:1071–1078 Takai H, Iwama R, Kobayashi S, Horiuchi H, Fukuda R, Ohta A (2012) Construction and characterization of a Yarrowia lipolytica mutant lacking genes encoding cytochromes P450 subfamily 52. Fungal Genet Biol 49:58–64 Tanaka A, Fukui S (1989) Metabolism of n-alkanes. In: The yeast. Academic, London, pp 261–287 Tenagy PJS, Iwama R, Kobayashi S, Ohta A et al (2015) Involvement of acyl-CoA synthetase genes in n-alkane assimilation and fatty acid utilization in yeast Yarrowia lipolytica. FEMS Yeast Res 15:fov031 Ueda T, Suzuki T, Yokogawa T, Nishikawa K, Watanabe K (1994) Unique structure of new serine tRNAs responsible for decoding leucine codon CUG in various Candida species and their putative ancestral tRNA genes. Biochimie 76:1217–1222 Van Bogaert IN, De Mey M, Develter D, Soetaert W, Vandamme EJ (2009) Importance of the cytochrome P450 monooxygenase CYP52 family for the sophorolipid-producing yeast Candida bombicola. FEMS Yeast Res 9:87–94 Vogel F, Gengnagel C, Kärgel E, Müller HG, Schunck WH (1992) Immunocytochemical localization of alkane-inducible cytochrome P-450 and its NADPH-dependent reductase in the yeast Candida maltosa. Eur J Cell Biol 57:285–291 Yamagami S, Iida T, Nagata Y, Ohta A, Takagi M (2001) Isolation and characterization of acetoacetyl-CoA thiolase gene essential for n-decane assimilation in yeast Yarrowia lipolytica. Biochem Biophys Res Commun 282:832–838 Yamagami S, Morioka D, Fukuda R, Ohta A (2004) A basic helix-loop-helix transcription factor essential for cytochrome P450 induction in response to alkanes in yeast Yarrowia lipolytica. J Biol Chem 279:22183–22189

Current View of The Mechanisms Controlling The Transcription of The TOL Plasmid Aromatic Degradation Pathways Patricia Domínguez-Cuevas and Silvia Marqués

Abstract

The TOL plasmid-encoded pathway for the degradation of toluene and derivatives is an archetype in bacterial transcription regulation. Six promoters belonging to different classes and several chromosome- and plasmid-encoded proteins are involved in maintaining optimal expression levels and synchronization with the global cell metabolism. The TOL-encoded regulators are the enhancer-binding protein XylR, which controls the σ54-dependent promoters of the upper pathway Pu and of xylS gene PS1, and the AraC family regulator XylS, which controls the σ32-σ38-dependent meta-cleavage pathway promoter Pm. Both regulators respond to the presence of a specific effector and activate transcription through different mechanisms. Much effort has been devoted to the elucidation of these processes. In this review, recent results are described and discussed in the light of the latest findings and models for homologous family proteins and their interrelationships with the cell metabolism.

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 2 The Upper Pathway: Onset of the Regulatory Cascade . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 3 The Meta-Cleavage Pathway: Current Understanding of the Activation Mechanism . . . . . . 7 3.1 New Insights into the Cross Regulation of the Meta- and Ortho-Cleavage Pathways for Benzoate Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10

P. Domínguez-Cuevas Department of Biology, University of Copenhagen, Copenhagen N., Denmark S. Marqués (*) Department of Environmental Protection, Estación Experimental del Zaidín, CSIC, Granada, Spain e-mail: [email protected] # Springer International Publishing AG 2017 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-319-39782-5_29-2

1

2

P. Domínguez-Cuevas and S. Marqués

3.2 A Family of Anti-activators Accompanying XylS/AraC Regulators . . . . . . . . . . . . . . . . . . 4 Integration in the Cell Regulatory Networks: Toward Optimization of Expression . . . . . . . . 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

11 11 14 15

Introduction

Pseudomonas putida mt-2 TOL plasmid pWW0 was one of the first plasmids described to contain an entire aromatic degradation pathway (Williams and Murray 1974). It codes for two catabolic routes responsible for toluene and benzoate degradation, the upper and meta-cleavage pathways, respectively (Assinder and Williams 1990), flanked by two insertion sequences (IS1246). The genes are organized in two operons located 10 kb apart (upper and meta-cleavage), clustered next to the divergent regulatory genes xylR and xylS, which coordinate the expression of the two pathways (Fig. 1a) (Greated et al. 2002). The regulatory network can be described as follows: when P. putida mt-2 cultures are exposed to toluene, the XylR regulator becomes active to promote transcription from two σ54-dependent promoters, the upper pathway promoter Pu and the xylS gene promoter PS1 (reviewed in Ramos et al. 1997). Activation of PS1 leads to increased synthesis of XylS, which in these conditions is able to promote expression from the meta-cleavage pathway promoter Pm even in the absence of its effector. This mechanism is known as the cascade regulatory loop (Marqués and Ramos 1993; Ramos et al. 1997). In the absence of toluene, transcription from Pm can be induced by the addition of benzoate and some substituted derivatives that activate XylS, a regulatory pathway known as the metaloop (Ramos et al. 1986; Inouye et al. 1987). Furthermore, toluene-dependent XylR induction of the upper pathway also leads to the metabolism of toluene to benzoate, the first substrate of the meta-cleavage pathway. In turn, and following a few minutes delay, this aromatic activates the XylS regulator to promote increased transcription from the meta-cleavage pathway promoter Pm (Marqués et al. 1994). A recent transcriptomic analysis of TOL pathway gene expression confirms the basic steps in this regulatory network and exposes previously overlooked subtleties of the pathway operon mRNA synthesis (Kim et al. 2016). The resulting synthesis of the pathway enzymes, however, takes several hours to reach maximum steady levels (Hugouvieux-Cotte-Pattat et al. 1990). The extensive analysis of these processes shows that a considerable number of chromosome- and plasmid-encoded proteins cooperate to maintain an optimal pathway expression level in every circumstance (Table 1). The general patterns of this network have already been revised (Marqués and Ramos 1993; Ramos et al. 1997; Ruíz et al. 2004; Daniels et al. 2008; Domínguez-Cuevas and Marqués 2010). The intricacy and refinement of the regulatory network allow for an appropriate synchronization with the global cell metabolism. Upon the well-established effector-dependent specific control mechanisms, increasing regulatory steps connecting the TOL pathway with the host metabolic network are continuously being uncovered, reflecting the complexity and fine control of the pathway. This makes of the TOL system an archetype in bacterial transcription

Current View of The Mechanisms Controlling The Transcription of The TOL. . .

a

3

b sensor

tra/rep

DNA-binding

AAA+

HTH

514 556

472

211

233

C

D

R S

c Tn4651

DNA-binding

effector binding/ dimerization

HTH-1

HTH-2

232-253

283-306

XylS 321

meta

B

204 214

IS

A IS

1

pWW0  kbp

XylR 1

Tn4653

upper

central

VR

Fig. 1 (a) The pWW0 TOL plasmid aromatic degradation pathway is organized in two operons and two regulatory genes included in two transposable elements inserting one within another. (b) Schematic presentation of XylR modular organization into three functional domains: a sensor domain (A), connected by a linker (B) to the central or AAA+ domain (C), and a DNA-binding domain (D). (c) Schematic presentation of the XylS protein functional domains: effector binding and dimerization domain and DNA-binding domain containing two HTH motives. The residue numbers delimiting each domain are indicated in each case

regulation, which has been exploited in fundamental and applied research fields. This chapter focuses on the most recent findings, especially those explaining the mechanistic and fine-tuned functioning of the pathway and its integration into the cell regulatory network.

2

The Upper Pathway: Onset of the Regulatory Cascade

XylR is the primary regulator of the TOL pathway. It belongs to the large AAA+ family of ATPases (Neuwald et al. 1999; Studholme and Dixon 2003) which includes, among others, the σ54-dependent promoter activator family known as bacterial enhancer-binding proteins (bEBPs). These proteins consist of three main domains: an N-terminal sensor domain (A) sensing the regulatory signal (e.g., the presence of an effector in the case of XylR (Delgado and Ramos 1994), connected through a B-linker to a central AAA+ domain (C) with ATPase activity, and a C-terminal DNA-binding domain (D) (Fig. 1b) (Bush and Dixon 2012). The role of the three main domains has been well established in XylR. Genetic analysis located the mutations altering the effector profile in the N-terminal domain (NTD) of the protein (Delgado and Ramos 1994; Salto et al. 1998; Garmendia et al. 2001, 2008). As for other bEBPs where the N-terminal domain exerts a negative control, a XylR protein devoid of the NTD shows constitutive activity (Pérez-Martín and De Lorenzo 1995a; Garmendia and de Lorenzo 2000). This domain given in trans exerts specific intramolecular repression, inhibiting ATP-binding capacity of the C domain; repression is released in the presence of an effector (Fernández et al. 1995; PérezMartín and De Lorenzo 1995a). Although in XylR physical interaction with the

4

P. Domínguez-Cuevas and S. Marqués

Table 1 Protein factors involved in TOL pathway regulation Factor XylR

Gene xylR

XylS

xylS

Sigma-70

rpoD

Sigma-38

rpoS

Sigma-32

rpoH

Sigma-54

rpoN

IHF

him, hip ptsN crc/ hfq

IIANtr Crc/Hfq

References

meta-cleavage pathway (+) xylR (+), xylS (+) meta-cleavage pathway (+)

Marqués et al. (1995)

meta-cleavage pathway (+)

Marqués et al. (1999)

Upper pathway (+), xylS (+)

Gallegos et al. (1996a), Marqués et al. (1998)

Upper pathway (+), xylS (
)

Holtel et al. (1995) Cases et al. (1999) Moreno et al. (2010, 2015)

FtsH

ftsH

General

HU

hupA, hupB pprA turA

General

xylS (+)

General General, low temperature General

Upper pathway (
) Upper pathway (
)

Carmona and de Lorenzo (1999), Sze et al. (2002) Pérez-Martín and de Lorenzo (1995b) Vitale et al. (2008) Rescalli et al. (2004)

Upper pathway (
) in E. coli

Zhang et al. (2014)

CRP

crp/ cya (vfr)

General General

Target (role)a Upper pathway (+), xylS (+)

Upper pathway, xylS Upper pathway (
), metacleavage pathway (
), xylR (
), xylS (
) Upper pathway (+)

PprA TurA

a

Category Pathway specific Pathway specific General, house keeping General, stress/ stationary General, stress General, nitrogen, other functions General

(+) positive effect; (
) negative effect

effector has not been analyzed, effector binding analysis in the phenol-responsive XylR homologue DmpR revealed that actually the interaction of labeled phenol with the N-terminal domain released C-domain ATPase activity repression (Shingler and Pavel 1995). Direct interaction with different effectors was proven by isothermal titration calorimetry (ITC) in MopR, a phenol-responsive XylR-DmpR homologue (Ray et al. 2016). Early structure prediction based on the alignment of 11 XylR family proteins and homology modeling suggested a fold similar to V4R domain (Pfam 02830) (Devos et al. 2002). This predicted structure was recently endorsed by the crystal structure of MopR NTD, the first structure available for a sensor domain of the XylR-NtrC subfamily of bEBPs (Ray et al. 2016). The NTD of both XylR and DmpR can be modeled using MopR structure, showing that the actual XylR residues

Current View of The Mechanisms Controlling The Transcription of The TOL. . .

5

involved in direct interaction with the effector are different from the previously suggested from genetic analysis. The specific role of several previously mutated residues could be inferred from the protein structure. Unexpectedly, the MopR crystal structure revealed the presence of a Zn-binding domain that was conserved in XylR and DmpR. According to the proposed model, this Zn-binding domain, together with the B-linker, would be directly involved in opening and closing an effector gate to the binding site and in triggering communication with the central domain, as suggested for several bEBPs (Bose et al. 2008; Ray et al. 2016). This is consistent with the previous suggestion, based on mutant analysis, of XylR B-linker influencing effector binding (Garmendia and de Lorenzo 2000). Structural analyses of several bEBPs show that this domain also regulates multimerization and couples the A-domain input signal with the C-domain ATPase activity (Bose et al. 2008). Although this has not been directly demonstrated in XylR, the strong homology among these proteins points toward a similar mechanism operating in this regulator. The AAA+ central domain (C) is the most conserved among bEBPs. It carries the ATP-binding motif, ATPase activity, and σ54 interaction determinants, features that are essential to σ54 promoter activation (Schumacher et al. 2006; Chen et al. 2008). According to the first XylR activation model based on the analysis of a truncated protein devoid of its N-terminal domain, the activation mechanism followed a cyclic sequence of events where ATP binding to the central domain triggered XylR multimerization at its binding site, followed by ATP hydrolysis, promoter activation, and return to the non-multimerized structure (Pérez-Martín and de Lorenzo 1996a). The recent availability of the crystal structure of several regulators of the family either devoid of their NTD or naturally lacking this domain has helped increase our knowledge of the multistep process leading to promoter activation by these proteins (Lee et al. 2003; Rappas et al. 2005; Sallai and Tucker 2005; reviewed in Bush and Dixon 2012). Although the mechanistic model has only been proposed for those members belonging to a two-component system or for those naturally lacking an NTD, XylR is likely to share many of their features. Proteins of this family generally bind DNA as dimers. After the A-domain repression has been released (in XylR, effector binding would trigger A-domain movement and initiate derepression), a conformational rearrangement induces protein oligomerization to a DNA-bound hexameric structure, which is followed by ATP binding to the regulator in the interface of two subunits. Interestingly, both adjacent subunits contribute to ATP binding and hydrolysis, thus explaining the previously observed ATPase activity dependence on protein oligomerization (Bordes et al. 2003; reviewed in Schumacher et al. 2006). Two C-domain specific loops pointing toward the inner pore of the hexameric ring undergo a conformational reorganization during ATP hydrolysis, reorienting the AAA+-family conserved GAFTGA motif present in one of the loops to allow contacts with σ54 (Bose et al. 2008; Wigneshweraraj et al. 2008). Mutations in these loops in XylR abolish ATP binding and hydrolysis (Pérez-Martin and de Lorenzo 1996a). Contacts between the regulator and the transcriptional machinery require that the two protein complexes are brought into proximity by DNA bending, generally facilitated by the integration host factor (IHF). Finally, σ54

6

P. Domínguez-Cuevas and S. Marqués

undergoes a structural remodeling, allowing the σ54-RNA polymerase (Eσ54) closed complex to isomerize to an open complex. As in all bEBPs, in XylR sequence the central domain is followed by the DNA-binding domain (D) encompassing a typical helix-turn-helix (HTH) structure which confers binding specificity on the target promoter (Inouye et al. 1988). Comparison of the structure of the HTH domains of a series of bEBPs shows that they all share a similar DNA recognition element formed by a three-helical bundle, where the C-terminal helix is the DNA-contacting element (Vidangos et al. 2013). This structure is conserved in XylR structure model. XylR dimers are always bound to the two upstream activation sequences (UASs) in Pu (Abril et al. 1991). As discussed above for other bEBPs, binding of ATP induces multimerization to a hexameric conformation, which is then able to hydrolyze ATP (Pérez-Martín and de Lorenzo 1996b). Binding of bEBPs to their UASs is thought to generate high local concentrations of activated regulator in the proximity of Eσ54-promoter complex. The relevant regulatory sequences in Pu span 108 bp, and the XylR binding site is composed of two UASs located between positions
120 and
175 (Holtel et al. 1990). Interestingly, these UASs, the furthermost sequence of the TOL xyl region located adjacent to IS1246, prevent read-through from upstream promoters, thus isolating pathway expression from external influence (Velázquez et al. 2006). UV laser footprint and atomic force microscopy confirmed binding of IHF between positions
52 and
79 (Valls et al. 2002), inducing the strong DNA bending needed for interaction between the UAS-bound regulator and the RNA polymerase (RNAP) machinery bound at
12/
24 (de Lorenzo et al. 1991). Early models explaining bEBP activation mechanism suggested direct interaction between the regulator and Eσ54, in most cases brought to proximity by IHF-assisted DNA looping. Interestingly, the regulator approaches the Eσ54 closed complex through the unbound face of Eσ54 binding site to contact σ54 and catalyze open complex formation, so that DNA appears sandwiched between Eσ54 and the regulator (Huo et al. 2006; Wigneshweraraj et al. 2008). Despite the profusion of bEBP structures and biochemical analysis available in the past years, the precise mechanism underlying energy coupling from regulator ATP hydrolysis to promoter melting and escape is not yet fully understood (see Bush and Dixon 2012 for a comprehensive review on bEBP structure and activation mechanism). The structure of the PS1 promoter slightly differs from the canonical σ54-promoter architecture, probably because it accommodates the two overlapping divergent promoters PR1 and PR2 responsible for xylR expression and PS2 xylS constitutive promoter which maintains XylS basal concentrations in the cell (Gallegos et al. 1996a). In fact, XylR UASs at position
133 to
207 of PS1 overlap the divergent
10/
35 Eσ70 binding sites of the two xylR promoters PR1 and PR2. Two consensus IHF binding sequences are found, overlapping the
12/
24 Eσ54 binding site and the UASs. As a consequence of this complex organization, expression of XylR regulator never achieves maximum levels (Marqués et al. 1998): when XylR binds its UASs to activate PS1, it represses its own synthesis (Holtel et al. 1992; Bertoni et al. 1998). Self-repression of XylR has been suggested to help buffering possible fluctuations in XylR synthesis levels in any condition (Koutinas et al. 2010); in fact,

Current View of The Mechanisms Controlling The Transcription of The TOL. . .

7

xylR gene expression levels were shown to be similar along the growth curve in the presence and absence of effector (Marqués et al. 1994). In contrast to Pu activation mechanism, the presence of IHF strongly represses PS1 activity (Marqués et al. 1998), which probably explains why PS1 expression levels have been estimated to be fourfold lower than Pu expression levels (Marqués et al. 1994). In PS1, the histonelike protein HU replaces the positive function of IHF by increasing a preexistent static bend in the DNA to bring into proximity the DNA-bound complexes of the regulator and Eσ54 (Pérez-Martín and de Lorenzo 1995b).

3

The Meta-Cleavage Pathway: Current Understanding of the Activation Mechanism

Expression of the meta-cleavage pathway is under the control of XylS-regulated Pm promoter. Expression along the growth curve is mediated by two stress sigma factors: σ32 (RpoH) in the early exponential phase and σ38 (RpoS) in the late exponential and stationary phases (Marqués et al. 1999). In fact, Pm
10/
35 RNAP binding sequence diverges considerably from the consensus defined for σ32, σ38, and σ70 factors (Domínguez-Cuevas et al. 2005). On one hand, XylS binding sites overlap the
35 region so this sequence is a compromise between the two binding consensus (González-Pérez et al. 2002); on the other hand, the
10 region of Pm must include the essential determinants for recognition by the two polymerases involved, Eσ32 and Eσ38. Unlike σ70, the amount of these two alternative sigma factors depends on the cell physiological state and requires stress conditions to reach effective amounts to compete with σ70 for core RNAP (Gross et al. 1998; Hengge-Aronis 2002). Global expression analyses of P. putida mt-2 revealed aromatic effectors such as toluene or 3-methylbenzoate (3MB) are good elicitors of the stress response (DomínguezCuevas et al. 2006). This general picture suggests the Pm promoter sequence has evolved to adapt the meta-cleavage pathway expression to the heat shock response triggered by the presence of toluene or 3MB (Domínguez-Cuevas et al. 2006). The increase in σ32 level during the heat shock response is strong but transient, so the Pm promoter sequence is optimized to accommodate the temporary Eσ32 RNAP. After this initial heat shock response, a second alternative RNAP associated with the stress/stationary sigma factor σ38 maintains Pm transcription at high levels. Thus, the roles of effector in Pm promoter activation mechanism are to activate XylS protein and to increase stress sigma factor levels to efficiently compete for core binding and promoter recognition (Fig. 2). XylS binding site at Pm is composed of two 15-bp direct repeats (positions
70 to
56 and
49 to
35) each divided in two sequence boxes A and B. Pm exhibits an intrinsic curvature centered in the A-track located between proximal boxes A and B, with an apparent bent angle of 35 , also observed in vivo (Gallegos et al. 1996b; González-Pérez et al. 2002). XylS belongs to the AraC family of transcription regulators (Gallegos et al. 1997; Tobes and Ramos 2002) and is composed of two separate functional domains; XylS mutants with altered effector specificity cluster in

8

P. Domínguez-Cuevas and S. Marqués COOH

CH3 R

CH3

R

Pu

activation XylR

σ54

XylR

upper

Eσ54

Effector

P S1

xylS RNAP core

overproduction

CH3

activation

XylS

XylS

COOH

XylS

XylS

CH3

XylS

meta-cleavage

Pm

Toluene

XylS

COOH

Effector

activation

XylS

Permanent Transient

XylS

CH3 3-methylbenzoate

Stress

Fast Slow

TCA cycle

Heat Shock Response 38

↑σ

↑σ

32

Eσ32 Eσ38

Fig. 2 Effector dual function in Pm activation. Red arrows indicate a direct positive effect on transcription. Light-green arrows indicate activation of the regulator. Black upward arrows indicate accumulation of a sigma factor

the N-terminal domain of the protein, indicating that this domain carries the effector recognition determinants (Ramos et al. 1990; Michán et al. 1992; Ruíz and Ramos 2002). In addition, mutations in residues Leu193, Leu194, and Ile205 on the C-terminal edge of this domain are impaired in XylS dimerization (Ruíz et al. 2003). On the other hand, genetic analysis located the DNA-binding domain at the C-terminal end of the protein, connected to the N-terminal domain by a short linker (Fig. 1c). It has been recently reported that in AraC the inter-domain linker plays an active role both in protein dimerization and affecting the interaction between the effector/dimerization and the DNA-binding domains (Seedorff and Schleif 2011). X-ray crystallographic structures of other AraC family proteins show that the DNA-binding domain is composed of seven α-helices folding in two HTH domains, which bind two adjacent segments of the major groove (Rhee et al. 1998; Kwon et al. 2000). XylS domains are functionally independent, so that XylS DNA-binding domain (XylSC) is able to activate transcription in spite of the absence of XylS N-terminal domain determinants, a process independent of the presence of effector (Domínguez-Cuevas et al. 2008b). Interestingly, newly reported data have shown that another member of the AraC/XylS family, VirF, is synthesized from two alternative translational start sites. The two produced protein forms play different functional roles; the full-length form activates Shigella spp. virulence system, while the short form, which includes the DNA-binding motif, negatively autoregulates virF expression itself (Di Martino et al. 2016). Intriguingly, reexamination of XylS open-

Current View of The Mechanisms Controlling The Transcription of The TOL. . .

9

reading frame disclosed the existence of two putative internal ATG start codons, which, hypothetically, could generate alterative shorter forms of the activator. Analysis of the XylS DNA-binding domain showed it binds DNA forming two complexes (CI and CII), corresponding to one or two XylS-C monomers bound to DNA, respectively (Domínguez-Cuevas et al. 2010). Affinity calculation, DNA bending angle estimation and footprinting assays of XylS C-terminal domain suggest the two monomers bind DNA sequentially and noncooperatively. The first XylS-C monomer binds Pm at the proximal site (closest to the RNAP binding site), raising Pm curvature from 35 to 50 . Simultaneously, the bent center shifts to the DNA region between XylS binding sites, and finally the binding of the second XylS-C monomer increases the DNA bending angle to 98 . This probably contributes to establish the XylS-RNAP contacts required for transcription activation (DomínguezCuevas and Marqués 2010). Our results indicate that sugar-phosphate backbone contacts greatly contribute to XylS/Pm binding strength (Domínguez-Cuevas et al. 2008b), so that Pm curvature around XylS monomers probably enhances nucleoprotein stability. In addition, XylS establishes base-specific contacts that are on the basis of unambiguous recognition of Pm direct repeats (Domínguez-Cuevas et al. 2008b). XylS dimer formation and DNA-binding capacity were enhanced in vivo by the presence of 3MB but became an effector-independent process at high protein concentration in vitro (Ruíz et al. 2003; Domínguez-Cuevas et al. 2008a). In fact, in the absence of inducer, expression from Pm increases as a function of XylS expression levels, reaching a maximum activity level that compares with the one under induced conditions (Zwick et al. 2013). These data support the proposal that XylS could exist in three states: monomers, dimers, and aggregates, being the dimeric form the only one active and able to induce transcription from the Pm promoter. This model suggests that XylS overproduction leads to the formation of inactive aggregates, which correlates with the absence of Pm induction at high intracellular XylS levels. As for the role of 3MB in XylS activation, data obtained with the two purified protein domains suggest intramolecular repression of XylS-NTD upon XylS-CTD DNA binding, which is released in the presence of 3MB (Domínguez-Cuevas et al. 2008a). Even though the mechanism for activation upon 3MB addition is not fully understood, these data are consistent with the role of arabinose during AraC activation. Upon arabinose binding, the AraC flexible arm restructures, which releases the DNA-binding domain from the dimerization domain, allowing transcription activation (Rodgers et al. 2009). The current model for XylS activation involves the following sequence of events: in the absence of 3MB, direct interaction between N- and C-terminal domains in XylS maintains the protein in an inactive state. The addition of 3MB releases N-terminal domain repression upon XylS-C, allowing XylS to bind DNA and to activate transcription. Thus, 3MB binding to XylS both triggers the conformational change favoring the dimerization and allows derepression of the DNA-binding domain. However, a XylS dimerization mutant able to bind DNA in the presence of 3MB remained inactive in transcription, indicating dimerization is an essential process in transcription activation. The mode of activation is slightly different in the absence of effector: at high XylS concentrations, XylS C-terminal domain

10

P. Domínguez-Cuevas and S. Marqués

derepression is favored, unmasking dimerization surface and DNA-binding determinants, which leads to Pm recognition and further activation.

3.1

New Insights into the Cross Regulation of the Metaand Ortho-Cleavage Pathways for Benzoate Degradation

Pseudomonas putida mt-2 encodes two alternative pathways for benzoate metabolism, the aforedescribed meta pathway and the chromosomally encoded ortho-cleavage pathway. The ortho pathway operon is transcribed from the Pben promoter in response to the BenR regulator, member of the AraC-XylS family of activators. The TOL pathway allows mineralization of toluene and m-xylene through the upper pathway. The methyl group of toluene and m-xylene is sequentially oxidized to render, respectively, benzoate or 3-methylbenzoate. Non-substituted benzoates can be degraded productively both through the ortho- and meta-cleavage pathways; however, methylated benzoates channeled via the ortho-cleavage pathway generate dead-end metabolites known as methyl-2-enelactones. This metabolic conflict posed when P. putida mt-2 cells, encoding both pathways, degrade 3-methylbenzoate has been analyzed in depth to conclude that it is overcome through a rather simple and effective regulatory solution. The pWW0-encoded XylS regulator is highly homologous to the chromosomally encoded BenR protein, to the point that both regulators show a degree of cross regulation (Kessler et al. 1994; Cowles et al. 2000; Domínguez-Cuevas et al. 2006). The operator sequences found in Pm and Pben are highly similar, however, while Pm has two complete operator sequences for XylS, Pben apparently presents three out of the four DNA boxes required for physiological XylS binding (Pérez-Pantoja et al. 2015). The lack of two complete operator sequences in Pben is sufficient to prevent activation of transcription by physiological levels of XylS (Pérez-Pantoja et al. 2015; Tsipa et al. 2016). In fact, generation of Pben promoter variants with two complete operator sequences resulted in increased activation levels in response to both inducers (Silva-Rocha and de Lorenzo 2012), due to improved recognition of the Pben promoter by 3MB-activated XylS. In connection with the simultaneous presence of the ortho and meta pathways, recent published data suggest that P. putida mt-2 contains an additional metabolic safety valve, the chromosomal catA2 gene, which helps counteracting the toxicity of high catechol concentrations (Jiménez et al. 2014). However, the most unexpected finding of a recent transcriptomic analysis of TOL operons in cells growing with different inducers was that the unsubstituted benzoate produced from toluene metabolism through the upper pathway was not able to induce the ortho pathway in the chromosome (Kim et al. 2016). Among the hypothesis suggested to explain this striking finding, physical channeling of TOL pathway metabolites in the cytoplasm toward the TOL enzymatic machinery opens up new interesting perspectives to analyze the spatial organization (compartmentalization) of cellular metabolism (de Lorenzo et al. 2015).

Current View of The Mechanisms Controlling The Transcription of The TOL. . .

3.2

11

A Family of Anti-activators Accompanying XylS/AraC Regulators

XylS belongs to the AraC/XylS family of transcriptional regulators. Several members of the AraC family involved in virulence gene regulation display an autoactivation mechanism (Martinez-Laguna et al. 1999; Porter et al. 2004; Morin et al. 2010) and, in contrast to XylS, AraC, or many other well-characterized members of the family, do not require effector binding for activation. Recently, a highly conserved family of small proteins named the AraC Negative Regulators (ANRs) has been shown to downregulate their AraC partners in pathogenic E. coli (Santiago et al. 2014, 2016). ANRs seem to play their regulatory roles by directly binding their cognate AraC partners, preventing DNA binding and activation of expression. Preliminary searches into the Pseudomonas genome and the TOL coding sequence did not render any putative candidates belonging to this new family of regulators. Nevertheless, we cannot discard the possibility that functional homologues of the ANR family members might exist and have not been identified to date.

4

Integration in the Cell Regulatory Networks: Toward Optimization of Expression

Table 1 lists the cell global regulators identified so far that are involved in TOL-mediated degradation of aromatic compounds. The abundance of host factors involved in TOL pathway regulation denotes a long coexistence of plasmid and host, which has led to the adaptation of plasmid gene expression to the cellular metabolism, taking advantage of the host machinery and using its network of general regulators. The main targets of this network are the σ54-dependent Pu and PS1 promoters and to a lesser extent Pm. Pu and PS1 respond to the availability of alternative carbon source, while Pm is optimized to maintain significant expression levels throughout the growth phase under induced conditions. The host factors affecting Pm expression have already been addressed above. Influence of global physiological conditions on Pu expression is observed in rich medium as a delay in the time course induction of the pathway after effector addition, which has also been called “exponential silencing” (reviewed in Cases and de Lorenzo 2005). This delay is also observed in PS1 and reflects the catabolite inhibition exerted by LB components, since it can be reproduced by the addition of casamino acids and of certain mixtures of amino acids (Marqués et al. 1994). This inhibition is at least partially dependent on the Crc and Hfq proteins (see below). Catabolite repression is also observed in defined minimal medium as reduced activation levels when a carbon source such as glucose, gluconate, or 2-ketoglutarate is present in addition to the effector (reviewed in Ruíz et al. 2004). These effects are mimicked in continuous culture, in which growth under carbon excess leads to total repression of the pathway in the presence of effector regardless of the carbon source used and regardless of the limiting substrate selected, while growth in carbon-limited conditions allows substantial expression when the effector is present (Duetz et al.

12

P. Domínguez-Cuevas and S. Marqués

1996). The mechanisms underlying this modulation have been analyzed for years, and a number of factors involved in the process have been described (Table 1). However, no clear picture of the regulatory network has emerged yet, and recent findings suggest that regulation is more intricate than initially considered. In Pseudomonas, the classical CRP-dependent regulation present in Enterobacteriaceae appears not to be involved in catabolite repression, rather the Pseudomonas CRP orthologue Vfr regulates processes such as quorum sensing response and virulence (Suh et al. 2002; Rojo 2010; Coggan and Wolfgang 2012). Although in some P. putida strains vfr was required for the assimilation of certain amino acids, organic acids, urea, and ammonia as carbon and/or nitrogen source (Daniels et al. 2010; Herrera et al. 2012), the function of the Vfr/cAMP tandem in this species is still unclear, although clearly unrelated to metabolic functions (Milanesio et al. 2011). In this sense, inactivation of vfr in P. putida KT2440 was clearly shown to be irrelevant for TOL pathway expression regulation through Pu and PS1 promoters (ArandaOlmedo et al. 2005). However, experiments in E. coli showed that Pu is sensitive to repression by CRP in a cAMP-dependent manner, probably by preventing direct contact between UAS-bound XylR with the promoter-bound RNA polymerase (Zhang et al. 2014). While it has been shown that P. putida Vfr maintains the capacity to bind DNA and cAMP and to contact RNAP, it seems to respond to different, so far unknown signals (Milanesio et al. 2011), which would explain the absence of effect of an vfr mutation on Pu and PS1 expression in its natural P. putida host. Three additional players in TOL expression have been considered as target of the global regulation. The first one was based on the regulatory network prevailing in Pseudomonas CF600 for the homologous system DmpR/Po, which involves the alarmone (p)ppGpp as main player (Sze and Shingler 1999). However, it seems this mechanism has only a marginal influence on XylR-mediated Pu expression (Carmona et al. 2000; Sze et al. 2002). The sigma factor σ54 was also explored as possible target for global regulation. In this sense, the detrimental effect of a mutation in the FtsH membrane protease gene on Pu activity suggested that the functionality of σ54, which is controlled by this protease, would influence Pu activity (Carmona and de Lorenzo 1999). However, comparative and combinatorial studies with the XylR/ Pu system and the related DmpR/Po tandem regulating phenol degradation in Pseudomonas sp. strain CF600 showed that the actual target of FtsH in Pu expression was the XylR regulator. FtsH, which belongs to the AAA+ family of chaperone-like ATPases, would be required for XylR correct folding or multimerization to form the regulator-Eσ54 complex (Sze et al. 2002). The third player is the PtsN protein. In most Gram-negative bacteria where the rpoN gene coding for σ54 is found, it appears clustered with three other genes: ptsN, which codes for the phosphoenolpyruvate: sugar phosphotransferase system (PTS) component EIIANtr (Deutscher et al. 2006), ptsP coding for EINtr, and ptsO coding for NPr. The phosphorelay cascade of this PTS system flows from phosphoenolpyruvate (PEP) through EINtr and NPr to EIIANtr. Inactivation of the ptsN gene leads to a partial release of glucose or succinatedependent Pu repression (Cases et al. 1999; Aranda-Olmedo et al. 2006). PtsN repressing activity on Pu depends on its phosphorylation level, where the non-phosphorylated form of the protein is the one exerting the repressive effect. In

Current View of The Mechanisms Controlling The Transcription of The TOL. . .

13

Pseudomonas, the PTS system is suggested to control carbon/nitrogen balance in the cell (Cases et al. 2007). Analysis of mutants in the carbon metabolism of P. putida showed that routing of the carbon sources through the Entner-Doudoroff pathway was required for carbon catabolite repression (Velázquez et al. 2004). Furthermore, interrupting this pathway by knocking out the eda gene for 2-dehydro-3-deoxyphosphogluconate aldolase increased the repressive effect of glucose, suggesting that the glucose metabolite 2-keto-3-deoxy-6-phosphogluconate (KDPG) was the cellular signal triggering PTS phosphorelay for Pu regulation, probably by controlling PtsN phosphorylation state (Velázquez et al. 2004; del Castillo and Ramos 2007). KDPG was also shown to be the signal triggering glucose-dependent carbon catabolite repression of phenylacetic acid metabolism in this strain (Kim et al. 2009). In addition, in P. putida the PTS system is interconnected with the transport system for fructose, a non-repressive carbon source (Pflüger and de Lorenzo 2008), suggesting that carbon flux through the Entner-Doudoroff pathway would be translated to the C/N PTS system through FruB (EIIAFru). The analysis of a series of P. putida metabolic mutants revealed that toluene and glucose exert a reciprocal repression of degradation pathways (del Castillo and Ramos 2007). Glucose repression of Pu expression requires ptsN, as it had previously been shown (Ruíz et al. 2004; Cases and de Lorenzo 2005). Interestingly, toluenemediated repression of glucose catabolism is mediated by the Crc protein, a global regulator that works together with the Hfq posttranscriptional regulator and controls carbon flow in Pseudomonas, playing a key role in catabolite repression (Collier et al. 1996; Rojo 2010). Although Crc was initially believed to directly bind RNA at a specific consensus sequence (termed the catabolite active motif), recent findings have shown that the actual role of Crc is to enhance and stabilize the complex formed between RNA and the RNA-binding protein Hfq. Hfq recognizes specific sites in the translation initiation region of target mRNAs, repressing translation (Sonnleitner and Blasi 2014). The effect of Crc stabilization of Hfq-RNA complexes at many target sites favors translational inhibition and explains the observed catabolite repression relief of crc mutants (Moreno et al. 2007, 2015; Madhusani et al. 2015). In P. putida, Crc, probably in cooperation with Hfq, has been shown to inhibit the assimilation of glucose and fructose in rich medium and to modulate the uptake of amino acids as well as their assimilation pathways, so that the use of the strain’s preferred amino acids is favored over the non-preferred ones (reviewed in Rojo 2010). Although a minor role of Crc in Pu repression in P. putida growing on rich medium had been described (Aranda-Olmedo et al. 2005), it was only after a thorough analysis of TOL pathway gene expression in a crc mutant that the role of this interesting protein in TOL pathway expression was elucidated (Moreno et al. 2010). In TOL, xylR and xylS mRNAs are targets for Crc/Hfq binding and repression of translation, which reduces the availability of these regulators in the cell, therefore tuning down overall expression of the pathway genes. Moreover, target sequences for Crc/Hfq were found overlapping the translation start site of several enzyme genes located in the upper and meta-cleavage polycistronic mRNAs, providing a mean to maintain an appropriate balance between the different pathway enzymes coded for in each messenger (Moreno et al. 2010).

14

P. Domínguez-Cuevas and S. Marqués

Finally, by assaying P. putida crude extracts, two proteins with the capacity to bind Pu promoter were identified. The LytTR family two-component response regulator PprA, with no known function in P. putida, is able to bind Pu promoter at a site overlapping the UASs for XylR binding, thus competing with this regulator (Vitale et al. 2008). As expected, in vivo the presence of PprA had a repressive effect on Pu expression. The signals sensed by PprA to modulate Pu expression are unknown so far. TurA, a small protein with structural homology to the nucleoid protein H-NS, was also shown to bind Pu and repress expression from this promoter, an effect that was strengthened at low temperature (Rescalli et al. 2004). This repression is unrelated to the previously described exponential silencing or catabolite repression control of Pu promoter. The targets of the global regulators of xyl pathway expression discussed above are summarized in Fig. 3.

5

Research Needs

After many years of thorough research, TOL plasmid catabolic pathways can be considered a perfect example of transcription regulation in bacteria. Future studies will focus on the subtleties of the system, with the aim to identify the minute processes involved in the key steps in transcription activation mechanisms, which will help us understand transcription regulation in bacteria. This will include the basic mechanisms leading to σ54 promoter activation, the contacts established between XylS and the two RNAP at the Pm promoter to initiate transcription, and the identification and analysis of new posttranscriptional regulation mechanisms (Velázquez et al. 2005). Understanding the processes underlying global control will definitely be the subject of thorough analysis in the future, especially to determine the role of the C/N PTS system, the final target of this phosphorelay and the possible additional elements involved. The behavior and performance of the system under natural conditions such as those found in real polluted environments will probably rely on the multiple signals that the bacteria encounter in these habitats, which are expected to be essentially different to those acting under the laboratory conditions analyzed so far. In this sense, our current knowledge of TOL pathway functioning is consistent enough to represent a perfect model to analyze the response of this biotechnologically relevant strain under real environment biodegradation conditions. Attempts in this sense are already underway (Svenningsen et al. 2016). A general issue barely addressed to date is how highly hydrophobic signal compounds such as hydrocarbons are spread in aqueous systems, how they approach and enter the cell, and how they find the regulator in the cytoplasm. In this sense, the physical channeling of TOL metabolites in the cytoplasm has been recently suggested (Kim et al. 2016) and deserves further analysis. Also, the role of biofilms and secreted proteins in the capacity of hydrocarbonoclastic bacteria to degrade hydrocarbons has been highlighted recently (Ennouri et al. 2016; Espinosa-Urgel and Marqués 2016), opening up new perspectives in the analysis of aromatic hydrocarbon degradation.

Current View of The Mechanisms Controlling The Transcription of The TOL. . .

15

FtsH TurA IHF PprA

σ38

σ54 PtsN

σ32

Pu

upper

Crc/Hfq

Pm

HU σ70 σ54 PtsN IHF σ70

meta-cleavage

Crc/Hfq

xylS PS2 PS1

Crc/Hfq

PR1

PR2

xylR

Crc/Hfq

Fig. 3 Current knowledge on global regulatory proteins involved in the control of TOL system at either the transcriptional or the translational level. Green arrows indicate positive effect over transcription. Red blunt-headed lines indicate negative effect over transcription, except for Crc and Hfq that exert their control posttranscriptionally, inhibiting mRNA translation in response certain carbon sources. The mRNA transcripts are depicted as a gray curved line

As evidenced throughout this review, the regulatory network of the TOL plasmid aromatic degradation pathway has become a paradigm of transcriptional regulation in bacteria and a model of integration of a horizontally acquired pathway in the host metabolism. The TOL regulatory network as a whole, as well as independent parts, has been extensively exploited with biotechnological purposes, in particular in the design of efficient biosensors and expression systems (Marqués et al. 2006; de las Heras and de Lorenzo 2011; de las Heras et al. 2012, 2015; Balzer et al. 2013). Furthermore, it has proven to be an excellent tool to develop logic models to explain the network layout and dynamics and to predict the circuit’s behavior upon different metabolic challenges using system biology approaches (Silva-Rocha et al. 2011, 2013). Acknowledgments This work was supported by the European Regional Development Fund FEDER and grant from the Spanish Ministry of Economy and Competitiveness (BIO2014-54361R).

References Abril MA, Buck M, Ramos JL (1991) Activation of the Pseudomonas TOL plasmid upper pathway operon. Identification of binding sites for the positive regulator XylR and for integration host factor protein. J Biol Chem 266:15832–15838 Aranda-Olmedo I, Ramos JL, Marqués S (2005) Integration of signals through Crc and PtsN in catabolite repression of Pseudomonas putida TOL plasmid pWW0. Appl Environ Microbiol 71:4191–4198 Aranda-Olmedo I, Marín P, Ramos JL, Marqués S (2006) Role of the ptsN gene product in catabolite repression of the Pseudomonas putida TOL toluene degradation pathway in chemostat cultures. Appl Environ Microbiol 72:7418–7421 Assinder SJ, Williams PA (1990) The TOL plasmids: determinants of the catabolism of toluene and the xylenes. Adv Microb Physiol 31:1–69

16

P. Domínguez-Cuevas and S. Marqués

Balzer S, Kucharova V, Megerle J, Lale R, Brautaset T, Valla S (2013) A comparative analysis of the properties of regulated promoter systems commonly used for recombinant gene expression in Escherichia coli. Microb Cell Factories 12:26 Bertoni G, Marqués S, De Lorenzo V (1998) Activation of the toluene-responsive regulator XylR causes a transcriptional switch between sigma54 and sigma70 promoters at the divergent Pr/Ps region of the TOL plasmid. Mol Microbiol 27:651–659 Bordes P, Wigneshweraraj SR, Schumacher J, Zhang X, Chaney M, Buck M (2003) The ATP hydrolyzing transcription activator phage shock protein F of Escherichia coli: identifying a surface that binds sigma 54. Proc Natl Acad Sci U S A 100:2278–2283 Bose D, Joly N, Pape T, Rappas M, Schumacher J, Buck M, Zhang X (2008) Dissecting the ATP hydrolysis pathway of bacterial enhancer-binding proteins. Biochem Soc Trans 36:83–88 Bush M, Dixon R (2012) The role of bacterial enhancer binding proteins as specialized activators of sigma54-dependent transcription. Microbiol Mol Biol Rev 76:497–529 Carmona M, De Lorenzo V (1999) Involvement of the FtsH (HflB) protease in the activity of sigma 54 promoters. Mol Microbiol 31:261–270 Carmona M, Rodríguez MJ, Martínez-Costa O, De Lorenzo V (2000) In vivo and in vitro effects of (p)ppGpp on the sigma(54) promoter Pu of the TOL plasmid of Pseudomonas putida. J Bacteriol 182:4711–4718 Cases I, De Lorenzo V (2005) Promoters in the environment: transcriptional regulation in its natural context. Nat Rev Microbiol 3:105–118 Cases I, Perez-Martin J, De Lorenzo V (1999) The IIANtr (PtsN) protein of Pseudomonas putida mediates the C source inhibition of the sigma54-dependent Pu promoter of the TOL plasmid. J Biol Chem 274:15562–15568 Cases I, Velázquez F, De Lorenzo V (2007) The ancestral role of the phosphoenolpyruvatecarbohydrate phosphotransferase system (PTS) as exposed by comparative genomics. Res Microbiol 158:666–670 Chen B, Sysoeva TA, Chowdhury S, Nixon BT (2008) Regulation and action of the bacterial enhancer-binding protein AAA+ domains. Biochem Soc Trans 36:89–93 Coggan KA, Wolfgang MC (2012) Global regulatory pathways and cross-talk control pseudomonas aeruginosa environmental lifestyle and virulence phenotype. Curr Issues Mol Biol 14:47–70 Collier DN, Hager PW, Phibbs PV Jr (1996) Catabolite repression control in the Pseudomonas. Res Microbiol 147:551–561 Cowles CE, Nichols NN, Harwood CS (2000) BenR, a XylS homologue, regulates three different pathways of aromatic acid degradation in Pseudomonas putida. J Bacteriol 182:6339–6346 Daniels C, Del Castillo T, Krell T, Segura A, Busch A, Lacal J, Ramos JL (2008) Cellular ecophysiology: genetics and genomics of accessing and exploiting hydrocarbons. In: Timmis KN (ed) Microbiology of hydrocarbons, oils, lipids, and derived compounds. Springer, Heidelberg Daniels C, Godoy P, Duque E, Molina-Henares MA, De La Torre J, Del Arco JM, Herrera C, Segura A, Guazzaroni ME, Ferrer M, Ramos JL (2010) Global regulation of food supply by Pseudomonas putida DOT-T1E. J Bacteriol 192:2169–2181 De Las Heras A, De Lorenzo V (2011) Cooperative amino acid changes shift the response of the sigma 54-dependent regulator XylR from natural m-xylene towards xenobiotic 2,4-dinitrotoluene. Mol Microbiol 79:1248–1259 De Las Heras A, Fraile S, De Lorenzo V (2012) Increasing signal specificity of the TOL network of Pseudomonas putida mt-2 by rewiring the connectivity of the master regulator XylR. PLoS Genet 8(10):e1002963. doi: 10.1371/journal.pgen.1002963 De Las Heras A, Martinez-Garcia E, Domingo-Sananes MR, De Lorenzo V (2015) Widening functional boundaries of the sigma(54) promoter Pu of Pseudomonas putida by defeating extant physiological constraints. Mol Biosys 11:734–742 De Lorenzo V, Herrero M, Metzke M, Timmis KN (1991) An upstream XylR and IHF induced nucleoprotein complex regulates the σ54-dependent Pu promotor of TOL plasmid. EMBO J 10:1159–1167

Current View of The Mechanisms Controlling The Transcription of The TOL. . .

17

De Lorenzo V, Sekowska A, Danchin A (2015) Chemical reactivity drives spatiotemporal organisation of bacterial metabolism. FEMS Microbiol Rev 39:96–119 Del Castillo T, Ramos JL (2007) Simultaneous catabolite repression between glucose and toluene metabolism in Pseudomonas putida is channeled through different signaling pathways. J Bacteriol 189:6602–6610 Delgado A, Ramos JL (1994) Genetic evidence for activation of the positive transcriptional regulator XylR, a member of the NtrC family of regulators, by effector binding. J Biol Chem 269:8059–8062 Deutscher J, Francke C, Postma PW (2006) How phosphotransferase system-related protein phosphorylation regulates carbohydrate metabolism in bacteria. Microbiol Mol Biol Rev 70:939–1031 Devos D, Garmendia J, De Lorenzo V, Valencia A (2002) Deciphering the action of aromatic effectors on the prokaryotic enhancer-binding protein XylR: a structural model of its N-terminal domain. Environ Microbiol 4:29–41 Di Martino ML, Romilly C, Wagner EG, Colonna B, Prosseda G (2016) One gene and two proteins: a leaderless mRNA supports the translation of a shorter form of the Shigella VirF regulator. MBio 7(6):e01860–16. doi:10.1128/mBio.01860-16. Domínguez-Cuevas P, Marqués S (2010) Transcriptional control of the TOL plasmid pathways. In: Timmis K (ed) Handbook of hydrocarbon and lipid microbiology. Spinger, Berlin/Heidelberg, pp 1130–1136 Domínguez-Cuevas P, Marín P, Ramos JL, Marqués S (2005) RNA polymerase holoenzymes can share a single transcription start site for the Pm promoter. Critical nucleotides in the
7 to
18 region are needed to select between RNA polymerase with sigma38 or sigma32. J Biol Chem 280:41315–41323 Domínguez-Cuevas P, González-Pastor JE, Marqués S, Ramos JL, De Lorenzo V (2006) Transcriptional tradeoff between metabolic and stress-response programs in Pseudomonas putida KT2440 cells exposed to toluene. J Biol Chem 281:11981–11991 Domínguez-Cuevas P, Marín P, Busby S, Ramos JL, Marqués S (2008a) Roles of effectors in XylSdependent transcription activation: intramolecular domain derepression and DNA binding. J Bacteriol 190:3118–3128 Domínguez-Cuevas P, Marín P, Marqués S, Ramos JL (2008b) XylS-Pm promoter interactions through two helix-turn-helix motifs: identifying XylS residues important for DNA binding and activation. J Mol Biol 375:59–69 Domínguez-Cuevas P, Ramos J-L, Marqués S (2010) Sequential XylS-CTD binding to the Pm promoter induces DNA bending prior to activation. J Bacteriol 192:2682–2690 Duetz WA, Marqués S, Wind B, Ramos JL, Van Andel JG (1996) Catabolite repression of the toluene degradation pathway in Pseudomonas putida harboring pWW0 under various conditions of nutrient limitation in chemostat culture. Appl Environ Microbiol 62:601–606 Ennouri H, D’abzac P, Hakil F, Branchu P, Naitali M, Lomenech AM, Oueslati R, Desbrieres J, Sivadon P, Grimaud R (2016) The extracellular matrix of the oleolytic biofilms of Marinobacter hydrocarbonoclasticus comprises cytoplasmic proteins and T2SS effectors that promote growth on hydrocarbons and lipids. Environ Microbiol 19(1):159–173 Espinosa-Urgel M, Marqués S (2016) New insights in the early extracellular events in hydrocarbon and lipid biodegradation. Environ Microbiol 19(1):15–18 Fernández S, De Lorenzo V, Pérez-Martín J (1995) Activation of the transcriptional regulator XylR of Pseudomonas putida by release of repression between functional domains. Mol Microbiol 16:205–213 Gallegos MT, Marqués S, Ramos JL (1996a) Expression of the TOL plasmid xylS gene in Pseudomonas putida occurs from a alpha 70-dependent promoter or from alpha 70- and alpha 54-dependent tandem promoters according to the compound used for growth. J Bacteriol 178:2356–2361 Gallegos MT, Marqués S, Ramos JL (1996b) The TACAN4TGCA motif upstream from the
35 region in the sigma70-sigmaS-dependent Pm promoter of the TOL plasmid is the minimum

18

P. Domínguez-Cuevas and S. Marqués

DNA segment required for transcription stimulation by XylS regulators. J Bacteriol 178:6427–6434 Gallegos MT, Schleif R, Bairoch A, Hofmann K, Ramos JL (1997) Arac/XylS family of transcriptional regulators. Microbiol Mol Biol Rev 61:393–410 Garmendia J, De Lorenzo V (2000) The role of the interdomain B linker in the activation of the XylR protein of Pseudomonas putida. Mol Microbiol 38:401–410 Garmendia J, Devos D, Valencia A, De Lorenzo V (2001) A la carte transcriptional regulators: unlocking responses of the prokaryotic enhancer-binding protein XylR to non-natural effectors. Mol Microbiol 42:47–59 Garmendia J, De Las Heras A, Galvao TC, De Lorenzo V (2008) Tracing explosives in soil with transcriptional regulators of Pseudomonas putida evolved for responding to nitrotoluenes. Microb Biotechnol 1:236–246 González-Pérez MM, Marqués S, Domínguez-Cuevas P, Ramos JL (2002) XylS activator and RNA polymerase binding sites at the Pm promoter overlap. FEBS Lett 519:117–122 Greated A, Lambertsen L, Williams PA, Thomas CM (2002) Complete sequence of the IncP-9 TOL plasmid pWW0 from Pseudomonas putida. Environ Microbiol 4:856–871 Gross CA, Chan C, Dombroski A, Gruber T, Shasrp M, Tupy J, Young B (1998) The functional and regulatory roles of sigma factors in transcription. Cold Spring Harb Symp Quant Biol 63:141–156 Hengge-Aronis R (2002) Signal transduction and regulatory mechanisms involved in control of the sigma(S) (RpoS) subunit of RNA polymerase. Microbiol Mol Biol Rev 66:373–395 Herrera MC, Daddaoua A, Fernández-Escamilla A, Ramos JL (2012) Involvement of the global Crp regulator in cyclic AMP-dependent utilization of aromatic amino acids by Pseudomonas putida. J Bacteriol 194:406–412 Holtel A, Abril MA, Marqués S, Timmis KN, Ramos JL (1990) Promoter upstream activator sequences are required for expression of the xylS gene and upper pathway operon on the Pseudomonas TOL plasmid. Mol Microbiol 4:1551–1556 Holtel A, Timmis KN, Ramos JL (1992) Upstream binding sequences of the XylR activator protein and integration host factor in the xylS gene promoter region of the Pseudomonas TOL plasmid. Nucleic Acids Res 20:1755–1762 Holtel A, Goldenberg D, Giladi H, Oppenheim AB, Timmis KN (1995) Involvement of IHF protein in expression of the Ps promoter of the Pseudomonas putida TOL plasmid. J Bacteriol 177:3312–3315 Hugouvieux-Cotte-Pattat N, Kohler T, Rekik M, Harayama S (1990) Growth-phase-dependent expression of the Pseudomonas putida TOL plasmid pWW0 catabolic genes. J Bacteriol 172:6651–6660 Huo YX, Tian ZX, Rappas M, Wen J, Chen YC, You CH, Zhang X, Buck M, Wang YP, Kolb A (2006) Protein-induced DNA bending clarifies the architectural organization of the sigma54dependent glnAp2 promoter. Mol Microbiol 59:168–180 Inouye S, Nakazawa A, Nakazawa T (1987) Expression of the regulatory gene xylS on the TOL pasmid is positively controlled by the xylR gene product. Proc Natl Acad Sci U S A 84:5182–5186 Inouye S, Nakazawa A, Nakazawa T (1988) Nucleotide sequence of the regulatory gene xylR of the TOL plasmid from Pseudomonas putida. Gene 66:301–306 Jiménez JI, Pérez-Pantoja D, Chavarréa M, Díaz E, De Lorenzo V (2014) A second chromosomal copy of the catA gene endows Pseudomonas putida mt-2 with an enzymatic safety valve for excess of catechol. Environ Microbiol 16:1767–1778 Kessler B, Marqués S, Köhler T, Ramos JL, Timmis KN, De Lorenzo V (1994) Cross talk between catabolic pathways in Pseudomonas putida: XylS-dependent and -independent activation of the TOL meta operon requires the same cis-acting sequences within the Pm promoter. J Bacteriol 176:5578–5582

Current View of The Mechanisms Controlling The Transcription of The TOL. . .

19

Kim J, Yeom J, Jeon CO, Park W (2009) Intracellular 2-keto-3-deoxy-6-phosphogluconate is the signal for carbon catabolite repression of phenylacetic acid metabolism in Pseudomonas putida KT2440. Microbiology 155:2420–2428 Kim J, Pérez-Pantoja D, Silva-Rocha R, Oliveros JC, De Lorenzo V (2016) High-resolution analysis of the m-xylene/toluene biodegradation subtranscriptome of Pseudomonas putida mt-2. Environ Microbiol 18:3327–3341 Koutinas M, Lam M-C, Kiparissides A, Silva-Rocha R, Godinho M, Livingston AG, Pistikopoulos EN, De Lorenzo V, Dos Santos VAPM, Mantalaris A (2010) The regulatory logic of m-xylene biodegradation by Pseudomonas putida mt-2 exposed by dynamic modelling of the principal node Ps/Pr of the TOL plasmid. Environ Microbiol 12:1705–1718 Kwon HJ, Bennik MH, Demple B, Ellenberger T (2000) Crystal structure of the Escherichia coli Rob transcription factor in complex with DNA. Nat Struct Biol 7:424–430 Lee SY, De La Torre A, Yan D, Kustu S, Nixon BT, Wemmer DE (2003) Regulation of the transcriptional activator NtrC1: structural studies of the regulatory and AAA+ ATPase domains. Genes Dev 17:2552–2563 Madhushani A, Del Peso-Santos T, Moreno R, Rojo F, Shingler V (2015) Transcriptional and translational control through the 50 -leader region of the dmpR master regulatory gene of phenol metabolism. Environ Microbiol 17:119–133 Marqués S, Ramos JL (1993) Transcriptional control of the Pseudomonas putida TOL plasmid catabolic pathways. Mol Microbiol 9:923–929 Marqués S, Holtel A, Timmis KN, Ramos JL (1994) Transcriptional induction kinetics from the promoters of the catabolic pathways of TOL plasmid pWW0 of Pseudomonas putida for metabolism of aromatics. J Bacteriol 176:2517–2524 Marqués S, Gallegos MT, Ramos JL (1995) Role of sigma S in transcription from the positively controlled Pm promoter of the TOL plasmid of Pseudomonas putida. Mol Microbiol 18:851–857 Marqués S, Gallegos MT, Manzanera M, Holtel A, Timmis KN, Ramos JL (1998) Activation and repression of transcription at the double tandem divergent promoters for the xylR and xylS genes of the TOL plasmid of Pseudomonas putida. J Bacteriol 180:2889–2894 Marqués S, Manzanera M, González-Pérez MM, Gallegos MT, Ramos JL (1999) The XylSdependent Pm promoter is transcribed in vivo by RNA polymerase with sigma 32 or sigma 38 depending on the growth phase. Mol Microbiol 31:1105–1113 Marqués S, Aranda-Olmedo I, Ramos JL (2006) Controlling bacterial physiology for optimal expression of gene reporter constructs. Curr Opin Biotechnol 17:50–56 Martinez-Laguna Y, Calva E, Puente JL (1999) Autoactivation and environmental regulation of bfpT expression, the gene coding for the transcriptional activator of bfpA in enteropathogenic Escherichia coli. Mol Microbiol 33:153–166 Michán C, Zhou L, Gallegos MT, Timmis KN, Ramos JL (1992) Identification of critical aminoterminal regions of XylS. The positive regulator encoded by the TOL plasmid. J Biol Chem 267:22897–22901 Milanesio P, Arce-Rodríguez A, Muñoz A, Calles B, De Lorenzo V (2011) Regulatory exaptation of the catabolite repression protein (Crp)-cAMP system in Pseudomonas putida. Environ Microbiol 13:324–339 Moreno R, Ruiz-Manzano A, Yuste L, Rojo F (2007) The Pseudomonas putida Crc global regulator is an RNA binding protein that inhibits translation of the AlkS transcriptional regulator. Mol Microbiol 64:665–675 Moreno R, Fonseca P, Rojo F (2010) The Crc global regulator inhibits the Pseudomonas putida pWW0 toluene/xylene assimilation pathway by repressing the translation of regulatory and structural genes. J Biol Chem 285:24412–24419 Moreno R, Hernández-Arranz S, La Rosa R, Yuste L, Madhushani A, Shingler V, Rojo F (2015) The Crc and Hfq proteins of Pseudomonas putida cooperate in catabolite repression and formation of ribonucleic acid complexes with specific target motifs. Environ Microbiol 17:105–118

20

P. Domínguez-Cuevas and S. Marqués

Morin N, Tirling C, Ivison SM, Kaur AP, Nataro JP, Steiner TS (2010) Autoactivation of the AggR regulator of enteroaggregative Escherichia coli in vitro and in vivo. FEMS Immunol Med Microbiol 58:344–355 Neuwald AF, Aravind L, Spouge JL, Koonin EV (1999) AAA+: a class of chaperone-like ATPases associated with the assembly, operation, and disassembly of protein complexes. Genome Res 9:27–43 Pérez-Martín J, De Lorenzo V (1995a) The amino-terminal domain of the prokaryotic enhancerbinding protein XylR is a specific intramolecular repressor. Proc Natl Acad Sci U S A 92:9392–9396 Pérez-Martín J, De Lorenzo V (1995b) The sigma 54-dependent promoter Ps of the TOL plasmid of Pseudomonas putida requires HU for transcriptional activation in vivo by XylR. J Bacteriol 177:3758–3763 Pérez-Martín J, De Lorenzo V (1996a) In vitro activities of an N-terminal truncated form of XylR, a sigma 54-dependent transcriptional activator of Pseudomonas putida. J Mol Biol 258:575–587 Pérez-Martín J, De Lorenzo V (1996b) ATP binding to the sigma 54-dependent activator XylR triggers a protein multimerization cycle catalyzed by UAS DNA. Cell 86:331–339 Pérez-Pantoja D, Kim J, Silva-Rocha R, De Lorenzo V (2015) The differential response of the Pben promoter of Pseudomonas putida mt-2 to BenR and XylS prevents metabolic conflicts in m-xylene biodegradation. Environ Microbiol 17:64–75 Pflüger K, De Lorenzo V (2008) Evidence of in vivo cross talk between the nitrogen-related and fructose-related branches of the carbohydrate phosphotransferase system of Pseudomonas putida. J Bacteriol 190:3374–3380 Porter ME, Mitchell P, Roe AJ, Free A, Smith DG, Gally DL (2004) Direct and indirect transcriptional activation of virulence genes by an AraC-like protein, PerA from enteropathogenic Escherichia coli. Mol Microbiol 54:1117–1133 Ramos JL, Stolz A, Reineke W, Timmis KN (1986) Altered effector specificities in regulators of gene expression: TOL plasmid xylS mutants and their use to engineer expansion of the range of aromatics degraded by bacteria. Proc Natl Acad Sci U S A 83:8467–8471 Ramos JL, Michán C, Rojo F, Dwyer D, Timmis K (1990) Signal-regulator interactions. Genetic analysis of the effector binding site of xylS, the benzoate-activated positive regulator of Pseudomonas TOL plasmid meta-cleavage pathway operon. J Mol Biol 211:373–382 Ramos JL, Marqués S, Timmis KN (1997) Transcriptional control of the Pseudomonas TOL plasmid catabolic operons is achieved through an interplay of host factors and plasmid- encoded regulators. Annu Rev Microbiol 51:341–373 Rappas M, Schumacher J, Beuron F, Niwa H, Bordes P, Wigneshweraraj S, Keetch CA, Robinson CV, Buck M, Zhang X (2005) Structural insights into the activity of enhancer-binding proteins. Science 307:1972–1975 Ray S, Gunzburg M, Wilce M, Panjikar S, Anand R (2016) Structural basis of selective aromatic pollutant sensing by the effector binding domain of MopR, an NtrC family transcriptional regulator. ACS Chem Biol 11:2357–2365 Rescalli E, Saini S, Bartocci C, Rychlewski L, De Lorenzo V, Bertoni G (2004) Novel physiological modulation of the Pu promoter of TOL plasmid: negative regulatory role of the TurA protein of Pseudomonas putida in the response to suboptimal growth temperatures. J Biol Chem 279:7777–7784 Rhee S, Martin RG, Rosner JL, Davies DR (1998) A novel DNA-binding motif in MarA: the first structure for an AraC family transcriptional activator. Proc Natl Acad Sci U S A 95:10413–10418 Rodgers ME, Holder ND, Dirla S, Schleif R (2009) Functional modes of the regulatory arm of AraC. Proteins 74:81–91 Rojo F (2010) Carbon catabolite repression in Pseudomonas: optimizing metabolic versatility and interactions with the environment. FEMS Microbiol Rev 34:658–684 Ruíz R, Ramos JL (2002) Residues 137 and 153 at the N terminus of the XylS protein influence the effector profile of this transcriptional regulator and the sigma factor used by RNA polymerase to stimulate transcription from its cognate promoter. J Biol Chem 277:7282–7286

Current View of The Mechanisms Controlling The Transcription of The TOL. . .

21

Ruíz R, Marqués S, Ramos JL (2003) Leucines 193 and 194 at the N-terminal domain of the XylS protein, the positive transcriptional regulator of the TOL meta-cleavage pathway, are involved in dimerization. J Bacteriol 185:3036–3041 Ruíz R, Aranda-Olmedo MI, Domínguez-Cuevas P, Ramos-González MI, Marqués S (2004) Transcriptional regulation of the toluene catabolic pathways. In: Ramos JL (ed) Pseudomonas. Kluwer Academic/Plenum Publishers, London, pp 509–537 Sallai L, Tucker PA (2005) Crystal structure of the central and C-terminal domain of the sigma(54)activator ZraR. J Struct Biol 151:160–170 Salto R, Delgado A, Michan C, Marques S, Ramos JL (1998) Modulation of the function of the signal receptor domain of XylR, a member of a family of prokaryotic enhancer-like positive regulators. J Bacteriol 180:600–604 Santiago AE, Ruiz-Perez F, Jo NY, Vijayakumar V, Gong MQ, Nataro JP (2014) A large family of antivirulence regulators modulates the effects of transcriptional activators in Gram-negative pathogenic bacteria. PLoS Pathog 10:e1004153 Santiago AE, Yan MB, Tran M, Wright N, Luzader DH, Kendall MM, Ruiz-Perez F, Nataro JP (2016) A large family of anti-activators accompanying XylS/AraC family regulatory proteins. Mol Microbiol 101:314–332 Schumacher J, Joly N, Rappas M, Zhang X, Buck M (2006) Structures and organisation of AAA+ enhancer binding proteins in transcriptional activation. J Struct Biol 156:190–199 Seedorff J, Schleif R (2011) Active role of the interdomain linker of AraC. J Bacteriol 193:5737–5746 Shingler V, Pavel H (1995) Direct regulation of the ATPase activity of the transcriptional activator DmpR by aromatic compounds. Mol Microbiol 17:505–513 Silva-Rocha R, De Lorenzo V (2012) Broadening the signal specificity of prokaryotic promoters by modifying cis-regulatory elements associated with a single transcription factor. Mol Biosys 8:1950–1957 Silva-Rocha R, De Jong H, Tamames J, De Lorenzo V (2011) The logic layout of the TOL network of Pseudomonas putida pWW0 plasmid stems from a metabolic amplifier motif (MAM) that optimizes biodegradation of m-xylene. BMC Syst Biol 5:591 Silva-Rocha R, Perez-Pantoja D, De Lorenzo V (2013) Decoding the genetic networks of environmental bacteria: regulatory moonlighting of the TOL system of Pseudomonas putida mt-2. ISME J 7:229–232 Sonnleitner E, Blasi U (2014) Regulation of Hfq by the RNA CrcZ in Pseudomonas aeruginosa carbon catabolite repression. PLoS Genet 10:e1004440 Studholme DJ, Dixon R (2003) Domain architectures of sigma54-dependent transcriptional activators. J Bacteriol 185:1757–1767 Suh SJ, Runyen-Janecky LJ, Maleniak TC, Hager P, Macgregor CH, Zielinski-Mozny NA, Phibbs PVJ, West SE (2002) Effect of vfr mutation on global gene expression and catabolite repression control of Pseudomonas aeruginosa. Microbiology 148:1561–1569 Svenningsen NB, Nicolaisen MH, Hansen HC, De Lorenzo V, Nybroe O (2016) Nitrogen regulation of the xyl genes of Pseudomonas putida mt-2 propagates into a significant effect of nitrate on m-xylene mineralization in soil. Microb Biotechnol 9:814–823 Sze CC, Shingler V (1999) The alarmone (p)ppGpp mediates physiological-responsive control at the sigma 54-dependent Po promoter. Mol Microbiol 31:1217–1228 Sze CC, Bernardo LM, Shingler V (2002) Integration of global regulation of two aromaticresponsive sigma(54)-dependent systems: a common phenotype by different mechanisms. J Bacteriol 184:760–770 Tobes R, Ramos JL (2002) AraC-XylS database: a family of positive transcriptional regulators in bacteria. Nucleic Acids Res 30:318–321 Tsipa A, Koutinas M, Pistikopoulos EN, Mantalaris A (2016) Transcriptional kinetics of the crosstalk between the ortho-cleavage and TOL pathways of toluene biodegradation in Pseudomonas putida mt-2. J Biotechnol 228:112–123

22

P. Domínguez-Cuevas and S. Marqués

Valls M, Buckle M, De Lorenzo V (2002) In vivo UV laser footprinting of the Pseudomonas putida sigma 54 Pu promoter reveals that integration host factor couples transcriptional activity to growth phase. J Biol Chem 277:2169–2175 Velázquez F, Di Bartolo I, De Lorenzo V (2004) Genetic evidence that catabolites of the EntnerDoudoroff pathway signal C source repression of the sigma54 Pu promoter of Pseudomonas putida. J Bacteriol 186:8267–8275 Velázquez F, Parro V, De Lorenzo V (2005) Inferring the genetic network of m-xylene metabolism through expression profiling of the xyl genes of Pseudomonas putida mt-2. Mol Microbiol 57:1557–1569 Velázquez F, Fernández S, De Lorenzo V (2006) The upstream-activating sequences of the sigma54 promoter Pu of Pseudomonas putida filter transcription readthrough from upstream genes. J Biol Chem 281:11940–11948 Vidangos N, Maris AE, Young A, Hong E, Pelton JG, Batchelor JD, Wemmer DE (2013) Structure, function, and tethering of DNA-binding domains in sigma(5)(4) transcriptional activators. Biopolymers 99:1082–1096 Vitale E, Milani A, Renzi F, Galli E, Rescalli E, De Lorenzo V, Bertoni G (2008) Transcriptional wiring of the TOL plasmid regulatory network to its host involves the submission of the sigma (54)-promoter Pu to the response regulator PprA. Mol Microbiol 69:698–713 Wigneshweraraj S, Bose D, Burrows PC, Joly N, Schumacher J, Rappas M, Pape T, Zhang X, Stockley P, Severinov K, Buck M (2008) Modus operandi of the bacterial RNA polymerase containing the sigma54 promoter-specificity factor. Mol Microbiol 68:538–546 Williams PA, Murray K (1974) Metabolism of benzoate and the methylbenzoates by Pseudomonas putida (arvilla) mt-2: evidence for the existence of a TOL plasmid. J Bacteriol 120:416–423 Zhang Y-T, Jiang F, Tian Z-X, Huo Y-X, Sun Y-C, Wang Y-P (2014) CRP-Cyclic AMP dependent inhibition of the xylene-responsive sigma(54)-Promoter Pu in Escherichia coli. Plos One 9: e86727. doi: 10.1371/journal.pone.0086727 Zwick F, Lale R, Valla S (2013) Regulation of the expression level of transcription factor XylS reveals new functional insight into its induction mechanism at the Pm promoter. BMC Microbiol 13:262

Genetics and Biochemistry of Biphenyl and PCB Biodegradation Loreine Agulló, Dietmar H. Pieper, and Michael Seeger

Abstract

Microorganisms are crucial for the removal of polychlorinated biphenyls (PCBs) from polluted environments. Microbial anaerobic dehalogenation of highly and moderately chlorinated biphenyls generates the subsequent less chlorinated congeners. Microbial aerobic degradation performed by enzymes of the biphenyl (bph) upper and lower pathways oxidizes moderately and low chlorinated biphenyls. These enzymes and their substrate specificities are discussed in Sect. 2.1. Biphenyl 2,3-dioxgenases (BDOs) are key enzymes of biphenyl pathways, which determine substrate range and extent of PCB degradation. In addition, the specificity of subsequent enzymes is also crucial for productive metabolism. Specific native and engineered BDOs possess a wide range of substrates, which permit their application for synthesis of fine organic chemicals including novel bioactive compounds. The metabolism of PCBs is described in detail for some model organisms, and the genetic organization of gene clusters of model organisms is described in Sect. 2.2. The sequenced genomes of some PCB-metabolizing organisms including the model strains Burkholderia xenovorans LB400 and Rhodococcus jostii RHA1 improve the understanding of their overall metabolism, physiology, and evolution as described in Sect. 2.3. This has also allowed a better evaluation into genome and proteome-wide defenses against PCB toxicity, which is summarized in Sect. 2.4. However, our knowledge on enzymes and genes involved in PCB metabolism is still rather fragmentary and an overview of

L. Agulló • M. Seeger (*) Laboratorio de Microbiología Molecular y Biotecnología Ambiental, Department of Chemistry & Center for Nanotechnology and Systems Biology & Centro de Biotecnología, Universidad Técnica Federico Santa María, Valparaíso, Chile e-mail: [email protected] D.H. Pieper Microbial Interactions and Processes Research Group, HZI – Helmholtz Centre for Infection Research, Braunschweig, Germany # Springer International Publishing AG 2017 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils and Lipids, Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-319-39782-5_30-1

1

2

L. Agulló et al.

the diversity of enzymes reported and mosaic routes is given in Sect. 2.5. Finally, strategies to optimize microorganisms for improved PCB degradation and bioremediation processes are discussed in Sects. 2.6 and 2.7.

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Aerobic Metabolism of PCBs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1 Upper Pathway Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Archetype bph Gene Clusters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Genome Analyses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4 Toxicity of PCBs and Their Metabolites and Bacterial Stress Response . . . . . . . . . . . . . 2.5 Metabolic Versatility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6 Optimized Enzymes and PCB-Degrading Organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7 Bioremediation of PCBs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1

2 4 4 11 12 14 15 17 19 20 21

Introduction

Preserving the environment for future generations is a main aim for sustainable development. The industrialization of many regions of the world has increased the environmental pollution. The removal of pollutants from the environment and the recovery of contaminated sites are major challenges of the XXI century. The Stockholm Convention of 2001 promotes the worldwide reduction and elimination of the emission of persistent organic pollutants (POPs) into the environment. PCBs, which are widely distributed in the environment, mainly in aquatic and soil ecosystems (Gomez-Gutiérrez et al. 2007; Palma-Fleming et al. 2008) were classified in the list of the 12 POPs for priority action. Biphenyl is an aromatic compound of two bound benzene rings, which occurs naturally in coal tar, crude oil, and natural gas. The industrial chlorination of biphenyl produces a mixture of PCBs carrying one to 10 chlorine atoms. There are 209 PCB congeners that differ in position and number of the chlorines. Industrial applications of PCBs started in 1929 in the USA by Monsanto. These compounds were not only used mainly as dielectric fluids in capacitors and transformers but also as flame retardants, plasticizers, and ink solvents. Commercial mixtures typically consisting of 40–70 congeners were sold under trade names as Askarel and Aroclor (Monsanto, USA, Canada, and UK), Clophen (Bayer, Germany), Kanechlor (Kanegafuchi, Japan), Phenoclor (Prodelec, France, and Spain), and Sovol and Sovtol (Orgsteklo, Orgsintez, former Soviet Union). More than 1.7 million tons of PCBs were produced worldwide (Stockholm Convention), and an important amount of these compounds have been released into the environment (Pieper and Seeger 2008).

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

3

Although adverse health effects were first recorded in the 1930s (Drinker et al. 1937), PCBs continued to be used for decades. Since then, PCBs have been shown to cause cancer (Mayes et al. 1998) and a number of serious effects on the immune, reproductive, nervous, and endocrine system (Faroon et al. 2001). Some coplanar PCBs have dioxin-like properties and are among the most toxic congeners. The toxicity, carcinogenicity, persistence, and tendency of PCBs to bioaccumulate are of increasing environmental and health concern in many countries (Pieper and Seeger 2008). In 2013 the International Agency for Research on Cancer (IARC) classified PCBs as human carcinogens (Lauby-Secretan et al. 2013). A study of a cohort of 24,865 capacitor-manufacturing workers exposed to polychlorinated biphenyls (PCBs) has associated PCB exposure with increased stomach, uterine, and prostate cancer and myeloma mortality (Ruder et al. 2014). Diverse full-scale applications of remediation technologies have been used for bioremediation of PCBs (Gomes et al. 2013). Technologies for the treatment of contaminated sites such as incineration, landfilling, thermal treatment (desorption, destruction, and vitrification), capping, and chemical dehalogenation are generally costly and usually involve dredging or excavation followed by disposal (Gomes et al. 2013; Fuentes et al. 2014). Microorganisms play a main role in the carbon cycle and in the removal of persistent organic pollutants from the environment. In situ and ex situ bioremediation has been applied successfully for the removal of petroleum contamination (Fuentes et al. 2014, 2015, 2016). For cleanup of PCB-contaminated environments, bioremediation is a promising technology (Pieper and Seeger 2008). Despite their chemical stability, diverse microbes have been reported as being capable to deal with PCBs and anaerobic consortia of microorganisms as well as aerobic bacteria biotransform or even mineralize PCBs. Generally, highly and moderately chlorinated PCBs are susceptible to a process termed reductive dehalogenation, in which PCBs are used as an alternative terminal electron acceptor in anaerobic respiration. The reductive dehalogenation of PCBs is congener-specific and, generally, involves selective dechlorination from para and meta positions, while chlorines at ortho position are preserved. However, ortho dechlorination of PCBs has also been reported. The first organisms capable to carry out such dehalogenations are available and belong to either the genus Dehalococcoides or Dehalobium (Cutter et al. 2001; Wu et al. 2002; Fennell et al. 2004). Dehalococcoides sp. strain CBDB1 is efficient for dechlorination of a wide range of PCB congeners in the environment (Adrian et al. 2009; Sowers and May 2013). Even though various reductive dehalogenases for dehalogenation of tetrachloroethene, vinyl chloride, or chlorobenzene (Neumann et al. 1996; Müller et al. 2004; Adrian et al. 2007) have been described, enzymes involved in reductive dehalogenation of PCBs remain to be identified. Lower and some moderately chlorinated PCBs are susceptible to aerobic metabolism.

4

2

L. Agulló et al.

Aerobic Metabolism of PCBs

Since the pioneering studies of Lunt and Evans (1970), diverse aerobic bacteria belonging to genera such as Pseudomonas, Burkholderia, Comamonas, Cupriavidus, Sphingomonas, Acidovorax, Rhodococcus, Arthrobacter, Acinetobacter, Corynebacterium, and Bacillus capable of using biphenyl as a sole source of carbon and energy and capable to oxidize PCBs have been described (Pieper and Seeger 2008). Diverse in situ assays have shown the biotransformation of a broad range of PCB congeners in the environmental (Xu et al. 2010; Tu et al. 2011; Pentyala et al. 2011; Gomes et al. 2013). Novel PCB degraders have been reported. Sinorhizobium meliloti biotransforms a broad range of PCB congeners (Tu et al. 2011).

2.1

Upper Pathway Enzymes

The dioxygenation of the aromatic ring constitutes the first step in the aerobic bacterial catabolism of aromatic compounds. This reaction destroys the aromatic system and functionalizes the molecule for further degradation (Furukawa, 2000; Overwin et al. 2015b). Based on the analysis of various biphenyl-degrading isolates, it could be deduced that, in general, lower chlorinated congeners are more easily transformed compared to higher chlorinated congeners and that PCB congeners with chlorines on one aromatic ring are more easily degraded than those bearing chlorine substituents on both aromatic rings. However, each isolate exhibits a particular activity spectrum with regard to the type and extent of PCB congeners metabolized, with some strains having a narrow spectrum and others, notably B. xenovorans LB400, being able to transform a broad range of congeners (Bopp 1986; Seeger et al. 1995a, b). The degradation of biphenyl and transformation of PCBs is usually catalyzed by enzymes encoded by the so-called biphenyl (bph) upper and lower pathways (Fig. 1).

2.1.1 Biphenyl 2,3-Dioxygenases Like the degradation of various other aromatics, the degradation of biphenyl is initiated by Rieske non-heme iron oxygenases, multicomponent enzyme complexes composed of a terminal oxygenase component (iron-sulfur protein [ISP]), and different electron transport proteins (a ferredoxin and a reductase or a combined ferredoxin-NADH-reductase) (Gibson and Parales 2000). Biphenyl 2,3-dioxygenases (BphA) usually belong to the toluene/biphenyl branch of Rieske non-heme iron oxygenases (Gibson and Parales 2000) where a ferredoxin (BphA3) and a ferredoxin reductase (BphA4) act as an electron transport system to transfer electrons from NADH to the terminal oxygenase, which consists of two subunits (BphA1A2), with the α-subunit being the major determinant of substrate specificity. The biphenyl 2,3-dioxygenases play a crucial role for the PCB degradation spectra. On one side, their regiospecificity of dioxygenation of the substrate determines the

5

Fig. 1 Pathway for biphenyl degradation

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

6

L. Agulló et al.

sites of attack by the subsequent enzymes of the pathway, while, on the other side, their substrates crucially determine the spectrum of PCB congeners that can be transformed by an organism. Studies on various biphenyl 2,3-dioxygenases have revealed considerable differences in their congener selectivity patterns, as well as their preference of the attacked ring (McKay et al. 1997; Seeger et al. 1999). The biphenyl pathway of strain LB400 oxidizes an unusually wide range of PCBs (from monochlorobiphenyls to 2,3,4,5,20 ,50 -hexachlorobiphenyl (Seeger et al. 1999)). Most primary catabolites that are dioxygenated by BphA of strain LB400 at ortho and meta carbons (Fig. 2) are further metabolized by the other enzymes of the upper pathway. In contrast, dioxygenation at meta and para positions results in channeling into a dead-end pathway. Dehalogenation by BphA of ortho-chlorinated (Fig. 2), ortho-brominated, and ortho-fluorinated biphenyls has been observed (Haddock et al. 1995; Seeger et al. 1995a), in addition to denitration and dehydroxylation (Seeger et al. 2001). Noteworthy, the dihydroxylation of natural and synthetic isoflavonoids and flavonoids by BphA of strain LB400 has also been described (Seeger et al. 2003; Overwin et al. 2015a).

2.1.2 cis-2,3-Dihydro-2,3-Dihydroxybiphenyl Dehydrogenases The second step in the metabolic pathway, the dehydrogenation of (chlorinated) cis2,3-dihydro-2,3-dihydroxybiphenyls (biphenyl 2,3-dihydrodiol) to give (chlorinated) 2,3-dihydroxybiphenyl, is catalyzed by cis-2,3-dihydro-2,3dihydroxybiphenyl dehydrogenases (BphB, Fig. 1). cis-Dihydrodiol dehydrogenases are involved in various aromatic degradation pathways. They are usually members of the family of short-chain alcohol dehydrogenases, generally of broad substrate specificity and able to transform several cis-dihydrodiol substrates (Rogers and Gibson 1977; Jouanneau and Meyer 2006). The cis-2,3-dihydro-2,3dihydroxybiphenyl dehydrogenase of strain LB400 is able to rearomatize isoflavonoids dihydroxylated by BphA, and the resulting products are assumed to have improved antioxidant properties (Arora et al. 1998). Interestingly, BphA from Burkholderia xenovorans LB400 catalyze dioxygenation of biphenyl 2,3-dihydrodiol (biphenyldienediol) in the nonoxidized ring to form biphenyl-bis-dienediol (Fig. 3) (Overwin et al. 2012). This metabolite is used as growth carbon source by B. xenovorans LB400 and Pseudomonas sp. strain B4-Magdeburg. BphB oxidizes both rings of the biphenyl-bis-dienediol in two successive steps into 2,3,20 ,30 -tetrahydroxybiphenyl (Overwin et al. 2012). In addition, BphA is able to oxidize 2,3-dihydroxybiphenyl produced in the biphenyl pathway to 2,3-dihydroxybiphenyl-4,6-diene-2,30 -diol, which is further transformed by BphB into 2,3,20 ,30 -tetrahydroxybiphenyl. 2.1.3 2,3-Dihydroxybiphenyl 1,2-Dioxygenases The ring cleavage of dihydroxylated aromatic intermediates can be catalyzed by enzymes from one of two structurally and mechanistically distinct enzyme classes. While intradiol dioxygenases, which cleave the aromatic nucleus between the hydroxyl substituents (ortho-cleavage), use non-heme Fe(III), extradiol dioxygenases, which cleave the aromatic nucleus adjacent to the hydroxyl

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

7

Fig. 2 Transformation of 4,40 -dichloro-, 2,20 -dichloro-, and 2,5,20 -trichlorobiphenyl by biphenyl 2,3-dioxygenase of B. xenovorans LB400. Unstable intermediates are shown in brackets. 4,40 Dichlorobiphenyl is exclusively subject to 2,3-dioxygenation yielding a 2,3-dihydrodiol as product. 2,20 -Dichlorobiphenyl is dioxygenated such that one of the vic-hydroxyl groups in the cis-dihydrodiol is bound to the same carbon as the chlorosubstituent. From such an unstable vic-dihydrodiol, the chlorosubstituent is spontaneously eliminated. 2,5,20 -Trichlorobiphenyl is subject to both 20 ,30 dioxygenation and 3,4-dioxygenation (Haddock et al. 1995; Seeger et al. 1995a, 1999, 2001)

8

L. Agulló et al.

Fig. 3 Conversion of biphenyl into 2,3,20 ,30 -tetrahydroxybiphenyl via two routes. Carbon numbering as used throughout this chapter indicated with compound 1. Compounds: 1, biphenyl; 2, biphenyldienediol [1-phenyl-cyclohexa-4,6-diene-cis-2,3-diol] (metabolite 1); 3a, 2,3-dihydroxybiphenyl; 3b, biphenyl-bisdienediol [1-(10 -cyclohexyl-40 ,60 -diene-20 ,30 -diol)cyclohexa-4,6-diene-2,3-diol] (metabolite 2); 4, 2,3-dihydroxybiphenyl-40 ,60 -diene-20 ,30 -diol [1-(10 -phenyl-20 ,30 -diol)-cyclohexa-4,6-diene-2,3-diol] (metabolite 3); 5, 2,3,20 ,30 -tetrahydroxybiphenyl (metabolite 4). The stereochemistry at carbons 20 and 30 of compounds 3b and 4 is proposed; it is based on the likely assumption that, for the second dioxygenation, the two rings simply exchange their positions within the BphA active site. Enzymes: BphA, biphenyl-2,3 dioxygenase (EC 1.14.12.18); BphB, cis-2,3-dihydrobiphenyl-2,3-diol dehydrogenase (EC 1.3.1.56) (Overwin et al. 2012)

substituents (meta-cleavage), typically use non-heme Fe(II) for cleavage (Harayama and Rekik 1989) even though Mn(II)-dependent extradiol dioxygenases have also been reported (Hatta et al. 2003). Among the extradiol dioxygenases, three types of enzymes could be identified. Type I extradiol dioxygenases belong to the vicinal oxygen chelate superfamily (Gerlt and Babbitt 2001), type II enzymes are exemplified by protocatechuate 4,5-dioxygenases and are often composed of two different subunits, and type III enzymes (such as gentisate 1,2-dioxygenase) belong to the cupin superfamily (Dunwell et al. 2001). Even though belonging to different families, all three types of extradiol dioxygenases share similar active sites, and all type I, type II, and various type III enzymes have the same iron ligands, two histidine and one glutamate, that constitute the 2-His 1-carboxylate structural motif. 2,3-Dihydroxybiphenyl 1,2-dioxygenases (BphC) involved in biphenyl degradation, usually belong to the subfamily 3A of type I extradiol dioxygenases (Eltis and Bolin 1996) and are specialized for transformation of 2,3-dihydroxybiphenyls (Fig. 1). Even though BphC enzymes differ in substrate specificity, they seem to be generally capable of transforming various chlorosubstituted derivatives (Dai et al. 2002; McKay et al. 2003). However, both 3,4-dihydroxybiphenyl and 20 -

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

9

chlorosubstituted 2,3-dihydroxybiphenyls strongly inhibit BphC enzymes (LloydJones et al. 1995; McKay et al. 2003). A special feature of extradiol dioxygenases is their susceptibility to inactivation due to a rapid oxidation of the active site ferrous iron into its ferric form with concomitant loss of activity (Vaillancourt et al. 2002). Specifically 20 -chlorosubstituted 2,3-dihydroxybiphenyls promote such inactivation and thus interfere with the degradation of other compounds (Dai et al. 2002). However, significant differences between different isoenzymes were observed (Fortin et al. 2005).

2.1.4

2-Hydroxy-6-Phenyl-6-Oxohexa-2,4-Dieneoate (HOPDA) Hydrolases The fourth step in the bph pathway is catalyzed by 2-hydroxy-6-phenyl-6-oxohexa2,4-dieneoate (HOPDA) hydrolase BphD, which hydrolyzes HOPDA to 2-hydroxypenta-2,4-dienoate and benzoate (Fig. 2). HOPDA hydrolases belong to the family of C–C hydrolase enzymes of the α/β-hydrolase enzyme superfamily (Ollis et al. 1992). Studies on B. xenovorans LB400 and Rhodococcus globerulus P6 BphDs have revealed that this enzyme may be a bottleneck for the metabolism of certain PCB congeners (Seeger et al. 1995b; Seah et al. 2000, 2001). Although some differences in turnover were observed, both enzymes were similar in that HOPDAs bearing chlorine substituents at the phenyl moiety were efficiently transformed, whereas HOPDAs bearing chlorine substituents on the dienoate moiety were poor substrates and competitively inhibit BphD (see Fig. 4). Studies suggest that this inhibition is due to inhibition of the histidine-mediated enol-keto tautomerization which precedes hydrolysis by BphD (Bhowmik et al. 2007). DxnB2 hydrolase from Sphingomonas wittichii RW1 catalyzes the hydrolysis of 3-Cl HOPDA more efficiently than BphD from B. xenovorans LB400 and Rhodococcus globerulus P6 (Seah et al. 2007; Ruzzini et al. 2013). Interestingly, DxnB2 is not inhibited by chlorinated HOPDAs, and the chlorine substituent is accommodated in the hydrophobic pocket (Ruzzini et al. 2013). 2.1.5 BphK Glutathione-S-Transferase BphK is a glutathione S-transferase (GST) that occurs in some bph pathways (Bartels et al. 1999). BphK was shown not to be essential for degradation of biphenyl (Bartels et al. 1999); however, this enzyme can catalyze dehalogenation of 4-chlorobenzoate (Fig. 4), the product of 4-chlorobiphenyl degradation by the enzymes BphA, BphB, BphC, and BphD (Gilmartin et al. 2003) suggesting that BphK was recruited to facilitate the degradation of PCBs. However, 3-chloro-2hydroxy-6-oxo-6-phenyl-2,4-dieneoates, compounds that are produced by the cometabolism of PCBs by BphA, BphB, and BphC (Fortin et al. 2006) and that inhibit BphD (Fig. 4), were significantly better substrates for the enzyme compared to 4-chlorobenzoate and were rapidly dehalogenated. Thus, BphK probably contributes to superior PCB-metabolizing activities by decreasing the inhibition of BphDs by chlorinated HOPDAs. A BphK modified at position 180 from an alanine to a

Fig. 4 Transformation of chlorosubstituted 2-hydroxy-6-oxo-6-phenyl-2,4-dieneoates (HOPDAs) by BphD and BphK gene products, exemplified by the metabolism of 4,40 -dichlorobiphenyl (Seah et al. 2000; Gilmartin et al. 2003; Fortin et al. 2006)

10 L. Agulló et al.

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

11

proline showed an increased GST activity toward diverse chlorinated pesticides (McGuinness et al. 2007).

2.1.6

Lower Pathways for the Degradation of 2-Hydroxypenta-2,4Dienoates and Benzoates The metabolism of (chloro)biphenyls by the biphenyl upper pathway results, in the best case, in the formation of (chlorinated) 2-hydroxypenta-2,4-dienoates and (chlorinated) benzoates (Fig. 1). 2-Hydroxypenta-2,4-dienoate is transformed by 2-hydroxypenta-2,4-dienoate hydratase (bphH), 4-hydroxy-2-oxovalerate aldolase (bphI), and an acetylating acetaldehyde dehydrogenase (bphJ) to pyruvate and acetyl-CoA (Fig. 1), which then can enter the Krebs cycle. The aldolase type II BphI forms a tetrameric complex with the dehydrogenase BphJ, channeling the toxic aldehyde from BphI to BphJ (Baker et al. 2011; Carere et al. 2011). Molecular determinants for volatile aldehyde products channeling by the BphI-BphJ complex have been reported (Carere et al. 2011). BphH, BphI, and BphJ enzymes allow growth of bacterial strains on biphenyls chlorinated at one aromatic ring only, which yield chlorinated benzoates as dead-end metabolites and unchlorinated 2-hydroxypenta-2,4-dienoate. If chlorinated 2-hydroxypenta-2,4-dienoate can be transformed by BphH has yet to be elucidated. Besides 2-hydroxypenta-2,4-dienoates, benzoates are generated during BphDcatalyzed hydrolysis of HOPDAs (Figs. 1 and 4). Benzoate is a growth substrate for a broad range of Actinobacteria and Proteobacteria and under aerobic conditions can be mineralized either via catechol and a 3-oxoadipate pathway or via 2,3-dihydroxydihydrobenzoyl-CoA and nonoxygenolytic cleavage of the aromatic ring. In contrast, chlorobenzoates, typically formed during metabolism of PCBs by the biphenyl upper pathway, are usually dead-end metabolites for PCB transforming bacteria.

2.2

Archetype bph Gene Clusters

Our knowledge on biphenyl degradation and PCB metabolism is significantly governed by analysis of some isolates which have been described in detail and are regarded as the archetype PCB degraders, among them, strains B. xenovorans LB400 and R. jostii RHA1, whose genomes have been deciphered. B. xenovorans LB400 (Mondello 1989), P. pseudoalcaligenes KF707 (Furukawa and Miyazaki 1986), and others harbor an operon comprising genes encoding enzymes of the biphenyl upper pathway, a glutathione S-transferase (bphK), and genes encoding enzymes involved in the transformation of 2-hydroxypenta-2,4-dienoate released during hydrolysis of HOPDA (Fig. 5). Regulation of these clusters is assumed to be mediated by an orf0 encoded GntR family transcriptional regulator (Watanabe et al. 2000). P. putida KF715 contains a bphABCD gene cluster (Hayase et al. 1990) (Fig. 5) which was suggested to have evolved from a LB400-type gene cluster. In LB400, the bph genes are located on a genomic island on the mega plasmid (Chain

12

L. Agulló et al.

et al. 2006). In Sphingobium fuliginis HC3, the bph genes are clustered in a plasmid (Hu et al. 2015). The presence of bph genes on mobile genetic elements indicate that these genes are able to move between genomes, thus allowing adaptation of microbial communities to PCBs. A second type of bph gene cluster was observed in Acidovorax sp. strain KKS102 (Kikuchi et al. 1994) and Cupriavidus oxalaticus A5 (Springael et al. 1993) (bphSEGF(orf4)A1A2A3A4BCD(orf1)A4) (Fig. 5). In these clusters, genes encoding enzymes involved in the transformation of 2-hydroxypenta-2,4-dienoate (designated bphEGF) are preceding genes encoding upper pathway enzymes, and the gene encoding the reductase subunit of biphenyl dioxygenase (bphA4) is localized at the end of the gene cluster. Like in LB400, regulation was shown to be dependent on a member of the GntR family of transcriptional regulators (BphS) (Mouz et al. 1999), and at least in C. oxalaticus A5, the bph genes are also located on a mobile genomic island (Toussaint et al. 2003). A third type bphAaAbAcAdCB gene cluster devoid of a gene encoding a HOPDA hydrolase was observed in R. jostii RHA1 (Masai et al. 1995) (Fig. 5) localized on the linear plasmid pRHL1 (Takeda et al. 2004). This catabolic gene cluster, like the similarly structured bph gene clusters of Rhodococcus sp. M5 (Peloquin and Greer 1993), is regulated by a two-component signal transduction system composed of a BphT response regulator and a BpdS sensor kinase, promoting transcriptional induction by a variety of aromatic compounds (Takeda et al. 2004). A nearly identical plasmid localized gene cluster has been shown to be involved in isopropylbenzene degradation by R. erythropolis BD2 (Stecker et al. 2003), indicating such gene clusters to be involved in the degradation of differently substituted aromatics. Figure 5 illustrates a fourth type of bph cluster (bphBCA1A2A3A4D) found in Rhodococcus sp. K37 and Rhodococcus sp. strain R04 (Yang et al. 2007; Taguchi et al. 2007), which differs from the bph cluster from Rhodococcus jostii RHA1.

2.3

Genome Analyses

The genomes of two potent PCB-degrading bacteria, B. xenovorans LB400 (Chain et al. 2006) and R. jostii RHA1 (McLeod et al. 2006), have been sequenced with the rational to better understand their overall physiology and to foster their applicability for bioremediation purposes. The LB400 genome has a size of 9.73 Mbp distributed over two circular chromosomes (4.87 Mbp and 3.36 Mbp, respectively) and a circular megaplasmid (1.47 Mbp). Strain RHA1 has a genome of 9.70 Mbp arranged on a linear chromosome (7.80 Mbp) and three linear plasmids (1.12 Mbp, 0.44 Mbp, and 0.33 Mbp, respectively). Both strains inhabit soil and plant rhizosphere niches. The large genomes of strains LB400 and RHA1 have evolved by different means. More than 20% of the genome of strain LB400 was recently acquired via horizontal gene transfer (HGT). In contrast, strain RHA1 evolved through ancient acquisition or gene duplication and acquired far fewer genes by recent HGT than LB400 (McLeod et al. 2006).

Fig. 5 Genetic organization of the bph gene clusters of B. xenovorans LB40, P. putida KF715, Rhodococcus sp. strain M5, Acidovorax sp. strain KKS102, Rhodococcus sp. K37, Bacillus sp. JF8, Sphingobium yanoikuyae B1, and of the bph and etb gene clusters of R. jostii RHA1

Genetics and Biochemistry of Biphenyl and PCB Biodegradation 13

14

L. Agulló et al.

Both bacterial strains have an unusually high metabolic versatility for degradation of aromatic compounds both with respect to peripheral routes activating aromatics for ring cleavage and for central routes channeling those intermediates into the Krebs cycle. Diverse active catabolic pathways for aromatic compounds from B. xenovorans LB400 have been reported (Méndez et al. 2011; Chirino et al., 2013; Romero-Silva et al. 2013; Agulló et al. 2017). The genes encoding enzymes of the biphenyl upper bph pathway are located in both strains on acquired and mobile genetic elements. In LB400 they are encoded by a genomic island on the megaplasmid, indicating that these genes were acquired via HGT (Chain et al. 2006). The genomic islands also provide strain LB400 with other highly specialized metabolic capabilities such as the abilities to degrade 2-aminophenol or 3-chlorocatechol. In strain RHA1, the bph genes are, like 11 of the 26 peripheral aromatic pathways, located on the plasmids (McLeod et al. 2006). The genomes of two additional PCB-degrading bacteria have been sequenced. Pseudomonas pseudoalcaligenes KF707 genome possesses an important number of genes involved in catabolic pathways of biphenyl/PCBs, phenol, benzoate, and other chloroaromatic compounds (Triscari-Barberi et al. 2012). Rhodococcus sp. WB1 genome contains a biphenyl/PCBs bphBA3A2A1CDA4 gene cluster and additional catabolic pathways genes for the degradation of nicotinate, nicotinamide, polycyclic aromatic hydrocarbons, toluene, nitrotoluene, and atrazine (Xu et al. 2016).

2.4

Toxicity of PCBs and Their Metabolites and Bacterial Stress Response

The toxicity of POPs and their catabolites for microorganisms is a major challenge for bioremediation processes (Blasco et al. 1995; Camara et al. 2004). PCBs are expected to accumulate in bacterial membranes due to their lipophilic character (Sikkema et al. 1995), and, in fact, PCBs decrease bacterial cell viability (Cámara et al. 2004). Noteworthy, some metabolic intermediates are even more toxic than PCBs. Degradation of specific PCB congeners by diverse bacteria is incomplete with a concomitant accumulation of different metabolic intermediates (Seeger et al. 1995a; Seah et al. 2000). Biotransformation of PCBs by BphA and BphB produces dihydrodiols and dihydroxybiphenyls, which are highly toxic for bacteria (Cámara et al. 2004). The increased polarity of dihydroxylated metabolites increases their aqueous solubility, contributing to this toxic effect. Hydroxylated PCB metabolites can affect the DNA content of bacteria, inhibiting bacterial cell separation (Hiraoka et al. 2002). The conversion of PCBs into products with increased toxicity is also known from the bioactivation of xenobiotics and drugs in higher organisms. In fact, the oxidation by cytochrome P450 generates reactive products that can be cytotoxic (Fig. 6). Moreover, chlorobenzoates, which are often dead-end products by PCB-metabolizing bacteria, can be transformed into deleterious downstream products. 3-Chlorocatechol can inactivate extradiol dioxygenases such as 2,3-dihydroxybiphenyl 1,2-dioxygenases (Vaillancourt et al. 2002), thus interfering with the biphenyl upper pathway. Channeling of 4-chlorocatechol into

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

15

the wide-spread 3-oxoadipate pathway can result in formation of the antibiotic protoanemonin (Blasco et al. 1995), and protoanemonin was assumed to be the reason for the poor survival of PCB cometabolizing organisms in soil microcosm studies (Blasco et al. 1997). Toxicity of PCBs as a direct result of the production of deleterious metabolites during cometabolism was also indicated in studies using the PCB degraders LB400 and RHA1 (Parnell et al. 2006). Although PCBs were shown to partition to the cell fraction, no significant effects were observed regarding viability or growth rate in either strain under non-PCB-degrading conditions, whereas significant straindependent differences were observed in cells metabolizing PCBs. Strain LB400 exhibited a high tolerance to PCB degradation-dependent toxicity, whereas RHA1 was highly sensitive. Evaluation of the genome- and proteome-wide defenses against PCB toxicity in LB400 showed induction of the molecular chaperones DnaK and GroEL during (chloro)biphenyl degradation (Agulló et al. 2007) and of DnaK and HtpG by 4-chlorobenzoate, a dead-end metabolite of the biphenyl upper pathway (Martínez et al. 2007), indicating that such exposure constitutes stressful conditions. The generation of reactive oxygen species (Chavez et al. 2004; Méndez 2017), probably resulting from the action of oxygenases in the metabolism, resulted in the induction of the alkyl hydroperoxide reductase AhpCF and other proteins indicating oxidative stress (Agulló et al. 2007; Méndez 2017). AhpCF detoxifies peroxides. The induction of a chloroacetaldehyde dehydrogenase (Denef et al. 2005) was suggested to reduce the concentration of toxic chlorinated aliphatic compounds resulting from PCB degradation. In order to establish optimized bioremediation processes for PCBs, it will be of paramount importance to overcome dead-end steps in the catabolic process and to balance the activities of enzymes involved in the degradation to avoid accumulation of toxic metabolites. Approaches to overcome a metabolic dead-end step (Saavedra et al. 2010) and to increase tolerance to oxidative stress (Ponce et al. 2011) of PCB-degrading bacteria have been successfully applied to improve biodegradation and bioremediation of PCBs and will be presented in Sect. 2.7. The isolation of novel PCB-degrading bacteria that also degrade chlorobenzoates such as Sphingobium fuliginis HC3 (Hu et al. 2015) will also provide improved catalysts for PCB bioremediation.

2.5 2.5.1

Metabolic Versatility

Diversity of Rieske Non-Heme Iron Oxygenases Involved in Biphenyl Metabolism More and more information becomes currently available that Rieske-type non-heme iron oxygenases outside of the archetype toluene/biphenyl branch are involved in biphenyl degradation. As an example, the bph operon of Bacillus sp. JF8 harbors a bphRDA1A2BC cluster (Mukerjee-Dhar et al. 2005) (Fig. 5) encoding enzymes only distantly related to enzymes of archetype Bph enzymes (Fig. 7), and BphA1 is more

16

L. Agulló et al.

Fig. 6 Biotransformation of PCBs into cytotoxic intermediates. Cytotoxic metabolites are boxed. Inhibition is indicated by a dashed arrow

closely related to naphthalene dioxygenases NidA from Rhodococcus sp. strain I24 (Larkin et al. 1999). Also, the Mn(II)-dependent BphC and BphD evidently belong to new subfamilies in the phylogeny of extradiol dioxygenases and hydrolases acting on extradiol cleavage products (Hatta et al. 2003; Mukerjee-Dhar et al. 2005). Analysis in Sphingobium yanoikuyae strain B1 revealed that a single ferredoxin and a single ferredoxin reductase, encoded by bphA3 and bphA4, respectively, can be shared by multiple oxygenase systems (Bae and Kim 2000), including biphenyl oxygenase encoded by the bphA1fA2f genes (Yu et al. 2007). In a phylogenetic analysis, BphA1f does not cluster with known BphAs, but is more related with PhnI from Sphingomonas sp. strain CHY-1, which was shown to be able to oxidize at least 8 PAHs made of 2-5 aromatic rings (Demaneche et al. 2004; Jakoncic et al. 2007) (Fig. 7). Accordingly, BphA1f is responsible for the capability of S. yanoikuyae B1 to dihydroxylate large aromatic compounds, such as chrysene and benzo[a]pyrene (Ferraro et al. 2007). Also the etbA1-encoded oxygenase α-subunit of R. jostii RHA1, only distantly related to previously characterized BphA1 proteins (see Fig. 7), has been implicated to be important for PCB metabolism as it is more active on highly chlorinated congeners than the bphAa-encoded one (Iwasaki et al. 2006) and obviously appropriate for both biphenyl and ethylbenzene transformation. Furthermore, another type of biphenyl oxygenase α-subunit has been discovered in Rhodococcus sp. strain K37 (Taguchi et al. 2007) (Fig. 7), evidencing that diversity of oxygenases involved in biphenyl degradation is highly underestimated. In general, it has to be considered that, despite the evolutionary adaptation of enzymes for specific substrates, the enzymes of a particular pathway often catalyze

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

17

the transformation of a range of substrate analogues, and specifically Rieske non-heme iron oxygenases are described by a broad substrate specificity. Among various other oxygenases, chlorobenzene dioxygenases (belonging, like biphenyl dioxygenases, to the toluene/biphenyl branch of Rieske non-heme iron oxygenases) (Raschke et al. 2001), naphthalene dioxygenases (belonging to the naphthalene family of Rieske non-heme iron oxygenases) (Fig. 7) (Barriault and Sylvestre 1999), phenanthrene dioxygenases (Kasai et al. 2003), or carbazole 1,9a dioxygenases (Nojiri et al. 1999) are capable to transform biphenyl. Additionally, culture-independent studies revealed the abundance of novel branches of Rieske-type non-heme iron oxygenases in contaminated sites, the importance and environmental function of which still remain to be elucidated (Taylor et al. 2002; Witzig et al. 2006), and studies on PCB-contaminated sites indicated novel undescribed types to be possibly important in situ (Leigh et al. 2007).

2.5.2 Mosaic Routes for Biphenyl Metabolism Metabolism of biphenyl and PCBs should not be regarded as a simple linear pathway, but often necessitates the complex interplay between different catabolic gene modules even inside single strains. As an example, the bph cluster of R. jostii RHA1 does not comprise a bphD gene, and such activity has to be recruited from elsewhere in the genome. In fact, three hydrolases were shown to be upregulated during growth of RHA1 on biphenyl (Goncalves et al. 2006) with one of them, termed BphD previously, shown to be capable to attack HOPDA (Yamada et al. 1998). P. putida strain CE2010 mineralizes biphenyl by a mosaic of tod (toluene) and cmt (cumate) pathways (Ohta et al. 2001). As previously reported, toluene dioxygenase (TodC1C2BA), toluene dihydrodiol dehydrogenase (TodD), and the meta-cleavage enzyme TodE have a significant cross-reactivity with biphenyl or metabolites produced during biphenyl degradation (Furukawa et al. 1993), whereas TodF 2-hydroxy-6-oxohepta-2,4-dienoate hydrolase cannot cope with HOPDA. Recruitment of a hydrolase active with HOPDA, such as in RHA1, allows CE21010 to mineralize biphenyl. The same holds for extradiol dioxygenases, especially in Rhodococcus, where the presence of multiple extradiol dioxygenase encoding genes has been reported (Taguchi et al. 2004; McLeod et al. 2006). The metabolic versatility of catabolic enzymes and pathways is an indication of the ongoing evolution of bacterial metabolism, thus endowing environmental microbes with the capabilities to deal with a broad range of pollutants.

2.6

Optimized Enzymes and PCB-Degrading Organisms

Pollution by PCBs typically consists of mixtures of congeners, and only a fraction of these can be attacked by known BphAs. Therefore, for improved PCB catabolic pathways, recruitment or generation of improved biphenyl 2,3-dioxygenases is required. The construction of chimeric BphA derivatives generated by the combination of gene segments of well-known PCB degraders enabled the identification of

18

L. Agulló et al.

Fig. 7 Dendrogram showing the relatedness of oxygenase α-subunits of Rieske non-heme iron oxygenases. α-Subunits of supposed or validated biphenyl 2,3-dioxygenases are indicated by a filled circle

key domains of these oxygenases (Kimura et al. 1997; Kumamaru et al. 1998) and generated biphenyl 2,3-dioxygenases with improved capacities (Erickson and Mondello 1993; Mondello et al. 1997; Suenaga et al. 1999, 2002). A directed evolution approach using random mutagenesis to specific segments allowed generating BphAs with increased turnover of PCBs, largely recalcitrant to attack by the parental enzyme (Zielinski et al. 2006). On the other side, the isolation of naturally occurring enzymatic activities by metagenomic methods which circumvent the cultivation of organisms has been used (Cámara et al. 2007). Combination of both approaches, the broad natural diversity and methods of artificial evolution by family shuffling of soil DNA encoding BphA segments, generates BphA variants with novel regioselectivities (Vezina et al. 2007). Even though enzyme optimizations have been mainly applied to biphenyl 2,3-dioxygenases, efforts have been also directed toward elucidation of pathway bottlenecks in downstream enzyme activities and in identifying optimized isoenzymes. As an example, a HOPDA hydrolase with novel specificities toward polychlorinated biphenyl metabolites, which specifically transformed 3-chlorosubstituted HOPDAs, compounds that inhibit archetype BphDs, was

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

19

recently characterized from S. wittichii RW1 (Seah et al. 2007). On the other side, a modified BphK with increased GST activity toward a broad range of chlorinated organic substrates (McGuinness et al. 2007) may be useful to improve PCB mineralization. As described above, most currently available microorganisms are capable to mineralize biphenyl, but only cometabolize PCBs due to the absence of enzymes necessary for mineralization of chlorobenzoates, generated through metabolism by the biphenyl upper pathway. The strategy of combining complementary metabolic activities for the development of microorganisms capable of mineralizing PCBs by combining an oxidative pathway for (chloro)biphenyl transformation (encoded by the bph genes) into (chloro)benzoate with a chlorobenzoate degradative pathway had been followed for various years. Several hybrid strains have been engineered by conjugative matings (Reineke 1998) of appropriate organisms or by introduction of the bph genes into chlorobenzoate degraders, usually using a degradative pathway for chlorobenzoates via the corresponding chlorocatechols. By cloning and expressing the genes encoding enzymes for ortho- and para-dechlorination of chlorobenzoates in biphenyl-degrading and chlorinated biphenyls co-metabolizing strains, derivatives capable of growing on and completely dechlorinating 2- and 4-chlorobiphenyl could also be obtained (Hrywna et al. 1999). However, it should be noted that novel isolates with interesting metabolic properties capable to mineralize some PCBs are still being isolated (Adebusoye et al. 2008). As mentioned above, the isolation of S. fuliginis strain HC3 that efficiently degrades PCBs without accumulation of dead-end intermediates was reported (Hu et al. 2015). Notably, the BphA-B4h hybrid enzyme based on biphenyl-dioxygenase from Pseudomonas sp. B4-Magdeburg and B. xenovorans LB400 is more efficient than BphA from strain LB400 for the dioxygenation of aromatic compounds (Overwin et al. 2015b). The BphA-B4h hybrid enzyme possesses a remarkable capacity for the double dioxygenation of various bicyclic aromatic compounds generating bis-DHDs (Fig. 3) (Overwin et al. 2016). The BphA-B4h hybrid dioxygenase is able to hydroxylate diverse flavonoids into novel products with interesting biological activities (Overwin et al. 2015a). The biotransformation of flavone, isoflavone, flavanone, and isoflavanol by the biphenyl dioxygenase from P. pseudoalcaligenes KF707 was reported (Seo et al. 2011).

2.7

Bioremediation of PCBs

Bioremediation of PCB-contaminated soils and sediments in microcosmos using aerobic and anaerobic bacteria has been reported (Singer et al. 2000; Bedard et al. 2007; Saavedra et al. 2010; Ponce et al. 2011; Sowers and May 2013; Payne et al. 2013). The genetically modified strain Cupriavidus pinatubonensis JMS34 containing the bph locus from strain LB400 is able to mineralize low chlorinated biphenyls in polluted soils (Saavedra et al. 2010). The addition of antioxidant compounds improved PCB degradation by B. xenovorans strain LB400 in soils (Ponce et al. 2011). Bioaugmentation of sediment microcosms and mesocosms indicates a high PCB degradation by aerobic B. xenovorans LB400 and anaerobic

20

L. Agulló et al.

Dehalobium chlorocoercia DF1 (Payne et al. 2013). An increase of β-Proteobacteria and Actinobacteria in soils polluted with PCBs has been reported, which correlates with an increase of two bphA1 gene subgroups from β-Proteobacteria (Comamonas, Burkholderia) and Actinobacteria (Rhodococcus), indicating the selection of PCB-degrading bacteria (Correa et al. 2010). Phytoremediation and natural attenuation of PCBs has also been reported (Gomes et al. 2013). A transgenic Nicotiana tabacum plant that expresses the bphC gene from Pandoraea pnomenusa B-356 was applied for the removal of 2,3-dihydroxybiphenyl (Novakova et al. 2010). Bioaugmentation with B. xenovorans LB400 combined with phytoremediation using switchgrass increased the degradation of PCBs in soils (Liang et al. 2014). However, in situ PCB bioremediation approaches have to be further studied and developed.

3

Research Needs

The isolation of novel PCB-degrading bacteria with improved capabilities for the mineralization of PCBs is still a challenge in order to optimize bioremediation of PCBs. Genome studies of PCB degraders provide novel insights into the catabolic capabilities and the adaptation of bacteria to these pollutants. The sequencing of genomes of additional PCB-degrading strains will provide the basis for the understanding of the physiology and adaptation of bacteria dealing with PCBs and their toxic metabolic intermediates and related processes such as biofilm formation. Biofilm formation by bacteria is crucial toward their application for bioremediation. Different PCB-bioremediation processes have been reported in the last decades. Nevertheless, the field is still under development. A higher number of bioremediation trials in soils, sediments, and aquatic systems at diverse scale-up levels including microcosms, macrocosms, and field studies are still required to find out crucial aspects and variables to improve these complex processes. Bioaugmentation, biostimulation, and bioventing are three of the most relevant technologies for bioremediation that should be further studied. For bioaugmentation, bacterial consortia or a combination of bacteria and fungi are main catalysts that should be applied in novel cleanup processes. Dynamics of microbial communities during bioremediation of PCBs in anaerobic and aerobic conditions should be addressed for the knowledge of main bioremediation actors and the most favorable scenarios for the cleanup of PCB-polluted environments. The knowledge of PCB degradation will require integration of single-microorganism analyses with environmental studies to understand functioning of complex microbial communities during the cleanup processes and to design novel bioremediation strategies. Acknowledgments M.S. gratefully acknowledges support from the grants FONDECYT (1070507, 1020221, 1110992, 1151174, 7020221, 7070174, 7080148, 7090079, and 7100027), USM (130522, 130836, 130948, 131109, 131342, 131562), MILENIO P04/007-F (MIDEPLAN), and CONICYT-BMBF. D.P. gratefully acknowledges support from the grant EU GOCE 003998 (BIOTOOL) and BACSIN.

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

21

References Adebusoye SA, Picardal FW, Ilori MO, Amund OO (2008) Evidence of aerobic utilization of diortho-substituted trichlorobiphenyls as growth substrates by Pseudomonas sp. SA-6 and Ralstonia sp. SA-4. Environ Microbiol 10:1165–1174 Adrian L, Rahnenführer J, Gobom J, Hölscher T (2007) Identification of a chlorobenzene reductive dehalogenase in Dehalococcoides sp. strain CBDB1. Appl Environ Microbiol 73 (23):7717–7724 Adrian L, Dudková V, Demnerová K, Bedard DL (2009) Dehalococcoides sp. strain CBDB1 extensively dechlorinates the commercial polychlorinated biphenyl mixture Aroclor 1260. Appl Environ Microbiol 75: 4516–4524 Agulló L, Cámara B, Martínez P, Latorre V, Seeger M (2007) Response to (chloro)biphenyls of the polychlorobiphenyl-degrader Burkholderia xenovorans LB400 involves stress proteins also induced by heat shock and oxidative stress. FEMS Microbiol Lett 267:167–175 Agulló L, Romero-Silva MJ, Domenech M, Seeger M (2017) p-Cymene promotes its catabolism through the p-cymene and the p-cumate pathways, activates a stress response and reduces the biofilm formation in Burkholderia xenovorans LB400. PLoS One 12(1):e0169544. doi:10.1371/journal.pone.0169544 Arora A, Nair MG, Strasburg GM (1998) Antioxidant activities of isoflavones and their biological metabolites in a liposomal system. Arch Biochem Biophys 356:133–141 Bae M, Kim E (2000) Association of a common reductase with multiple aromatic terminal dioxygenases in Sphingomonas yanoikuyae strain B1. J Microbiol 38:40–43 Baker P, Carere J, Seah SY (2011) Probing the molecular basis of substrate specificity, stereospecificity, and catalysis in the class II pyruvate aldolase, BphI. Biochemistry 50:3559–3569 Barriault D, Sylvestre M (1999) Catalytic activity of Pseudomonas putida strain G7 naphthalene 1,2-dioxygenase on biphenyl. Int Biodeterior Biodegrad 44:33–37 Bartels F, Backhaus S, Moore ERB, Timmis KN, Hofer B (1999) Occurrence and expression of glutathione-S-transferase-encoding bphK genes in Burkholderia sp. strain LB400 and other biphenyl-utilizing bacteria. Microbiology 145:2821–2834 Bedard DL, Ritalahti KM, Loffler FE (2007) The Dehalococcoides population in sediment-free mixed cultures metabolically dechlorinates the commercial polychlorinated biphenyl mixture Aroclor 1260. Appl Environ Microbiol 73:2513–2521 Bhowmik S, Horsman GP, Bolin JT, Eltis LD (2007) The molecular basis for inhibition of BphD, a C–C bond hydrolase involved in polychlorinated biphenyls degradation: large 3-substituents prevent tautomerization. J Biol Chem 282:36377–36385 Blasco R, Wittich R-M, Mallavarapu M, Timmis KN, Pieper DH (1995) From xenobiotic to antibiotic. Formation of protoanemonin from 4-chlorocatechol by enzymes of the 3-oxoadipate pathway. J Biol Chem 270:29229–29235 Blasco R, Mallavarapu M, Wittich RM, Timmis KN, Pieper DH (1997) Evidence that formation of protoanemonin from metabolites of 4-chlorobiphenyl degradation negatively affects the survival of 4-chlorobiphenyl-cometabolizing microorganisms. Appl Environ Microbiol 63:427–434 Bopp L (1986) Degradation of highly chlorinated PCBs by Pseudomonas strain LB400. J Ind Microbiol 1:23–29 Cámara B, Herrera C, González M, Couve E, Hofer B, Seeger M (2004) From PCBs to highly toxic metabolites by the biphenyl pathway. Environ Microbiol 6:842–850 Cámara B, Seeger M, González M, Standfuss-Gabisch C, Kahl S, Hofer B (2007) Generation of a hybrid dioxygenase showing improved oxidation of polychlorobiphenyls by a widely applicable approach. Appl Environ Microbiol 73:2682–2689 Carere J, Baker P, Seah S (2011) Investigating the molecular determinants for substrate channeling in BphI – BphJ, an aldolase – dehydrogenase complex from the polychlorinated biphenyls degradation pathway. Biochemistry 50:8407–8416 Chain PS, Denef VJ, Konstantinidis KT, Vergez LM, Agulló L, Reyes VL, Hauser L, Córdova M, Gomez L, González M, Land M, Lao V, Larimer F, LiPuma JJ, Mahenthiralingam E, Malfatti

22

L. Agulló et al.

SA, Marx CJ, Parnell JJ, Ramette A, Richardson P, Seeger M, Smith D, Spilker T, Sul WJ, Tsoi TV, Ulrich LE, Zhulin IB, Tiedje JM (2006) Burkholderia xenovorans LB400 harbors a multireplicon, 9.73-Mbp genome shaped for versatility. Proc Natl Acad Sci U S A 103:15280–15287 Chavez FP, Lünsdorf H, Jerez CA (2004) Growth of polychlorinated-biphenyl-degrading bacteria in the presence of biphenyl and chlorobiphenyls generates oxidative stress and massive accumulation of inorganic polyphosphate. Appl Environ Microbiol 70:3064–3072 Chirino B, Strahsburger E, Agulló L, González M, Seeger M (2013) Genomic and functional analyses of the 2-aminophenol catabolic pathway and partial conversion of its substrate into picolinic acid in Burkholderia xenovorans LB400. PLoS ONE 8:e75746 Correa PA, Lin L, Just CL, Hu D, Hornbuckle KC, Schnoor JL, Van Aken B (2010) The effects of individual PCB congeners on the soil bacterial community structure and the abundance of biphenyl dioxygenase genes. Environ Int 36(8):901–906 Cutter LA, Watts JEM, Sowers KR, May HD (2001) Identification of a microorganism that links its growth to the reductive dechlorination of 2,3,5,6-chlorobiphenyl. Environ Microbiol 3:699–709 Dai S, Vaillancourt F, Maaroufi H, Drouin N, Neau D, Snieckus V, Bolin J, Eltis L (2002) Identification and analysis of a bottleneck in PCB biodegradation. Nat Struct Biol 9:934–939 Demaneche S, Meyer C, Micoud J, Louwagie M, Willison JC, Jouanneau Y (2004) Identification and functional analysis of two aromatic-ring-hydroxylating dioxygenases from a Sphingomonas strain that degrades various polycyclic aromatic hydrocarbons. Appl Environ Microbiol 70:6714–6725 Denef VJ, Patrauchan MA, Florizone C, Park J, Tsoi TV, Verstraete W, Tiedje JM, Eltis LD (2005) Growth substrate- and phase-specific expression of biphenyl, benzoate, and C1 metabolic pathways in Burkholderia xenovorans LB400. J Bacteriol 187:7996–8005 Drinker CK, Warren MF, Bennet GA (1937) The problem of possible systemic effects from certain chlorinated hydrocarbons. J Ind Hyg Toxicol 19:283–311 Dunwell JM, Culham A, Carter C, Sos-Aguirre C, Goodenough PW (2001) Evolution of functional diversity in the cupin superfamily. Trends Biochem Sci 26:740–746 Eltis LD, Bolin JT (1996) Evolutionary relationships among extradiol dioxygenases. J Bacteriol 178:5930–5937 Erickson BD, Mondello FJ (1993) Enhanced biodegradation of polychlorinated biphenyls after sitedirected mutagenesis of a biphenyl dioxygenase gene. Appl Environ Microbiol 59:3858–3862 Faroon O, Jones D, de Rosa C (2001) Effects of polychlorinated biphenyls on the nervous system. Toxicol Ind Health 16:305–333 Fennell DE, Nijenhuis I, Wilson SF, Zinder SH, Haggblom MM (2004) Dehalococcoides ethenogenes strain 195 reductively dechlorinates diverse chlorinated aromatic pollutants. Environ Sci Technol 38:2075–2081 Ferraro D, Daniel J, Brown EN, Yu CL, Parales RE, Gibson DT, Ramaswamy S (2007) Structural investigations of the ferredoxin and terminal oxygenase components of the biphenyl 2,3-dioxygenase from Sphingobium yanoikuyae B1. BMC Struct Biol 9:10 Fortin PD, Horsman G, Yang H, Eltis LD (2006) A glutathione S-transferase catalyzes the dehalogenation of inhibitory metabolites of polychlorinated biphenyls. J Bacteriol 188:4424–4430 Fortin PD, Lo ATF, Haro MA, Kaschabek SR, Reineke W, Eltis LD (2005) Evolutionarily divergent extradiol dioxygenases possess higher specificities for polychlorinated biphenyl metabolites. J Bacteriol 187:415–421 Furukawa K, Miyazaki T (1986) Cloning of a gene cluster encoding biphenyl and chlorobiphenyl degradation in Pseudomonas pseudoalcaligenes. J Bacteriol 166:392–398 Furukawa K, Hirose J, Suyama A, Zaiki T, Hayashida S (1993) Gene components responsible for discrete substrate specificity in the metabolism of biphenyl (bph operon) and toluene (tod operon). J Bacteriol 175:5224–5232 Fuentes S, Méndez V, Aguila P, Seeger M (2014) Bioremediation of petroleum hydrocarbons: catabolic genes, microbial communities, and applications. Appl Microbiol Biotechnol 98 (11):4781–4794

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

23

Fuentes S, Ding GC, Cárdenas F, Smalla K, Seeger M (2015) Assessing environmental drivers of microbial communities in estuarine soils of the Aconcagua River in Central Chile. FEMS Microbiol Ecol 91:fiv110. doi:10.1093/femsec/fiv110 Fuentes S, Barra B, Caporaso JG, Seeger M (2016) From rare to dominant: a fine-tuned soil bacterial bloom during petroleum hydrocarbon bioremediation. Appl Environ Microbiol 82 (3):888–896 Furukawa K (2000) Biochemical and genetic bases of microbial degradation of polychlorinated biphenyls (PCBs). J Gen Appl Microbiol 46:283–296 Gerlt JA, Babbitt PC (2001) Divergent evolution of enzymatic function: mechanistically diverse superfamilies and functionally distinct suprafamilies. Annu Rev Biochem 70:209–246 Gibson DT, Parales RE (2000) Aromatic hydrocarbon dioxygenases in environmental biotechnology. Curr Opin Biotechnol 11:236–243 Gilmartin N, Ryan D, Sherlock O, Dowling DN (2003) BphK shows dechlorination activity against 4-chlorobenzoate, an end-product of bph-promoted degradation of PCBs. FEMS Microbiol Lett 222:251–255 Gomes H, Dias-Ferreira C, Ribeiro A (2013) Overview of in situ and ex situ remediation technologies for PCB-contaminated soils and sediments and obstacles for full-scale application. Sci Total Environ 445-–446:237–260 Gomez-Gutiérrez A, Garnacho E, Bayona JM, Albaigés J (2007) Assessment of the Mediterranean sediments contamination by persistent organic pollutants. Environ Pollut 148:396–408 Goncalves E, Hara H, Miyazawa D, Davies J, Eltis LD, Mohn WW (2006) Transcriptomic assessment of isoenzymes in the biphenyl pathway of Rhodococcus sp. strain RHA1. Appl Environ Microbiol 72:6283–6193 Haddock JD, Horton JR, Gibson DT (1995) Dihydroxylation and dechlorination of chlorinated biphenyls by purified biphenyl 2,3-dioxygenase from Pseudomonas sp. strain LB400. J Bacteriol 177:20–26 Harayama S, Rekik M (1989) Bacterial aromatic ring-cleavage enzymes are classified into two different gene families. J Biol Chem 264:15328–15333 Hatta T, Mukerjee-Dhar G, Damborsky J, Kiyohara H, Kimbara K (2003) Characterization of a novel thermostable Mn(II)-dependent 2,3-dihydroxybiphenyl 1,2-dioxygenase from a polychlorinated biphenyl- and naphthalene-degrading Bacillus sp JF8. J Biol Chem 278:21483–21492 Hayase N, Taira K, Furukawa K (1990) Pseudomonas putida KF715 bphABCD operon encoding biphenyl and polychlorinated biphenyl degradation: cloning analysis, and expression in soil bacteria. J Bacteriol 172:1160–1164 Hiraoka Y, Yamada T, Tone K, Futaesaku Y, Kimbara K (2002) Flow cytometry analysis of changes in the DNA content of the polychlorinated biphenyl degrader Comamonas testosteroni TK102: effect of metabolites on cell-cell separation. Appl Environ Microbiol 68:5104–5112 Hrywna Y, Tsoi TV, Maltseva OV, Quensen JF, Tiedje JM (1999) Construction and characterization of two recombinant bacteria that grow on ortho- and para-substituted chlorobiphenyls. Appl Environ Microbiol 65:2163–2169 Hu J, Qian M, Zhang Q, Cui J, Yu C, Su X et al (2015) Sphingobium fuliginis HC3: a novel and robust isolated biphenyl- and polychlorinated biphenyls-degrading bacterium without dead-end intermediates accumulation. PLoS ONE 10(4):e0122740. doi:10.1371/journal.pone.0122740 Iwasaki T, Miyauchi K, Masai E, Fukuda M (2006) Multiple-subunit genes of the aromatic-ringhydroxylating dioxygenase play an active role in biphenyl and polychlorinated biphenyl degradation in Rhodococcus sp. strain RHA1. Appl Environ Microbiol 72:5396–5402 Jakoncic J, Jouanneau Y, Meyer C, Stojanoff V (2007) The catalytic pocket of the ringhydroxylating dioxygenase from Sphingomonas CHY-1. Biochem Biophys Res Commun 352:861–866 Jouanneau Y, Meyer C (2006) Purification and characterization of an arene cis-dihydrodiol dehydrogenase endowed with broad substrate specificity toward polycyclic aromatic hydrocarbon dihydrodiols. Appl Environ Microbiol 72:4726–4734

24

L. Agulló et al.

Kasai Y, Shindo K, Harayama S, Misawa N (2003) Molecular characterization and substrate preference of a polycyclic aromatic hydrocarbon dioxygenase from Cycloclasticus sp. strain A5. Appl Environ Microbiol 69:6688–6697 Kikuchi Y, Yasukochi Y, Nagata Y, Fukuda M, Takagi M (1994) Nucleotide sequence and functional analysis of the meta-cleavage pathway involved in biphenyl and polychlorinated biphenyl degradation in Pseudomonas sp. strain KKS102. J Bacteriol 176:4269–4276 Kimura N, Nishi A, Goto M, Furukawa K (1997) Functional analyses of a variety of chimeric dioxygenases constructed from two biphenyl dioxygenases that are similar structurally but different functionally. J Bacteriol 179:3936–3943 Kumamaru T, Suenaga H, Mitsuoka M, Watanabe T, Furukawa K (1998) Enhanced degradation of polychlorinated biphenyls by directed evolution of biphenyl dioxygenase. Nat Biotechnol 16:663–666 Lauby-Secretan B, Loomis D, Grosse Y, El Ghissassi F, Bouvard V, Benbrahim-Tallaa L, Guha N, Baan R, Mattock H, Straif K et al (2013) Carcinogenicity of polychlorinated biphenyls and polybrominated biphenyls. Lancet Oncol 14:287–288 Larkin MJ, Allen CCR, Kulakov LA, Lipscomb DA (1999) Purification and characterization of a novel naphthalene dioxygenase from Rhodococcus sp strain NCIMB12038. J Bacteriol 181:6200–6204 Leigh MB, Pellizari VH, Uhlík O, Sutka R, Rodrigues J, Ostrom NE, Zhou J, Tiedje JM (2007) Biphenyl-utilizing bacteria and their functional genes in a pine root zone contaminated with polychlorinated biphenyls (PCBs). ISME J 1:134–148 Liang Y, Meggo R, Hu D, Schnoor JL, Mattes TE (2014) Enhanced polychlorinated biphenyl removal in a switchgrass rhizosphere by bioaugmentation with Burkholderia xenovorans LB400. Ecol Eng 71:215–222 Lloyd-Jones G, Ogden RC, Williams PA (1995) Inactivation of 2,3-dihydroxybiphenyl 1,2-dioxygenase from Pseudomonas sp. strain CB406 by 3,4-dihydroxybiphenyl (4-phenylcatechol). Biodegradation 6:11–17 Lunt D, Evans WC (1970) The microbial metabolism of biphenyl. Biochem J 118:54–55 Martínez P, Agulló L, Hernández M, Seeger M (2007) Chlorobenzoate inhibits growth and induces stress proteins in the PCB-degrading bacterium Burkholderia xenovorans LB400. Arch Microbiol 188:289–297 Masai E, Yamada A, Healy JM, Hatta T, Kimbara K, Fukuda M, Yano K (1995) Characterization of biphenyls catabolic genes of Gram-positive polychlorinated biphenyls degrader Rhodococcus sp. strain RHA1. Appl Environ Microbiol 61:2079–2085 Mayes BA, McConnell EE, Neal BH, Brunner MJ, Hamilton SB, Sullivan TM, Peters AC, Ryan MJ, Toft JD, Singer AW, Brown JF, Menton RG, Moore JA (1998) Comparative carcinogenicity in Sprague-Dawley rats of the polychlorinated biphenyl mixtures aroclors 1016, 1242, 1254, and 1260. Toxicol Sci 41:62–76 McKay DB, Seeger M, Zielinski M, Hofer B, Timmis KN (1997) Heterologous expression of biphenyl dioxygenase-encoding genes from a Gram-positive broad-spectrum polychlorinated biphenyl degrader and characterization of chlorobiphenyl oxidation by the gene products. J Bacteriol 179:1924–1930 McGuinness MC, Mazurkiewicz V, Brennan E, Dowling DN (2007) Dechlorination of pesticides by a specific bacterial glutathione S-transferase, BphK LB400: potential for bioremediation. Eng Life Sci 7(6):611–615 McKay DB, Prucha M, Reineke W, Timmis KN, Pieper DH (2003) Substrate specificity and expression of three 2,3-dihydroxybiphenyl 1,2-dioxygenases from Rhodococcus globerulus strain P6. J Bacteriol 185:2944–2951 McLeod MP, Warren RL, Hsiao WW, Araki N, Myhre M, Fernandes C, Miyazawa D, Wong W, Lillquist AL, Wang D, Dosanjh M, Hara H, Petrescu A, Morin RD, Yang G, Stott JM, Schein JE, Shin H, Smailus D, Siddiqui AS, Marra MA, Jones SJ, Holt R, Brinkman FS, Miyauchi K, Fukuda M, Davies JE, Mohn WW, Eltis LD (2006) The complete genome of Rhodococcus

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

25

sp. RHA1 provides insights into a catabolic powerhouse. Proc Natl Acad Sci U S A 103:15582–15587 Méndez V, Agulló L, González M, Seeger M (2011) The homogentisate and homoprotocatechuate central pathways are involved in 3- and 4-hydroxyphenylacetate degradation by Burkholderia xenovorans LB400. PLoS ONE 6:e17583 Méndez V (2017) Molecular mechanisms of the adaptive response of Burkholderia xenovorans LB400 to oxidative stress induced by oxidants exposure and aromatic metabolism. Biotechnology PhD thesis, Universidad Técnica Federico Santa María & Pontificia Universidad Catolica de Valparaiso, Valparaiso, Chile Mondello FJ (1989) Cloning and expression in Escherichia coli of Pseudomonas strain LB400 genes encoding polychlorinated biphenyl degradation. J Bacteriol 171:1725–1732 Mondello FJ, Turcich MP, Lobos JH, Erickson BD (1997) Identification and modification of biphenyl dioxygenase sequences that determine the specificity of polychlorinated biphenyl degradation. Appl Environ Microbiol 63:3096–3103 Mouz S, Merlin C, Springael D, Toussaint A (1999) A GntR-like negative regulator of the biphenyl degradation genes of the transposon Tn4371. Mol Gen Genet 262:790–799 Mukerjee-Dhar G, Shimura M, Miyazawa D, Kimbara K, Hatta T (2005) bph genes of the thermophilic PCB degrader, Bacillus sp. JF8: characterization of the divergent ringhydroxylating dioxygenase and hydrolase genes upstream of the Mn-dependent BphC. Microbiology 151:4139–4151 Müller JA, Rosner BM, von Abendroth G, Meshulam-Simon G, McCarty PL, Spormann AM (2004) Molecular identification of the catabolic vinyl chloride reductase from Dehalococcoides sp. strain VS and its environmental distribution. Appl Environ Microbiol 70:4880–4888 Neumann A, Wohlfarth G, Diekert G (1996) Purification and characterization of tetrachloroethene reductive dehalogenase from Dehalospirillum multivorans. J Biol Chem 271:16515–16519 Nojiri H, Nam JW, Kosaka M, Morii KI, Takemura T, Furihata K, Yamane H, Omori T (1999) Diverse oxygenations catalyzed by carbazole 1,9a-dioxygenase from Pseudomonas sp strain CA10. J Bacteriol 181:3105–3113 Novakova M, Mackova M, Antosova Z, Viktorova J, Szekeres M, Demnerova K, Macek T (2010) Cloning the bacterial bphC gene into Nicotiana tabacum to improve the efficiency of phytoremediation of polychlorinated biphenyls. Bioeng Bugs 1(6):419–423 Ohta Y, Maeda M, Kudo T (2001) Pseudomonas putida CE2010 can degrade biphenyl by a mosaic pathway encoded by the tod operon and cmtE, which are identical to those of P. putida F1 except for a single base difference in the operator-promoter region of the cmt operon. Microbiology 147:31–41 Ollis DL, Cheah E, Cygler M, Dijkstra B, Frolow F, Franken SM, Harel M, Remington SJ, Silman I, Schrag J, Sussman JL, Verschueren KHG, Goldman A (1992) The alpha/beta hydrolase fold. Protein Eng 5:197–211 Overwin H, González M, Méndez V, Seeger M, Wray V, Hofer B (2012) Dioxygenation of the biphenyl dioxygenation product. Appl Environ Microbiol 78:4529–4532 Overwin H, González M, Méndez V, Cárdenas F, Seeger M, Hofer B (2015a) Stepwise conversion of flavonoids by engineered dioxygenases and dehydrogenase: characterization of novel biotransformation products. Enzym Microb Technol 81:63–71 Overwin H, Standfuß-Gabisch C, González M, Méndez V, Seeger M, Reichelt J, Wray V, Hofer B (2015b) Permissivity of the biphenyl specific aerobic bacterial metabolic pathway towards analogues with various steric requirements. Microbiology 161:1844–1856 Overwin H, González M, Méndez V, Seeger M, Wray V, Hofer B (2016) An aryl dioxygenase shows remarkable double dioxygenation capacity for diverse bis-aryl compounds, provided they are carbocyclic. Appl Microbiol Biotechnol 100:8053–8061. doi:10.1007/s00253-016-7570-0 Payne RB, Fagervold SK, May HD, Sowers KR (2013) Remediation of polychlorinated biphenyl impacted sediment by concurrent bioaugmentation with anaerobic halorespiring and aerobic degrading bacteria. Environ Sci Technol 47(8):3807–3815

26

L. Agulló et al.

Palma-Fleming H, Cornejo C, González M, Pérez V, González M, Gutierrez E, Sericano JL, Seeger M (2008) Polycyclic aromatic hydrocarbons and polychlorinated biphenyls from the coastal reef of Valdivia and Valparaíso region Chile. J Chil Chem Soc 53:1393–1398 Parnell JJ, Park J, Denef V, Tsoi T, Hashsham S, Quensen J III, Tiedje JM (2006) Coping with polychlorinated biphenyl (PCB) toxicity: physiological and genome-wide responses of Burkholderia xenovorans LB400 to PCB-mediated stress. Appl Environ Microbiol 72:6607–6614 Peloquin L, Greer CW (1993) Cloning and expression of the polychlorinated biphenyl-degradation gene cluster from Arthrobacter M5 and comparison to analogous genes from Gram-negative bacteria. Gene 125:35–40 Pentyala SN, Rebecchi M, Mishra S, Rahman A, Stefen R, Rebecchi J, Kodavanti PS (2011) Polychlorinates biphenyls: In situ bioremediation from the environment. GR Reddy, SJF Flora, and RM Basha (ed.) Environ Pollut Ecol Hum Hlth, Narosa Publishing House, New Delhi, India, Chapter 1:249–262 Pieper DH, Seeger M (2008) Bacterial metabolism of polychlorinated biphenyls. J Mol Microbiol Biotechnol 15:121–138 Ponce BL, Latorre VK, González M, Seeger M (2011) Antioxidant compounds improved PCB-degradation by B. xenovorans strain LB400. Enzym Microb Technol 49:509–516 Raschke H, Meier M, Burken JG, Hany R, Muller MD, Van der Meer JR, Kohler HPE (2001) Biotransformation of various substituted aromatic compounds to chiral dihydrodihydroxy derivatives. Appl Environ Microbiol 67:3333–3339 Reineke W (1998) Development of hybrid strains for the mineralization of chloroaromatics by patchwork assembly. Annu Rev Microbiol 52:287–331 Rogers JE, Gibson DT (1977) Purification and properties of cis-toluene dihydrodiol dehydrogenase from Pseudomonas putida. J Bacteriol 130:1117–1124 Romero-Silva MJ, Méndez V, Agulló L, Seeger M (2013) Genomic and functional analyses of the gentisate and protocatechuate ring-cleavage pathways and related 3-hydroxybenzoate and 4-hydroxybenzoate peripheral pathways in Burkholderia xenovorans LB400. PLoS ONE 8: e56038 Ruder AM, Hein MJ, Hopf NB, Waters MA (2014) Mortality among 24,865 workers exposed to polychlorinated biphenyls (PCBs) in three electrical capacitor manufacturing plants: a ten-year update. Int J Hyg Environ Health 217:176–187 Ruzzini AC, Bhowmik S, Yam KC, Ghosh S, Bolin JT, Eltis LD (2013) The lid domain of the MCP hydrolase DxnB2 contributes to the reactivity toward recalcitrant PCB metabolites. Biochemistry 52(33):5685–5695 Saavedra JM, Acevedo F, González M, Seeger M (2010) Mineralization of PCBs by the genetically modified strain Cupriavidus necator JMS34 and its application for bioremediation of PCBs in soil. Appl Microbiol Biotechnol 87(4):1543–1554 Seah SY, Ke J, Denis G, Horsman GP, Fortin PD, Whiting CJ, Eltis LD (2007) Characterization of a C–C bond hydrolase from Sphingomonas wittichii RW1 with novel specificities towards polychlorinated biphenyl metabolites. J Bacteriol 189:4038–4045 Seah SYK, Labbe G, Nerdinger S, Johnson MR, Snieckus V, Eltis LD (2000) Identification of a serine hydrolase as a key determinant in the microbial degradation of polychlorinated biphenyls. J Biol Chem 275:15701–15708 Seah SYK, Labbe G, Kaschabek SR, Reifenrath F, Reineke W, Eltis LD (2001) Comparative specificities of two evolutionarily divergent hydrolases involved in microbial degradation of polychlorinated biphenyls. J Bacteriol 183:1511–1516 Seeger M, Timmis KN, Hofer B (1995a) Degradation of chlorobiphenyls catalyzed by the bphencoded biphenyl-2,3-dioxygenase and biphenyl-2,3-dihydrodiol-2,3-dehydrogenase of Pseudomonas sp. LB400. FEMS Microbiol Lett 133:259–264 Seeger M, Timmis KN, Hofer B (1995b) Conversion of chlorobiphenyls into phenylhexadienoates and benzoates by the enzymes of the upper pathway for polychlorobiphenyl degradation

Genetics and Biochemistry of Biphenyl and PCB Biodegradation

27

encoded by the bph locus of Pseudomonas sp. strain LB400. Appl Environ Microbiol 61:2654–2658 Seeger M, Cámara B, Hofer B (2001) Dehalogenation, denitration, dehydroxylation, and angular attack on substituted biphenyls and related compounds by a biphenyl dioxygenase. J Bacteriol 183:3548–3555 Seeger M, Zielinski M, Timmis KN, Hofer B (1999) Regiospecificity of dioxygenation of di- to pentachlorobiphenyls and their degradation to chlorobenzoates by the bph-encoded catabolic pathway of Burkholderia sp. strain LB400. Appl Environ Microbiol 65:3614–3621 Seeger M, González M, Cámara B, Muñoz L, Ponce E, Mejias L, Mascayano C, Vasquez Y, Sepulveda-Boza S (2003) Biotransformation of natural and synthetic isoflavonoids by two recombinant microbial enzymes. Appl Environ Microbiol 69:5045–5050 Seo J, Kang S, Kim M, Han J, Hur H-G (2011) Flavonoids biotransformation by bacterial non-heme dioxygenases, biphenyl and naphthalene dioxygenase. Appl Microbiol Biotechnol 91:219–228 Sikkema J, de Bont JAM, Poolman B (1995) Mechanisms of membrane toxicity of hydrocarbons. Microbiol Rev 59:201–222 Singer AC, Gilbert ES, Luepromchai E, Crowley DE (2000) Bioremediation of polychlorinated biphenyl-contaminated soil using carvone and surfactant-grown bacteria. Appl Microbiol Biotechnol 54(6):838–843 Sowers K, May HD (2013) In situ treatment of PCBs by anaerobic microbial dechlorination in aquatic sediment: are we there yet? Curr Opin Biotechnol 24(3):482–488 Springael D, Kreps S, Mergeay M (1993) Identification of a catabolic transposon, Tn4371, carrying biphenyl and 4-chlorobiphenyl degradation genes in Alcaligenes eutrophus A5. J Bacteriol 175:1674–1681 Stecker C, Johann A, Herzberg C, Averhoff B, Gottschalk G (2003) Complete nucleotide sequence and genetic organization of the 210-kilobase linear plasmid of Rhodococcus erythropolis BD2. J Bacteriol 185:5269–5274 Suenaga H, Nishi A, Watanabe T, Sakai M, Furukawa K (1999) Engineering a hybrid pseudomonad to acquire 3,4-dioxygenase activity for polychlorinated biphenyls. J Biosci Bioeng 87:430–435 Suenaga H, Watanabe T, Sato M, Ngadiman FK (2002) Alteration of regiospecificity in biphenyl dioxygenase by active-site engineering. J Bacteriol 184:3682–3688 Taguchi K, Motoyama M, Kudo T (2004) Multiplicity of 2,3-dihydroxybiphenyl dioxygenase genes in the Gram-positive polychlorinated biphenyl degrading bacterium Rhodococcus rhodochrous K37. Biosci Biotechnol Biochem 68:787–795 Taguchi K, Motoyama M, Iida T, Kudo T (2007) Polychlorinated biphenyl/biphenyl degrading gene clusters in Rhodococcus sp. K37, HA99, and TA431 are different from well-known bph gene clusters of Rhodococci. Biosci Biotechnol Biochem 71:1136–1144 Takeda H, Yamada A, Miyauchi K, Masai E, Fukuda M (2004) Characterization of transcriptional regulatory genes for biphenyl degradation in Rhodococcus sp. strain RHA1. J Bacteriol 186:2134–2146 Taylor PM, Medd JM, Schoenborn L, Hodgson B, Janssen PH (2002) Detection of known and novel genes encoding aromatic ring- hydroxylating dioxygenases in soils and in aromatic hydrocarbon-degrading bacteria. FEMS Microbiol Lett 216:61–66 Toussaint A, Merlin C, Monchy S, Benotmane MA, Leplae R, Mergeay M, Springael D (2003) The biphenyl- and 4-chlorobiphenyl-catabolic transposon Tn4371, a member of a new family of genomic islands related to IncP and Ti plasmids. Appl Environ Microbiol 69:4837–4845 Triscari-Barberi T, Simone D, Calabrese FM, Attimonelli M, Hahn KR, Amoako KK, Turner RJ, Fedi S, Zannonia D (2012) Genome sequence of the polychlorinated-biphenyl degrader Pseudomonas pseudoalcaligenes KF707. J Bacteriol 194(16):4426–4427 Tu C, Teng Y, Luo Y, Li X, Sun X, Li Z, Liu W, Christie P (2011) Potential for biodegradation of polychlorinated biphenyls (PCBs) by Sinorhizobium meliloti. J Hazard Mater 186 (2–3):1438–1444

28

L. Agulló et al.

Vaillancourt FH, Labbe G, Drouin NM, Fortin PD, Eltis LD (2002) The mechanism-based inactivation of 2,3-dihydroxybiphenyl 1,2- dioxygenase by catecholic substrates. J Biol Chem 277:2019–2027 Vezina J, Barriault D, Sylvestre M (2007) Family shuffling of soil DNA to change the regiospecificity of Burkholderia xenovorans LB400 biphenyl dioxygenase. J Bacteriol 189:779–788 Watanabe T, Inoue R, Kimura N, Furukawa K (2000) Versatile transcription of biphenyl catabolic bph operon in Pseudomonas pseudoalcaligenes KF707. J Biol Chem 275:31016–31023 Witzig R, Junca H, Hecht HJ, Pieper DH (2006) Assessment of toluene/biphenyl dioxygenase gene diversity in benzene-polluted soils: links between benzene biodegradation and genes similar to those encoding isopropylbenzene dioxygenases. Appl Environ Microbiol 72:3504–3514 Wu Q, Watts JE, Sowers KR, May HD (2002) Identification of a bacterium that specifically catalyzes the reductive dechlorination of polychlorinated biphenyls with doubly flanked chlorines. Appl Environ Microbiol 68:807–812 Xu Y, Yu M, Shen A (2016) Complete genome sequence of the polychlorinated biphenyl degrader Rhodococcus sp. WB1. Genome Announc 4(5):e00996–e00916 Xu L, Teng Y, Li ZG, Norton JM, Luo YM (2010) Enhanced removal of polychlorinated biphenyls from alfalfa rhizosphere soil in a field study: the impact of a rhizobial inoculum. Sci Total Environ 408(5):1007–1013 Yamada A, Kishi H, Sugiyama K, Hatta T, Nakamura K, Masai E, Fukuda M (1998) Two nearly identical aromatic compound hydrolase genes in a strong polychlorinated biphenyl degrader, Rhodococcus sp. strain RHA1. Appl Environ Microbiol 64:2006–2012 Yang X, Liu X, Xie F, Zhang G, Qian S (2007) Characterization and functional analysis of a novel gene cluster involved in biphenyl degradation in Rhodococcus sp. strain R04. J Appl Microbiol 103:2214–2224 Yu C, Liu W, Ferraro D, Brown E, Parales JV, Ramaswamy S, Zylstra GJ, Gibson DT, Parales RE (2007) Purification, characterization and crystallization of the components of a biphenyl dioxygenase system from Sphingobium yanoikuyae B1. J Ind Microbiol Biotechnol 34:311–324 Zielinski M, Kahl S, Standfuss-Gabisch C, Cámara B, Seeger M, Hofer B (2006) Generation of novel-substrate-accepting biphenyl dioxygenases through segmental random mutagenesis and identification of residues involved in enzyme specificity. Appl Environ Microbiol 72:2191–2199

Phylogenomics of Aerobic Bacterial Degradation of Aromatics D. Pérez-Pantoja, R. Donoso, H. Junca, B. González, and D. H. Pieper

Abstract

Aromatic compounds are widely distributed in nature. They are found as lignin components, aromatic amino acids, and xenobiotic compounds, among others. Microorganisms, mostly bacteria, degrade an impressive variety of such chemical structures. Various aerobic aromatic catabolic pathways have been reported in bacteria, which typically consist of activation of the aromatic ring through oxygenases or CoA ligases and ring cleavage of di- or trihydroxylated intermediates or dearomatized CoA derivatives. We survey almost 900 sequenced bacterial genomes available in 2008 for the presence of genes encoding key enzymes of aromatic metabolic pathways, including ring-cleavage enzymes as well as enzymes activating aromatics or dearomatizing CoA derivatives. The metabolic diversity is discussed from two angles: the spread of such key activities among different bacterial phyla and the overall metabolic potential of members of bacterial genera.

D. Pérez-Pantoja Departamento de Bioquímica y Biología Molecular, Facultad de Ciencias Biológicas, Universidad de Concepción, Concepción, Chile e-mail: [email protected] R. Donoso • B. González Facultad de Ingeniería y Ciencias, Universidad Adolfo Ibáñez, Santiago, Chile e-mail: [email protected]; [email protected] H. Junca Research Group Microbial Ecology: Metabolism, Genomics & Evolution, Microbiomas Foundation, Chia, Colombia e-mail: [email protected] D.H. Pieper (*) Microbial Interactions and Processes Research Group, HZI – Helmholtz Centre for Infection Research, Braunschweig, Germany e-mail: [email protected] # Springer International Publishing AG 2016 F. Rojo (ed.), Aerobic Utilization of Hydrocarbons, Oils and Lipids, Handbook of Hydrocarbon and Lipid Microbiology , DOI 10.1007/978-3-319-39782-5_33-1

1

2

D. Pérez-Pantoja et al.

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 2 Aerobic Aromatic Catabolic Routes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 3 Sequenced Bacterial Genomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 4 Spread of Members of Gene Families . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 5 Metabolism Diversity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42

1

Introduction

A few non-mutually exclusive choices are possible to address the analysis of the genetic basis of bacterial degradation of aromatic compounds. One is to select a few well-studied bacterial catabolic models and go in depth into their genetic organization of aromatic catabolism genes (Jimenez et al. 2002; Pérez-Pantoja et al. 2008). Another approach is to select a few central catabolic pathways and to assess the similarities and differences in gene organization, substrate range, and regulatory elements, among the bacteria where such pathways have been described. A third possibility is to look for all the aromatic catabolism pathways present in bacteria, searching in the growing database of sequenced bacterial genomes. The latter, by definition, is a less in-depth analysis but has the broader coverage possible today. We selected the latter approach, because we think it provides clues on the distribution of catabolic properties among bacterial phyla, gives some hints on the ecological functions of specific bacterial groups, defines underscored research objectives, and gives a better overview of the genetic basis of bacterial catabolism of aromatics. The phylogenomic approach to study the organization of aromatic degradation is based on the selection of sequences of key catabolic functions to fish into the sequenced genome database, followed by refinement of the positive scores. With this information, the genomes can be analyzed in terms of presence/absence of catabolic abilities among bacterial groups, new enzyme families based on the sequence similarity be defined, new putative functions be suggested, and evolutionary links among different groups of sequences be addressed (for an appropriate novel database see Duarte et al. 2014). Of course such approach has some limitations, as most of the new data are not supported by biochemical or genetic studies. To minimize such limitations, the selected sequence probes were derived from both biochemical and genetic well-studied systems. One of the main purposes of the following material is to provide to the reader new research venues to get a deeper knowledge on bacterial catabolism of aromatics.

2

Aerobic Aromatic Catabolic Routes

Bacterial degradation of aromatic compounds and their haloaromatic derivatives has been well studied (Duarte et al. 2014). Various pathways for degradation of these compounds by bacteria have been reported. The activation of the aromatic ring

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

3

commonly proceeds by members of one of three superfamilies: the Rieske non-heme iron oxygenases usually catalyzing the incorporation of two oxygen atoms (although some members of this superfamily also catalyze monooxygenations) (Gibson and Parales 2000), the flavoprotein monooxygenases (van Berkel et al. 2006), and the soluble diiron multicomponent oxygenases (Leahy et al. 2003). Further metabolism is achieved through di- or trihydroxylated aromatic intermediates. Alternatively, activation is mediated by CoA ligases and the formed CoA derivatives are subjected to oxygenations. This can proceed through 2-aminobenzoyl-CoA monooxygenase/ reductase, an enzyme that catalyzes both monooxygenation and hydrogenation, and where the N-terminal part of the protein shows similarities to single-component flavin monooxygenases (Buder and Fuchs 1989). Alternatively, the aromatic CoA derivative is attacked by multicomponent enzymes, where the oxygenase subunits belong to the diiron oxygenases, like in phenylacetyl-CoA (Ismail et al. 2003) or benzoyl-CoA oxygenase (Zaar et al. 2004). Various further key reactions channeling aromatics to central di- or trihydroxylated intermediates, such as the processing of side chains or demethylations, will not be discussed here. The further aerobic degradation of di- or trihydroxylated intermediates can be catalyzed by either intradiol or extradiol dioxygenases. While all intradiol dioxygenases described thus far belong to the same superfamily, members of at least three different families are reported to be involved in the extradiol ring cleavage of hydroxylated aromatics. Type I extradiol dioxygenases (e.g., catechol 2,3-dioxygenases) belong to the vicinal oxygen chelate superfamily enzymes (Gerlt and Babbitt 2001), the type II or LigB superfamily of extradiol dioxygenases which comprise among other protocatechuate 4,5-dioxygenases (Sugimoto et al. 1999) and the type III enzymes such as gentisate dioxygenases which comprise enzymes belonging to the cupin superfamily (Dunwell et al. 2000). However, even though belonging to different families, all three types of extradiol dioxygenases share similar active sites and all type I, type II, and various type III enzymes have the same iron ligands, two histidine and one glutamate, that constitute the 2-His 1-carboxylate structural motif. The benzoquinol 1,2-dioxygenase from the 4-hydroxyacetophenone-degrading Pseudomonas fluorescens ACB that displays no significant sequence identity with known dioxygenases may constitute the prototype of a novel fourth class of Fe 2+-dependent dioxygenases (Moonen et al. 2008).

3

Sequenced Bacterial Genomes

As of September 2008 approximately 1,000 genomes have been sequenced and three quarters of them finished. For the purpose of this review, we concentrated on genomes that were simultaneously represented in both the Integrated Microbial Genomes (IMG) database at DOE Joint Genome Institute (JGI) (img.jgi.doe.gov/cgi-bin/pub/main.cgi? page=home) and the National Center for Biotechnology Information (NCBI) database at National Institute of Health (NIH) (www.ncbi.nlm.nih.gov/sutils/genom_table.cgi), summing up to 822 genomes. The number of representatives of the bacterial phyla in these public databases was highly variable: from a very few members from the phyla

4

D. Pérez-Pantoja et al.

Aquificae (2) Acidobacteria (2), Chlamydiae (11), Chlorobi (10), Chloroflexi (8), Deinococcus/Thermus (4), Fusobacteria (2), Lentisphaerae (2), Planctomycetes (3), Spirochaetes (9), Thermotogae (6), and Verrucomicrobia (1); the medium represented phyla: Actinobacteria (53), Bacteroidetes (28), Cyanobacteria (40), and the Proteobacteriales δ- (23) and ε- classes (28); and the highly represented phylum Firmicutes (182) and the α- (112) β- (71) and γ- (223) classes of Proteobacteria (besides two unclassified Proteobacteria). Despite of that, the number of bacterial genomes is now significant to search for the presence/absence of the main catabolic pathways for aromatic compounds to provide a reasonable idea about the spread of these catabolic abilities among the main phylogenetic groups.

4

Spread of Members of Gene Families

4.1

Intradiol Dioxygenases

The intradiol cleavage of catechol to muconate and of protocatechuate to 3-carboxymuconate by catechol 1,2-dioxygenases and protocatechuate 3,4-dioxygenases, respectively, is a central reaction in the metabolism of various aromatic compounds (Fig. 1). Hydroxybenzoquinol (1,2,4-trihydroxybenzene) is also a central intermediate in the degradation of a variety of aromatic compounds such as resorcinol (Fig. 1), with hydroxybenzoquinol 1,2-dioxygenase as key enzyme, catalyzing intradiol cleavage to form 3-hydroxy-cis,cis-muconate and its tautomer, maleylacetate. Among the different groups of enzymes significant metabolic cross-reactivity is usually not observed. Phylogenetic analysis of the deduced protein sequences of intradiol dioxygenases encoded in the genomes of bacteria sequenced so far showed the presence of seven clusters as indicated in Fig. 2. Based on biochemical or genetically validated representatives, cluster 1 comprises hydroxybenzoquinol dioxygenases, cluster 2 proteobacterial catechol 1,2-dioxygenases, cluster 3 actinobacterial catechol 1,2-dioxygenases, and clusters 5 and 7 the α- and β-subunits of protocatechuate 3,4-dioxygenases, respectively. Enzymes of cluster 6 are obviously related to the β-subunits of protocatechuate dioxygenases, however, in no case genes encoding these enzymes are clustered with genes encoding putative α- subunits, and the function of these enzymes remains to be elucidated. Similarly, the function of enzymes of cluster 4 wait for clarification. Intradiol dioxygenases are nearly exclusively found in two phyla, the Actinobacteria and the Proteobacteria. However, protocatechuate 3,4-dioxygenases were observed in one of the two sequenced Deinococci, i.e., Deinococcus geothermalis DSM 11300 and one of the two sequenced Acidobacteria, i.e., Solibacter usitatus Ellin6076. Considering the wide spread of Acidobacteria in the environment, their involvement in aromatic degradation under natural conditions has to be considered. Actually, Acidobacteria have been implied to be involved in the biogeochemical cycles of rhizosphere soil (Lee et al. 2008). Regarding catechol 1,2-dioxygenases, where two lineages have previously been described (Eulberg et al. 1997), phylogenetic analysis confirmed that cluster

Fig. 1 Aerobic metabolism of aromatics via di- or trihydroxylated intermediates, or via CoA derivatives. Peripheral hydroxylation reactions can be catalyzed by flavoprotein monooxygenases, Rieske non-heme iron oxygenases or soluble diiron oxygenases. Alternatively, aromatics can be activated through CoA ligases

Phylogenomics of Aerobic Bacterial Degradation of Aromatics 5

6

D. Pérez-Pantoja et al.

3 enzymes are restricted to members of the order Actinomycetales of the Actinobacteria, and catechol intradiol cleavage pathways were observed in the majority of Corynebacteria, Arthrobacter, Mycobacteria, and Nocardiaceae. Usually, Actinobacteria possessing a catechol intradiol cleavage pathway also harbor a protocatechuate intradiol cleavage. However, Streptomyces strains seem to be endowed only with the protocatechuate branch. A hydroxybenzoquinol pathway seems to

Fig. 2 Evolutionary relationships among intradiol dioxygenases. The evolutionary history was inferred using the neighbor joining method after alignment of sequences using MUSCLE (Edgar 2004). All positions containing alignment gaps and missing data were eliminated only in pairwise sequence comparisons. Wedges represent enzyme clusters as described in the text. Deduced protein sequences not falling inside the defined clusters are also indicated. Wedge length is a measure of evolutionary distance from the common ancestor. Phylogenetic analyses were conducted in MEGA4 (Tamura et al. 2007). Cluster 1 comprises hydroxybenzoquinol dioxygenases, cluster 2 proteobacterial catechol 1,2-dioxygenases, cluster 3 actinobacterial catechol 1,2-dioxygenases, and clusters 5 and 7 the α- and β-subunits of protocatechuate 3,4-dioxygenases, respectively. The functions of enzymes of clusters 4 and 6 remain to be elucidated

ä Fig. 1 (continued) followed by dearomatization catalyzed by members of the flavoprotein monooxygenases or soluble diiron oxygenases. 4-Hydroxyphenylpyruvate dioxygenase is indicated by a Central di- or trihydroxylated intermediates are subjected to ring cleavage by intradiol dioxygenases or extradiol dioxygenases of the vicinal chelate superfamily, the LigB superfamily or the cupin superfamily. Ring-cleavage products are channeled to the Krebs cycle via central reactions

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

7

be spread only in Corynebacteria and out of the Mycobacteria, only Mycobacterium smegmatis and M. vanbaalenii are endowed with such a pathway. As shown in Table 1, intradiol dioxygenases can be identified in 11 out of 19 α-proteobacterial, 2 out of 10 β-proteobacterial, and 4 out of 29 γ-proteobacterial families and are absent in δ- or ε- proteobacteria. Significant differences in gene spread were observed among families. Catechol intradiol pathways are observed in nearly all Pseudomonas strains and are absent only from the genomes of P. syringae and P. mendocina. The last one is also the only Pseudomonas strain devoid of a protocatechuate intradiol pathway. Similarly, both protocatechuate and catechol pathways are observed in all Burkholderia genomes. Interestingly, catechol intradiol cleavage pathways were only exceptionally observed in α-Proteobacteria. In contrast, a catechol pathway is absent in Rhizobiaceae, which, however, often bear a hydroxybenzoquinol pathway. Also Bradyrhizobiaceae, none of which has a catechol pathway, are usually endowed with a hydroxybenzoquinol pathway except for Nitrobacter strains.

4.2

EXDO I Family

The extradiol ring cleavage of catechol is typically catalyzed by type I extradiol dioxygenases (EXDO I), which belong to the vicinal oxygen chelate superfamily (Gerlt and Babbitt 2001). The EXDO I family comprises enzymes that catalyze the dioxygenolytic ring fission of the catecholic derivatives in several bacterial monoand polyaromatics biodegradation pathways (Eltis and Bolin 1996; Duarte et al. 2014) (Fig. 1) like those involved in degradation of benzene, toluene, phenol, biphenyl, naphthalene, dibenzofuran, 4-hydroxyphenylacetate, p-cymene, or diterpenoid compounds such as abietate. They catalyze the meta-cleavage of catechol to 2-hydroxymuconic semialdehyde (catechol 2,3-dioxygenases, C23O), of 2,3-dihydroxybiphenyl (2,3-dihydroxybiphenyl 1,2-dioxygenases, BphC), 1,2-dihydroxynaphthalene (NahC), homoprotocatechuate (homoprotocatechuate 2,3-dioxygenases, HpaD), 2,3-dihydroxy-p-cumate (2,3-dihydroxy-p-cumate-3,4dioxygenases CmtC), and 7-oxo-11,12-dihydroxydehydroabietate (DitC), among others (see also Fig. 1). In many cases, the respective genes are localized in catabolic pathway gene clusters such that their actual function can easily be deduced. However, in various cases multiple EXDO I activities are observed in a single strain and often their function remains unproven (Maeda et al. 1995). Here, the names are given to the enzymes according to the preferential activity observed, but in many cases there is a range of structurally similar substrates that can be metabolized by the same enzyme with varying catalytic efficacies and the “natural” substrate has not yet been identified. Because genome annotations pipelines are in many cases using the NCBI Conserved Domains Database (CDD), which is in turn, interconnected with the Wellcome Trust Sanger Institute Pfam database descriptions, all EXDO I genes found in the genome sequences are recognized and annotated with the superfamily

8

D. Pérez-Pantoja et al.

Table 1 Intradiol dioxygenases observed in genomes of Proteobacteria

Class α α α α α α α α α α α β β γ γ γ γ

Family Caulobacteraceae (2) Aurantimonadaceae (2) Bradyrhizobiaceae (11) Brucellaceae (6) Methylobacteriaceae (3) Phyllobactriaceae (3) Rhizobiaceae (6) Rhodobacteraceae (24) Xanthobacteriaceae (2) Acetobacteraceae (3) Sphingomonadaceae (5) Burkholderiaceae (43) Comamonadaceae (8) Oceanosprillaceae (3) Moraxellaceae (5) Pseudomonadaceae (19) Xanthomonadaceae (11)

Protocatechuate 3,4-dioxygenase (Pca34) ++ +

Catechol 1,2-dioxygenase (Cat12)

( ) ++ +

Hydroxybenzoquinol dioxygenase (Hqu)

++

+

++ ++ ++

++ +

++

+

++

+ ( )

+

( )

++

++

+

( )

+

+

+

+

+ ++

( )

+ ++ +

++; More than 60% of the sequenced genomes of these bacterial taxa comprise a gene encoding the mentioned activity (number of sequenced representatives is given in parentheses); +, between 20% and 60%; ( ), less than 20%; , not observed

name as Glyoxalase/bleomycin resistance protein/dioxygenase (InterPro: IPR004360, pfam00903: Glyoxalase). However, in the majority of cases, a more precise annotation of several genomic sequences as EXDO I would be possible, as they show conservation of the Prosite PS00082 extradiol ring-cleavage dioxygenases signature [GNTIV]-x-H-x(5,7)-[LIVMF]-Y-x(2)-[DENTA]-P-x[GP]-x(2,3)-E. Phylogenetic analysis of the deduced protein sequences of EXDO I encoded in the genomes of bacteria sequenced so far, and retrieved after iterative PSI Blast searches using representative proteins of major clusters where a function has been described as seeds show the presence of three major evolutionary lineages (Fig. 3).

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

9

One of these lineages (cluster 1) comprises nearly all EXDO I proteins of validated function. Ten subclusters (A–J) grouping proteins associated with different substrate specificities can be differentiated. Subcluster 1A comprises enzymes experimentally validated as C23O. Interestingly, there is a high redundancy in genomes, as the 28 identified genes are observed in only 18 strains. Out of these, 13 strains belong to the β-proteobacteria and C230 is mainly observed in Burkholderia, Cupriavidus, and Ralstonia genomes. This contrasts previous reports on C23Os, which were predominantly characterized from Pseudomonas strains (Eltis and Bolin 1996). However, in none of the sequenced Pseudomonas a homologous gene is observed. It has, however, to be noted that most of such genes have previously been reported on plasmids rather than in the chromosome of the strains, such as the case for P. putida KT2440 where the IncP-9 TOL plasmid pWW0 is present (Williams and Murray 1974), but not included in the same genome project. It is also interesting to note that the Actinobacterium R. jostii RHA1 has a predicted C23O of this kind. Subcluster 1B groups putative homoprotocatechuate 2,3-dioxygenases of the actinobacterial lineage. As expected from literature, the respective encoding genes are present in Actinobacteria (Vetting et al. 2004), and observed in 5 out of 53 genomes. They are absent from any β- and γ-proteobacterial genomes, but surprisingly most abundant in α-proteobacterial genomes (16 genomes), specifically in Bradyrhizobiaceae and Rhodobacteraceae, even though proteobacterial homoprotocatechuate 2,3-dioxygenases are generally assumed to be members of the LigB family (see below) (Roper and Cooper 1990). It is also interesting to note that such genes were found also outside the Actinobacteria and Proteobacteria, and are present in both sequenced Deinococcus and in both sequenced Thermus strains as well as in three Bacillaceae. Subcluster 1C groups proteins related to BphC of Bacillus sp. JF8 involved in biphenyl degradation by this strain (Hatta et al. 2003). Related proteins are not encoded in any of the sequenced Bacilli, but astonishingly in all four genomes available of Chloroflexaceae strains and in a few actinobacterial species, including one protein of R. jostii RHA1, however, not having a taxonomically linked distribution in lower levels. Similarly, proteins related to NahC 1,2-dihydroxynaphthalene dioxygenase of Bacillus sp. JF8 (Miyazawa et al. 2004) (subcluster 1D) are not observed in any Bacillus species, but encoded in four α-proteobacterial genomes. Also the three subcluster 1E proteins, where no closely related proteins have been characterized so far, are encoded in two α-proteobacterial genomes. Subcluster 1F proteins are encoded by all 34 genomes available of Burkholderia and various other proteobacterial genomes, however, their actual function still remains to be elucidated. Subcluster 1G comprises proteins such as DntD of Burkholderia sp. DNT responsible for meta-cleavage of trihydroxytoluene, which is also active on catechol (Haigler et al. 1999) but includes as well various proteins of proven activity against 2,3-dihydroxybiphenyl such as BphC3 and BphC4 of R. jostii RHA1, both being reported as being practically inactive with catechol (Sakai et al. 2002). Similar proteins are mainly observed in genomes of Actinobacteria, with R. jostii RHA1

10

D. Pérez-Pantoja et al.

Fig. 3 Evolutionary relationships among type I extradiol dioxygenases (EXDO I). Subcluster 1A comprises catechol 2,3-dioxygenases, subcluster 1B putative homoprotocatechuate 2,3-dioxygenases, subcluster 1C proteins related to BphC of Bacillus sp. JF8, subcluster 1D proteins related to NahC of Bacillus sp. JF8, subcluster 1G proteins related to DntD of Burkholderia sp. DNT or BphC3 and BphC4 of R. jostii RHA1, subcluster 1H proteins similar to those capable to cleave 2,3-dihydroxy-p-cumate, subcluster 1I proteins related to those involved in diterpenoid degradation, and subcluster 1J enzymes with similarities to those being active mainly against bicyclic and higher condensed dihydroxylated aromatics. Subcluster 2B comprises so-called one-domain extradiol dioxygenases and cluster 3 proteins related to LinE chlorobenzoquinol 1,2-dioxygenases and PcpA 2,6-dichlorobenzoquinol 1,2-dioxygenases. However, the function of the majority of enzymes of cluster 3 as well as of enzymes of subclusters 1E, 1F, and 2A remains to be elucidated

harboring three of such genes, and α- and β-Proteobacteria. Proteins similar to those capable to cleave 2,3-dihydroxy-p-cumate (subcluster 1H) are only found in four genomes including P. putida F1 reported to exhibit such activity (Eaton 1996) and B. xenovorans LB400, indicating that it is not a widespread activity. Similarly, proteins related to those involved in diterpenoid degradation (subcluster 1I) (Martin and Mohn 2000) are not common in the genomes analyzed, showing only hits in Caulobacter sp. K31 and the already described activity of B. xenovorans LB400 (Smith et al. 2007). Subcluster 1J comprises a variety of enzymes with similarities to members of subfamilies I.4, I.5, and I.3.E being active mainly against bicyclic and higher condensed dihydroxylated aromatics (Eltis and Bolin 1996). An overall of 68 such proteins could be observed to be encoded in thus far sequenced genomes. Respective genes are observed in 11 of 17 Mycobacterial genomes, which is not astonishing, as various sequenced Mycobacteria were selected for their capability to mineralize polycyclic aromatics. They are also observed in all three Nocardiaceae

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

11

genomes, with R. jostii RHA1 harboring six such genes. In addition, eight α-, eight β-, and five γ-proteobacterial strains harbor such enzyme. Out of the Pseudomonas, it was observed only in the P. putida F1 genome (Zylstra et al. 1988). The majority of the approximately 100 protein sequences conforming cluster 2 contain the Prosite PS00082 extradiol ring-cleavage dioxygenase signature described above. Subcluster 2B comprises BphC6 of R. jostii RHA1 (ABO34703) and other previously characterized so-called one-domain extradiol dioxygenases such as BphC2 and BphC3 from R. globerulus P6 with reported activity against 2,3-dihydroxybiphenyl (Asturias and Timmis 1993) (subfamily I.1 as defined by Eltis and Bolin (Eltis and Bolin 1996)). However, besides BphC6 of strain RHA1, no further enzyme of this type was found to be encoded in the genomes analyzed, and proteins with similarity to subcluster 2A proteins have not yet been functionally characterized. Ring-cleavage dioxygenases involved in the turnover of (chloro)benzoquinols and (chloro)hydroxybenzoquinols have been identified from various microorganisms degrading γ-hexachlorocyclohexane or chlorophenols, and comprise LinE chlorobenzoquinol/benzoquinol 1,2-dioxygenases, which preferentially cleaves aromatic rings with two hydroxyl groups at para positions (Miyauchi et al. 1999) and PcpA 2,6-dichlorobenzoquinol 1,2-dioxygenases (Xu et al. 1999). These proteins are comprised in cluster 3, and are the only validated extradiol dioxygenases observed in this cluster. Compared to cluster 1, cluster 3 is so divergent that even the Superfam HMM system recognizes the validated LinE/PcpA sequences as part of the Glyoxalase/bleomycin resistance protein/dioxygenase superfamily but belonging to the family of Glyoxalase I (lactoylglutathione lyase). Only the genomes of Cupriavidus necator H16 and JMP134 contain sequences that may have encode chlorobenzoquinol dioxygenases. It should be noted that one of the sequences of C. necator JMP134 is clustered with a gene similar to the one described from P. putida HS12 encoding nitrobenzene nitroreductase, which is also clustered with a putative benzoquinol extradiol dioxygenase (Park and Kim 2000).

4.3

Lig B Superfamily

A second family of extradiol dioxygenases is the so-called LigB family (Sugimoto et al. 1999; Duarte et al. 2014). LigB type extradiol dioxygenases are well established as being responsible for the degradation of protocatechuate via the protocatechuate 4,5-dioxygenase pathway. Protocatechuate dioxygenases are composed of two distinct subunits, with the active site being located in the β-subunit. Also, proteobacterial homoprotocatechuate 2,3-dioxygenases as the one described in Escherichia coli (Roper and Cooper 1990) belong to the type II or LigB superfamily of extradiol dioxygenases whereas actinobacterial homoprotocatechuate 2,3-dioxygenases are supposed to belong to the EXDO I (Vetting et al. 2004). A further well-documented group of LigB-type extradiol dioxygenases are the 2,3-dihydroxyphenylpropionate 1,2-dioxygenases which, like LigB-type homoprotocatechuate dioxygenases, consist only of one type of subunit (Diaz et al.

12

D. Pérez-Pantoja et al.

2001). Recent analyses have revealed various other substrates that are cleaved by LigB-type extradiol dioxygenases. Aminophenol 1,6-dioxygenases (Fig. 1) are like protocatechuate 4,5-dioxygenases, composed of two distinct subunits, with the β-subunits containing the active site (Takenaka et al. 1997). Gallate dioxygenases have so far been described in S. paucimobilis SYK-6 (Kasai et al. 2005) and P. putida KT2440 (Nogales et al. 2005), and are specific for this substrate and do not transform protocatechuate, whereas gallate transformation by protocatechuate 4,5-dioxygenases has been reported. Both gallate dioxygenases have sizes significantly larger than those of the β-subunits of protocatechuate dioxygenases. Analysis of the primary structure revealed that the N-terminal regions showed a significant amino acid sequence identity with the β-subunit of protocatechuate 4,5-dioxygenases, whereas the C-terminal region has similarity to the corresponding small α-subunit (Nogales et al. 2005). It was therefore suggested that gallate dioxygenases are two-domain proteins that have evolved from the fusion of large and small subunits. Additional LigB-type enzymes have been described to be involved in the degradation of methylgallate (Kasai et al. 2004) or of bi- and polycyclic aromatics (Laurie and Lloyd-Jones 1999). Phylogenetic analysis of the deduced protein sequences of LigB-type proteins encoded in the genomes of bacteria sequenced so far allowed the identification of six clusters (Fig. 4). Cluster 1 comprises three subclusters, which contain protocatechuate 4,5-dioxygenase β-subunits (Fig. 4, cluster 1A), gallate dioxygenases (cluster 1B), and a group of related proteins where no member has been characterized thus far (cluster 1C). Respective genes were nearly exclusively observed in α-, β-, and γ-Proteobacteria and only 1 of the 53 analyzed actinobacterial genomes (Arthrobacter sp. FB24) has a protocatechuate 4,5-dioxygenase encoding gene. Protocatechuate 4,5-dioxygenases are predominantly observed in Comamonadaceae and Bradyrhizobiaceae, specifically Bradyrhizobium and Rhodopseudomonas strains and are mainly composed of two distinct subunits as evidenced by two subsequent genes encoding the respective subunits. However, putative gene fusions are observed in Arthrobacter and Verminephrobacter. Even though one of the two gallate dioxygenases characterized so far was reported in a Sphingomonas strain (Kasai et al. 2005), gallate dioxygenase encoding genes are not observed in any of the 112 sequenced α-Proteobacteria and are thus not a dominant trait in this group. In contrast, gallate dioxygenases are obviously encoded in the genomes of three of four sequenced P. putida strains. The supposed gallate dioxygenases are mainly fusions of α- and β-subunits, like in P. putida KT2440 (Nogales et al. 2005), however, seem to consist of separate subunits in Xanthomonas and Chromohalobacter. Dioxygenases belonging to the third subcluster are usually composed of α- and β-subunits, and are in 10 out of 12 cases encoded in genomes, which also encode a protocatechuate 4,5-dioxygenase pathway. A second cluster (cluster 2, Fig. 4) comprises enzymes most closely related to those involved in bi- and polycyclic aromatic degradation such as PhnC involved in the degradation of polycyclic aromatics by Burkholderia sp. strain RP007 (Laurie and Lloyd-Jones 1999), CarBb involved in the degradation of carbazol by P. resinovorans

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

13

Fig. 4 Evolutionary relationships among LigB-type dioxygenases. Subcluster 1A comprises protocatechuate 4,5-dioxygenase β-subunits, subcluster 1B gallate dioxygenases, cluster 2, enzymes most closely related to PhnC of Burkholderia sp. strain RP007 or CarBb of P. resinovorans CA10, cluster 3 enzymes related to DesZ of Sphingomonas paucimobilis SYK-6, cluster 4 2,3-dihydroxyphenylpropionates 1,2-dioxygenases, cluster 5 the β- and α-subunits (clusters 5A and B, respectively) of 2-aminophenol 1,6-dioxygenases, and cluster 6 homoprotocatechuate 2,3-dioxygenases. The function of enzymes of subcluster 1C remains to be elucidated

CA10 (Sato et al. 1997a), or BphC6 involved in the degradation of fluorene by Rhodococcus rhodochrous K37 (Taguchi et al. 2004). However, no clear association with a capability to degrade such compounds was evident, and the respective enzymes are spread among very different groups of Actinobacteria and Proteobacteria. The corresponding genes are absent from strains selected for genome sequencing due to their exceptional capability to degrade aromatics such as M. vanbaalenii Pyr, M. gilvium PYR-GCK, R. jostii RHA1, or B. xenovorans LB400. Cluster 3 comprises enzymes related to DesZ methylgallate dioxygenase of Sphingomonas paucimobilis SYK-6, where 7 out of 11 proteins are observed in Mycobacterium strains, however, their function remains to be elucidated. A fourth cluster obviously comprises 2,3-dihydroxyphenylpropionate 1,2-dioxygenases. The respective enzymes are most dominantly observed to be encoded in the genomes of Enterobacteriaceae, and specifically observed in 13 out of 18 E. coli strains sequenced and in Shigella sonnei. Interestingly, related enzymes are also observed to be encoded by 9 out 17 Mycobacterial genomes. Their function, however, remains to be proven.

14

D. Pérez-Pantoja et al.

A fifth cluster comprises 2-aminophenol 1,6-dioxygenases (Fig. 4, clusters 5A and B comprising the β- and α-subunits, respectively). Only two of these enzymes are observed to be encoded by previously sequenced genomes, i.e., B. xenovorans LB400 and P. putida W619, indicating such pathways to be present only in very few specialized bacteria. In contrast, homoprotocatechuate 2,3-dioxygenases (cluster 6) are observed to be widespread, and in contrast to previous assumptions that LigBtype homoprotocatechuate 2,3-dioxygenases were restricted to proteobacteria, homologues are also observed in two Actinobacteria, and the genomic context suggest that those enzymes actually are part of a functional homoprotocatechuate pathway. A homologue is also observed in Bacillus licheniformis.

4.4

Cupin Dioxygenases

Several extradiol dioxygenases of aromatic degradation pathways have been described to belong to the cupin superfamily (Dunwell et al. 2000; Duarte et al. 2014) sharing a common architecture and including key enzymes such as gentisate 1,2-dioxygenase (involved in the degradation of salicylate or 3-hydroxybenzoate, Fig. 1), homogentisate 1,2-dioxygenase (involved in the degradation of phenylalanine and tyrosine) (Arias-Barrau et al. 2004) and 3-hydroxyanthranilate 3,4-dioxygenase (involved in tryptophan degradation) (Kurnasov et al. 2003; Muraki et al. 2003). The phylogenomic analysis of this type of dioxygenases in the genomes of bacteria sequenced so far shows that homogentisate dioxygenase is the enzyme with the broadest distribution in bacterial families. This may be explained by the key role in the degradation of the aromatic amino acids phenylalanine and tyrosine in several organisms, including eukaryotes. Putative genes encoding this enzyme are strongly represented in Proteobacteria, being identified in 10 out of 19 α-, 5 out of 10 β-, 16 out of 29 γ-, and 4 out of 11 δ-proteobacterial families, although they were absent in ε-proteobacteria. In the families Bradyrhizobiaceae, Rhizobiaceae, Alcaligenaceae, Burkholderiaceae, Shewanellaceae, Legionellaceae, Pseudomonadaceae, and Vibrionaceae, a respective gene can be observed in nearly all genomes sequenced. Homogentisate 1,2-dioxygenase was the unique aromatic ring-cleavage enzyme found in sequenced representatives of the families Hyphomonadaceae, Neisseriaceae, Aeromonadaceae, Idiomarinaceae, Moritellaceae, Chromatiaceae, Legionellaceae, Hahellaceae, Bdellovibrionaceae, Cystobacteraceae, and Nannocystaceae. In addition, genes putatively encoding homogentisate 1,2-dioxygenase are also found in members of the non-proteobacterial orders Actinomycetales, Flavobacteriales, Sphingobacteriales, and Bacillales. Gentisate 1,2-dioxygenase is the ring-cleavage enzyme involved in catabolism of salicylate and 3-hydroxybenzoate, among other aromatics (Fig. 1). In comparison to homogentisate 1,2-dioxygenases, gentisate 1,2-dioxygenases show a narrow distribution in bacterial families of proteobacteria being identified only in six α-, three β-, and three γ-proteobacterial families and being absent from δ- and ε-proteobacteria. The number of members with putative gentisate 1,2-dioxygenase genes inside the 12 proteobacterial families owing this enzyme is also significantly lower than the percentage

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

15

of homogentisate 1,2-dioxygenase carrying members. Inside the Comamonadaceae however, six out of eight members harbor a gentisate 1,2-dioxygenase, but only one a homogentisate dioxygenase. Similarly, homogentisate dioxygenases are absent from the genomes of Enterobacteriaceae, although Salmonella, Serratia, and some E. coli strains are endowed with a gentisate dioxygenase. In addition to Proteobacteria, gentisate 1,2-dioxygenase genes can be found in Corynebacteriaceae, Micrococcaceae, Mycobacteriaceae, Nocardiaceae, and Bacillaceae. 3-Hydroxyanthranilate 3,4-dioxygenase catalyzes the conversion of 3-hydroxyanthranilate to 2-amino-3-carboxymuconic semialdehyde during tryptophan degradation via the kynurenine pathway. This extradiol dioxygenase is the cupin-type dioxygenase with the narrowest distribution since it is only found and with a low representativity in Brucellaceae, Rhodobacteraceae, Sphingomonadaceae, Burkholderiaceae, Shewanellaceae, Xanthomonadaceae, and Myxococcaceae in Proteobacteria and in Flavobacteriaceae, Flexibacteraceae, and Bacillaceae in non-proteobacterial families.

4.5

Other Extradiol Dioxygenases

Recently, a novel Fe 2+-dependent dioxygenase, benzoquinol 1,2-dioxygenase, which is a α 2β 2 heterotetramer where the α- and β-subunits displayed no significant sequence identity with other dioxygenases and which catalyzes the ring fission of a wide range of benzoquinols to the corresponding 4-hydroxymuconic semialdehydes, has been described in P. fluorescens ACB (Moonen et al. 2008). Putative genes encoding both subunits of benzoquinol 1,2-dioxygenase show a highly narrow distribution since they are almost exclusively found in Burkholderia with the exceptions of P. luminescens subsp. laumondii TTO1 and P. aeruginosa PA7 strains, in spite to be originally identified in a 4-hydroxyacetophenone-degrading P. fluorescens strain (Moonen et al. 2008). The origin of this type of dioxygenase remains to be clarified.

4.6

Diiron Oxygenases

Soluble diiron oxygenases comprise an evolutionary-related family of enzymes capable to monooxygenate benzene/toluene to phenol/methylphenol and phenols to catechols (Leahy et al. 2003). Sequence comparisons of the respective α-subunits with the PaaA oxygenase subunit of phenylacetyl-CoA oxygenase and the BoxB oxygenase of benzoyl-CoA oxygenase strongly suggest that also these enzymes belong to the family of soluble diiron oxygenases. Benzene/toluene monooxygenases and phenol monooxygenases of the soluble diiron oxygenase family are enzyme complexes including an electron transport system comprising a reductase (and, in some cases, a ferredoxin), a catalytic effector and a terminal heteromultimeric oxygenase composed by α, β, and γ subunits whose α-subunits are assumed to be the site of substrate hydroxylation (Leahy et al. 2003).

16

D. Pérez-Pantoja et al.

According to the presence of genes putatively coding for α subunit, benzene/toluene multicomponent monooxygenase are found almost exclusively in β-Proteobacteria, including Burkholderia, Cupriavidus, Ralstonia, Methylibium, and Dechloromonas strains with the only exceptions of Bradyrhizobium sp. BTAi1 and Frankia sp. CcI3. In the β-proteobacterial strains, the benzene/toluene multicomponent monooxygenase are associated with a phenol/methylphenol multicomponent monooxygenase. On the other hand, the phenol/methylphenol multicomponent monooxygenases showed a slightly broader distribution since in addition to the above mentioned strains, such genes are also identified in Acidovorax and Verminephrobacter strains and even in γ-proteobacterial families such as Alteromonadaceae and Pseudomonadaceae. In contrast to the limited distribution of the above described multicomponent monooxygenases, multicomponent phenylacetyl-CoA oxygenases are broadly distributed in Proteobacteria being identified in 6 out of 19 α-, 5 out of 10 β-, and 8 out of 29 γ-proteobacterial families. They are, however, absent from δ- and ε-proteobacteria. The families Rhodobacteraceae, Bradyrhizobiaceae, Alcaligenaceae, Burkholderiaceae, Rhodocyclaceae, Enterobacteriaceae, and Pseudomonadaceae include a significant number of strains with such genes. Several representatives are also found in non-proteobacterial families, predominantly Actinobacteria such as Streptomycetaceae, Pseudonocardiaceae, Nocardiaceae, Micrococcaceae, Corynebacteriaceae, Brevibacteriaceae, and Acidothermaceae, and also in Flavobacteriaceae and Bacillaceae families. Benzoyl-CoA oxygenase encoding genes are exclusively found in some families of the α- and β-proteobacteria: Bradyrhizobiaceae, Rhodospirillaceae, Comamonadaceae, Burkholderiaceae, and Rhodocyclaceae, and predominantly in the last two families in which the pathway was also originally described (Denef et al. 2004; Zaar et al. 2004).

4.7

Flavoprotein Monooxygenases

Flavoprotein monooxygenases are involved in a wide variety of biological processes including biosynthesis of antibiotics and siderophores or biodegradation of aromatics. They have been classified according to sequence and structural data in six classes (van Berkel et al. 2006), with classes A, D, and F being of special importance for aromatic degradation. Class A enzymes are considered to be widely distributed in different bacterial taxa and typically ortho- or para-hydroxylate aromatic compounds that contain an activating hydroxyl- or amino-group (van Berkel et al. 2006). In fact, it is interesting to note that according to genome annotations, a huge set of bacteria contain enzymes capable of 4-hydroxybenzoate 3-hydroxylation, salicylate 1-hydroxylation or 2,4-dichlorophenol 6-hydroxylation. Regarding the fact that the capability to mineralize chloroaromatics is not widespread in bacteria and chlorocatechol genes, usually necessary to achieve mineralization of chloroaromatics are, among the sequenced genomes only observed in the two bacteria well studied for such capability, i.e., B. xenovorans LB400 (Chain et al.

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

17

2006) and C. necator JMP134 (Pérez-Pantoja et al. 2008), the annotated widespread of enzymes involved in dichlorophenol degradation is astonishing. A phylogenetic analysis of proteins related to enzymes of class A flavoproteins using proteins of documented function (salicylate 1-hydroxylases, 3-hydroxybenzoate 4-hydroxylases, 2-aminobenzoyl-CoA monooxygenases/reductases, 4-hydroxybenzoate 3-hydroxylases, among others) as seeds show that these oxygenases can be grouped into six distinct protein clusters (enzymes related to UbiH involved in ubiquinone biosynthesis will not be discussed here). Only one of these clusters comprises enzymes, which, based on characterized representatives, can be assumed to catalyze a single defined activity, i.e., the 3-hydroxylation of 4-hydroxybenzoate. As with the majority of aromatic degradative properties, the respective enzymes are predominantly observed in Actinobacteria and Proteobacteria. However, they are also observed in one of two Acidobacteria, in Pedobacter of the Bacteroidetes, in one Deinococcus and in 1 of 28 Bacillaceae. No other monocomponent flavoprotein monooxygenases discussed in this section are observed in these orders. Among the Actinobacteria, 4-hydroxybenzoate 3-hydroxylases are observed in roughly one third of the families, including Arthrobacter and Streptomyces, but interestingly were absent from any of the 17 Mycobacterium analyzed. It is a dominant trait in α-Proteobacteria, specifically in Bradyrhizobiaceae and Rhodobacteraceae. Also among β-Proteobacteria, all 34 Burkholderia, three Cupriavidus, four Ralstonia, and six out of eight Comamonadaceae are endowed with such capability. In contrast, such activity is rare in γ-Proteobacteria with the exception of Pseudomonadaceae, where 17 out of 18 strains (exception again P. mendocina) have a 4-hydroxybenzoate 3-hydroxylase. Similarly, such activity is spread among Acinetobacter and Xanthomonas strains. Among the Enterobacteriaceae, only Klebsiella pneumoniae and Serratia proteomaculans have a 4-hydroxybenzoate 3-hydroxylase. Also the aminobenzoyl-CoA pathway (Altenschmidt and Fuchs 1992) seems to be strongly represented among the thus far sequenced bacteria. In a phylogenetic analysis, the aminobenzoyl-CoA oxygenases seem to be related to salicylyl-CoA 5-hydroxylase from Streptomyces sp. WA46 (Ishiyama et al. 2004) channeling salicylate to gentisate. However, in contrast to the organization in strain WA46 where the oxygenase encoding gene is clustered with a gentisate dioxygenase, function as a salicylyl-CoA 5-hydroxylase can be suggested only in a few cases, such as in S. wittichii RW1, since a gentisate pathway is absent from the genomes of various strains including the two Streptomyces strains sequenced. Overall, homologues to aminobenzoyl-CoA oxygenases are observed in 44 genomes comprising Actinobacteria (five genomes) such as Streptomyces or Saccharopolyspora erythraea NRRL 2338. In Proteobacteria this pathway is absent in γ-Proteobacteria, but it is observed in Plesiocystis pacifica SIR-1 (a δ-proteobacterium). The pathway is abundant in β-Proteobacteria such as Azoarcus strains, where this metabolic route was initially established (Altenschmidt and Fuchs 1992), but also in Comamonadaceae (six of eight genomes), Ralstonia (all four genomes), Cupriavidus (all three genomes), and α-proteobacteria such as

18

D. Pérez-Pantoja et al.

Bradyrhizobium strains (all three genomes) or Rhodobacteraceae (11 of 24 genomes). A large number of genes in bacterial genomes (nearly 100) are annotated as encoding salicylate 1-hydroxylases. However, a phylogenetic analysis taking into account validated salicylate 1-hydroxylases, identified only two of such proteins (amino acid sequence identity >40% to validated NahG proteins [Yen and Gunsalus 1982]) encoded in the genome of A. baylyi ADP1 (as previously described [Jones et al. 2000]) and P. putida GB-1 (see Fig. 5, cluster 1). Also enzymes related to NahW, a second evolutionary lineage of salicylate 1-hydroxylases (Bosch et al. 1999b) are scarce and only seven homologues (four of them encoded by Burkholderia genomes) are identified (sequence identity >35%) (see Fig. 5, cluster 5). In contrast, various enzymes (observed in 22 genomes) clustered with enzymes of proven function as 3-hydroxybenzoate 6-hydroxylases (Fig. 5, cluster 10) and were observed, among others, in three Corynebacteria, two Arthrobacter, seven Burkholderiaceae, and three Comamonadaceae strains. Other enzymes annotated as salicylate hydroxylases (16) show high similarity (>60% identity) and cluster together with 6-hydroxynicotinate 3-monooxygenase of P. fluorescens TN5 (Nakano et al. 1999) such that their function as salicylate hydroxylases is questionable (Fig. 5, cluster 2). The same holds true for a further more than 100 additional sequences, out of which 69 (Fig. 5, cluster 6–9) are, among enzymes with validated function, phylogenetically most closely related to 3-hydroxybenzoate 6-hydroxylases. However their genomic contexts indicate different functions. A similar situation holds for enzymes annotated as 3-hydroxyphenylpropionate monooxygenases. An overall of 24 proteins showed significant similarity (>40% identity) with respective validated enzymes and, in phylogenetic analysis, clustered together in one evolutionary branch. These enzymes are predominantly observed in Mycobacterium (seven genomes) and Enterobacteriaceae (mainly E. coli, 11 genomes, but also in K. pneumoniae and S. sonnei), as well as in B. vietnamiensis, B. xenovorans, C. necator JMP134, and P. putida W619. Other enzymes annotated as 3-hydroxyphenylpropionate monooxygenases show significant similarity to either resorcinol monooxygenase of C. glutamicum (Huang et al. 2006) or to GdmM involved in formation of the geldanamycin benzoquinoid system by S. hygroscopicus AM 3672 (Rascher et al. 2005) and are thus highly improbable to function as 3-hydroxyphenylpropionate monooxygenase. A 3-hydroxyphenylacetate 6-hydroxylase forming homogentisate has been recently described in P. putida U being composed of the hydroxylase and a small coupling protein, constituting a novel type of two-component hydroxylase, distinct from the classical two-component flavoprotein monooxygenases (Arias-Barrau et al. 2005). Seventeen homologues (>40% sequence identity, clustering on the same phylogenetic branch) are observed in 16 of the so far sequenced genomes and usually two subsequent genes encoding for the coupling protein and the monooxygenase can be identified. Interestingly, in contrast to the first and thus far only observation in Pseudomonas, such genes are absent from all 17 sequenced Pseudomonas strains and all other γ-proteobacterial genomes but frequently found in

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

19

Fig. 5 Evolutionary relationships among proteins related to NahG, or NahW-type salicylate 1-hydroxylases and 3-hydroxybenzoate 6-hydroxylases. Clusters 1 and 5 comprise salicylate 1-hydroxylases related to NahG or NahW salicylate 1-hydroxylases, cluster 10 3-hydroxybenzoate 6-hydroxylases, and cluster 2 enzymes related to 6-hydroxynicotinate 3-monooxygenase of Pseudomonas fluorescens TN5. The function of enzymes of other clusters remains to be elucidated

Burkholderia (5 of 34 genomes), Cupriavidus (two of three genomes), and Comamonadaceae (four out of eight genomes). Also, various flavoprotein monooxygenases are annotated as 2,4-dichlorophenol hydroxylases. However, enzymes related to valid 2,4-dichlorophenol hydroxylases (>40% sequence identity) also comprise phenol hydroxylases such as PheA from Pseudomonas sp. strain EST1001, which transforms phenol and 3-methylphenol, but not 2,4-dichlorophenol (Nurk et al. 1991), ChqA chlorobenzoquinol monooxygenase of Pimelobacter simplex (AY822041), HpbA 2-hydroxybiphenyl-3monooxygenase from P. azelaica HBP1, which is capable of oxidizing various 2-substituted phenols, but not phenol (Suske et al. 1997), OhpB 3-(2-hydroxyphenyl)propionic acid monooxygenase from R. aetherivorans I24 (DQ677338) and MhqA methylbenzoquinol monooxygenase from Burkholderia NF100 (Tago et al. 2005). Thus, enzymes of this group typically share the capability to transform 2-substituted phenols, but are obviously recruited for different metabolic routes and involve pathways where the ring-cleavage substrate is a dihydroxylated compound, but also routes where the ring-cleavage substrate is trihydroxylated. The function of these proteins, therefore, cannot be deduced from

20

D. Pérez-Pantoja et al.

similarity measures or from phylogenetic analysis. An overall of 18 proteins can be identified as belonging to this cluster, and beside the two characterized 2,4-dichlorophenol hydroxylases from C. necator JMP134 only two genomes (Rhizobium leguminosarum and Bradyrhizobium sp. ORS278) comprise proteins clustering with 2,4-dichlorophenol hydroxylases. However, the genetic environment of the encoding genes does not give a direct support for such a function. Further proteins of this cluster are observed to be scattered among Actinobacteria and Proteobacteria with R. jostii RHA1 encoding for three of such proteins. Interestingly, a distinct group of flavoprotein monooxygenases exhibiting approximately 30% of sequence identity to the above described monooxygenases is also typically annotated as phenol hydroxylases. This annotation seems to be due to some similarity to the phenol hydroxylase (30–35% identity) of Trichosporon cutaneum (Enroth et al. 1994), however, phylogenetic analysis shows that a set of 29 proteins (typically with identities >50%) is most closely related to proteins of validated function as 3-hydroxybenzoate 4-hydroxylases, previously assumed to be restricted to Comamonas strains (Hiromoto et al. 2006). In fact, inside the β-proteobacteria such genes are only observed in C. testosteroni and B. phymatum, however, also three γ-Proteobacteria harbor such gene, and 3-hydroxybenzoate-4-hydroxylases seem to be frequently encoded in the genome of α-Proteobacteria (12 genomes), specifically in Bradyrhizobium strains (all three genomes) and Rhodobacteraceae (6 out of 24 genomes). Also seven Actinobacteria seem to harbor such activity (among them two Corynebacterium species and both sequenced Arthrobacter strains), indicating this activity to be more widespread than previously thought. Nearly 20 enzymes were annotated as pentachlorophenol monooxygenases, an activity previously reported, for example, in Sphingobium chlorophenolicum (Cai and Xun 2002). However, none of these proteins showed sequence identities >35% to validated PcpB proteins, and only a group of enzymes typically encoded in Burkholderia genomes could be shown to be evolutionary related, however, their function as PCP monooxygenases seems highly improbable. Styrene monooxygenases (StyA) have been identified in various Pseudomonas strains (Beltrametti et al. 1997), and were classified as Class E flavoprotein monooxygenases, however, they are evolutionary related to the Class A flavoprotein monooxygenases (van Berkel et al. 2006). Interestingly, none of the sequenced Pseudomonas strains harbor such a gene. Eight phylogenetically related proteins are observed in genome sequencing projects, however their function as such monooxygenases remains speculative. Two-component aromatic hydroxylases such as 4-hydroxyphenylacetate 3-hydroxylases from E. coli (Diaz et al. 2001) consisting of an oxidoreductase and an oxygenase were classified as type D flavoprotein monooxygenases (Ballou et al. 2005) and have no structural or sequence similarities to the single-component enzymes described above. Iterative Psi-blast searches identified nearly 100 of such enzymes putatively involved in aromatic metabolism to be encoded in sequenced genomes and phylogenetic analysis indicated the presence of eight evolutionary lines (see Fig. 6).

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

21

Two of the branches contain the proteobacterial (Fig. 6, cluster 1) and non-proteobacterial (Fig. 6, cluster 7) 4-hydroxyphenylacetate 3-hydroxylases with an identity of members of the different cluster of approximately 30%. Proteins located on the same phylogenetic branch as validated 4-hydroxyphenylacetate 3-hydroxylases from Thermus or Geobacillus (Hawumba et al. 2007; Kim et al. 2007) are observed in only three Actinobacteria, but in both sequenced Deinococci and in both Thermus strains. It is also a dominant trait in Bacillaceae (13 out of 28 genomes). Among the Proteobacteria, 4-hydroxyphenylacetate 3-hydroxylation by enzymes of this cluster is a trait nearly exclusively observed in γ-proteobacteria, predominantly in Enterobacteriaceae (19 out of 61 genomes) and Pseudomonas (5 out of 18 genomes), and outside of this group only in two α-proteobacteria. The cluster of proteins most closely related to these proteobacterial 4-hydroxyphenylacetate 3-hydroxylases (50–60% identity) comprises those with high similarity to phenol hydroxylase PheA of Geobacillus thermoleovorans (Duffner and Muller 1998), R. erythropolis (CAJ01325), 4-nitrophenol hydroxylase of Rhodococcus sp. PN1 (Takeo et al. 2003), an enzyme which also acts as a phenol hydroxylase, and 4-coumarate 3-hydroxylase of Saccarothrix espanaensis involved in the formation of caffeic acid (Takeo et al. 2003) (see Fig. 6, cluster 2). Interestingly, respective genes are practically absent from proteobacteria and only observed in Photorhabdus and Saggitula, but observed in one of the two Thermus strains sequenced, in all Chloroflexaceae and in some Actinobacteria such as R. jostii RHA1, which harbors four homologues. A further group of proteins show similarity to PvcC, previously assumed to be involved in pyoverdin synthesis, but recently shown to be involved in the formation of pseudoverdine and paerucumarin by P. aeruginosa (Takeo et al. 2003) (Fig. 6, cluster 3). Interestingly, respective genes and gene clusters are exclusively observed in P. aeruginosa, B. mallei, B. pseudomallei, and B. thailandensis. A further cluster of six proteins, also typically annotated as 4-hydroxyphenylacetate 3-hydroxylases is related to TcpA 2,4,6-trichlorophenol monooxygenases of C. necator JMP134 (Sanchez and Gonzalez 2007), however, the function of these proteins also remains to be elucidated (Fig. 6, cluster 8). A different type of two-component aromatic hydroxylases consisting also of a reductase and an oxygenase has been described recently (Thotsaporn et al. 2004). This type has been also classified as type D flavoprotein monooxygenases (Ballou et al. 2005) but it is able to use FMN, FAD, and riboflavin for hydroxylation in contrast to HpaB, PheA, and TcpA, which specifically uses only reduced FAD (Thotsaporn et al. 2004). The best studied representative of this group is 4-hydroxyphenylacetate 3-hydroxylase from A. baumannii but it shows very low identity with the 4-hydroxyphenylacetate 3-hydroxylases described previously in E. coli, P. aeruginosa, or T. thermophilum (Thotsaporn et al. 2004). Although the different types of 4-hydroxyphenylacetate 3-hydroxylase catalyze the same reaction, they have significant differences in the details of the mechanisms involved (Ballou et al. 2005). Genes putatively coding for enzymes similar to the A. baumannii-type of 4-hydroxyphenylacetate 3-hydroxylase are found in some strains of α- and

22

D. Pérez-Pantoja et al.

Fig. 6 Evolutionary relationships among the large subunits of two-component flavoprotein monooxygenases related to 4-hydroxyphenylacetate 3-hydroxylase from Escherichia coli. Clusters 1 and 7 comprise 4-hydroxyphenylacetate 3-hydroxylases of proteobacteria and non-proteobacteria, cluster 2 proteins related with PheA phenol hydroxylase of Geobacillus thermoleovorans, and cluster 3 proteins with similarity to PvcC of P. aeruginosa (Takeo et al. 2003) (Fig. 6, cluster 3). The function of enzymes of other clusters remains to be elucidated

γ-proteobacteria: S. stellata, R. sphaeroides, Marinomonas sp., V. shilonii, V. vulnificus, A.vinelandii, P. entomophila, and one P. putida strain. Additional enzymes of this kind of two-component aromatic hydroxylases includes naphthoate 2-hydroxylase (NmoAB) described in Burkholderia sp. JT1500 (Deng et al. 2007) with homologous genes in some Bradyrhizobium and Cupriavidus strains and resorcinol hydroxylase from Rhizobium sp. MTP-10005 (GraAD) (Yoshida et al. 2007) with homologous genes in the related strains A. tumefaciens and R. leguminosarum and in the β-proteobacterium Polaromonas sp. JS666.

4.8

Rieske Non-heme Iron Oxygenases

The so-called Rieske non-heme iron oxygenases are one of the key families of enzymes important for aerobic activation and thus degradation of aromatics such as benzoate, benzene, toluene, phthalate, naphthalene, or biphenyl (Fig. 1) (Gibson and Parales 2000; Duarte et al. 2014). Members of this family also catalyze monooxygenations, such as salicylate 1- or salicylate 5-hydroxylases or demethylations,

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

23

such as vanillate O-demethylases. They are multicomponent enzyme complexes consisting of a terminal oxygenase component (iron-sulfur protein [ISP]) and electron transport proteins (a ferredoxin and a reductase or a combined ferredoxinNADH-reductase). The catalytic ISPs are usually heteromultimers composed of a large α-subunit containing a Rieske-type [2Fe-2S] cluster, with a mononuclear nonheme iron oxygen activation center, and a substrate-binding site modulating substrate specificity and a small β-subunit, however, some enzymes, such as phthalate 4,5-dioxygenases contain an oxygenase composed only of α-subunits. Phylogenetic analyses of Rieske non-heme iron oxygenases show that sequences obtained in our searches can be grouped into three main divergent clusters or divisions, where only two of them comprise proteins of validated function and are thus discussed here. One of these two divisions comprises the so-called phthalate family including vanillate demethylases (Gibson and Parales 2000). Four clusters of this division contain oxygenases of proven function to dioxygenate aromatics, i.e., phthalate 4,5-dioxygenases (Nomura et al. 1992), isophthalate dioxygenase (Wang et al. 1995), phenoxybenzoate dioxygenase (Dehmel et al. 1995), and carbazol dioxygenase (Sato et al. 1997b). Genes putatively encoding phthalate 4,5-dioxygenases are nearly exclusively observed in β-proteobacteria (seven genomes) except for an amazing five homologues possibly encoded in the genome of Rhodobacterales bacterium HTCC2654. Similarly, genes putatively encoding isophthalate dioxygenases are predominantly observed in β-proteobacterial genomes (overall in five), but also in one γ-proteobacterium and in two α-proteobacteria, among them strain HTCC2654. A similar spread is observed for enzymes related to phenoxybenzoate dioxygenase (observed in seven β-, four α-, and one γ-Proteobacterium). Genes putatively encoding carbazol dioxygenases are not observed in any sequenced genome. Most of the currently characterized Rieske non-heme iron oxygenases are concentrated in a well-defined division (see Fig. 7). The significant amount of validly described enzymes allows assignment of putative functions to most of the respective enzymes encoded in sequenced genomes. Benzoate dioxygenases (cluster A1) are most widely distributed and can be observed in the genomes of Actinobacteria as well as α-, β-, and γ-proteobacteria. Most importantly, such enzymes are observed in 32 out of 34 Burkholderia strains, 14 out of 18 Pseudomonas strains, and 4 out of 17 Mycobacteria. Anthranilate can be transformed either by two-component anthranilate dioxygenases such as the one described from Acinetobacter baylyi ADP1 (Eby et al. 2001) (cluster A2) or by three-component anthranilate dioxygenase as the one from Burkholderia cepacia DBO1 (Chang et al. 2003) (cluster E5). Genome analysis clearly showed that two-component dioxygenases are obviously restricted to γ-proteobacteria and are only observed in seven Pseudomonas genomes and, as described, in A. baylyi. In contrast, three-component anthranilate dioxygenases are exclusively observed in Burkholderia genomes and present in 31 out of 34 sequenced strains. Cluster A3 comprises proteins phylogenetically related with known p-cumate dioxygenases. These sequence relatives are found in five of the sequenced Pseudomonas genomes but also in S. wittichi RW1 and B. xenovorans LB400.

24

D. Pérez-Pantoja et al.

Fig. 7 Evolutionary relationships among the α-subunits of Rieske non-heme iron oxygenases excluding phthalate family enzymes. A function can be assigned to proteins of some of the clusters shown as follows: cluster A1, benzoate dioxygenases; cluster A2, two component anthranilate dioxygenases; cluster A3, proteins related with p-cumate dioxygenases; cluster B3, aniline dioxygenases; cluster C1, NidA-type dioxygenases; cluster C2, phthalate 3,4-dioxygenases; cluster C3, proteins related with diterpenoid dioxygenases; cluster C5, NahA-type naphthalene dioxygenases; cluster 6, proteins related with ethylbenzene dioxygenase from R. jostii RHA1; cluster C8, 3-phenylpropionate dioxygenases; cluster C9, benzene/toluene/isopropylbenzene/ biphenyl dioxygenases; cluster E1, salicylate 5-hydroxylases; cluster E2, 2-chlorobenzoate dioxygenases; cluster E3, terephthalate dioxygenases; cluster E4, salicylate 1-hydroxylases; and cluster E5, three component anthranilate dioxygenases. The function of enzymes of other clusters remains to be elucidated

Cluster B3 comprises proteins similar to aniline dioxygenases, and similar sequences are found only in Nocardioides sp. JS614 and Bradyrhizobium sp. BTAi1, indicating a very restricted distribution of such activity. Further related sequences, where no specific function can be postulated (clusters B1, B2, and B4) were predominantly observed in Burkholderiaceae. Proteins of cluster C1 exhibit similarity to proteins involved in the degradation of polycylic aromatics by Actinobacteria, exemplified by NidA of M. vanbaalenii PYR-1 (Stingley et al. 2004a) and thus putatively have a function in degradation of polycyclic aromatics. In accordance with this assumption, respective proteins are found to be encoded in the genomes of five environmental Mycobacteria and up to four different such proteins are observed per genome. As NidA-like proteins, also sequences putatively encoding phthalate 3,4-dioxygenases (Stingley et al. 2004b)

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

25

(cluster C2) are exclusively observed in Actinobacteria, differentiating them from β-proteobacteria which obviously degrade phthalate by phthalate 4,5-dioxygenases. Phthalate 3,4-dioxygenases were observed to be encoded in genomes of Mycobacteria comprising a NidA sequence, but also in M. avium strains, R. jostii RHA1, and Arthrobacter sp. FB24. Group C3 proteins, comprising diterpenoid dioxygenases-like proteins (Martin and Mohn 1999) are having a very restricted distribution in the genomes available so far, being found only in Caulobacter sp. K31, Sphingomonas sp. SKA58, S. wittichii RW1, and B. xenovorans LB400 genomes (Smith et al. 2007). Naphthalene and phenanthrene dioxygenases related to NahA of P. stutzeri AN10 (Bosch et al. 1999a) have previously been observed in various Pseudomonas, Sphingomonas, Burkholderia, Cycloclasticus, Acidovorax, and Ralstonia isolates. The genomic survey indicates such activities (see cluster C5) not to be widespread and similar sequences are only observed in genomes of N. aromaticivorans DSM 12444, Acidovorax sp. JS42, and P. naphthalenivorans CJ2. Also sequences related to ethylbenzene dioxygenase from strain RHA1 (Iwasaki et al. 2006) (cluster C6) are additionally observed only in of Azotobacter vinelandii AvOP and N. aromaticivorans DSM 12444. Sequences indicating to encode 3-phenylpropionate dioxygenases (cluster C8) are exclusively observed in Enterobacteriaceae, and interestingly observed in all Shigella spp. strains (seven genomes) and 11 of 17 E. coli. Cluster C9 is composed of benzene/toluene/isopropylbenzene/biphenyl dioxygenases (Witzig et al. 2006), enzymes typically involved in the degradation of the respective compounds, where a broad set of both proteobacterial and actinobacterial isolates is available. Respective sequences are only observed in the four genomes of strains previously reported to harbor such activity ( P. putida F1, B. xenovorans LB400, P. napthalenivorans CJ2, and R. jostii RHA1). Cluster E comprises enzymes acting on ortho- or para-substituted benzoates and include salicylate 5-hydroxylases (Fuenmayor et al. 1998) (cluster E1), salicylate 1-hydroxylases (Pinyakong et al. 2003) (cluster E4), 2-chlorobenzoate dioxygenases (cluster E2), three-component anthranilate dioxygenases (cluster E5, see above), and terephthalate dioxygenases (Sasoh et al. 2006) (cluster E3). Respective sequences are nearly exclusively observed in β-proteobacteria and in Sphingomonads out of the α-proteobacteria and only terephthalate dioxygenases are also observed in Actinobacteria, i.e., R. jostii RHA1 and Arthrobacter aurescens T1, which corresponds with various reports of Rhodococci being capable of degrading terephthalate. Terephthalate dioxygenases are also observed in B. xenovorans LB400 and C. testosteroni with members of last mentioned genus also often being implicated in terephthalate degradation (Sasoh et al. 2006). Salicylate 5-hydroxylases were observed in two Cupriavidus strains, both Polaromonas strains, and both R. solanacearum isolates in accordance with such activity being first described from a Ralstonia strain (Fuenmayor et al. 1998). Also S. wittichii RW1 seems to harbor such activity. In contrast, a Rieske-type salicylate 1-hydroxylase was only observed in N. aromaticivorans DSM 12444, also in accordance with the fact that such activities so far have only been described in

26

D. Pérez-Pantoja et al.

Sphingomonads. Also putative 2-chlorobenzoate 1,2-dioxygenases are rare and a putative homologue is only observed in the genome of B. xenovorans LB400.

5

Metabolism Diversity

A very exciting question can be addressed based on the phylogenomic analyses carried out here: What is the diversity of catabolic properties within phylogenetic groups? However, before answering such question, a definition about the “unit of catabolic diversity” must first be addressed. The first unit level is pathway diversity. It refers to the presence in one bacterium or bacterial group of different ways to degrade one compound (i.e., intradiol versus extradiol ring cleavage; classical aromatic ring oxidation versus a CoA-dependent pathway, etc.). This level of diversity is the thickest and provides the most powerful versatility because it allows the microorganism to choose among very different ways to metabolize the compound. The second level of “unit of diversity” is the enzymatic diversity. It refers to the same biochemical reaction or catabolic step carried out by completely different enzymes. For example, enzymes belonging to three different families can perform phenol conversion to catechol: single-component flavoprotein monoxygenases, diiron oxygenases, or two-component monooxygenases. This level of catabolic diversity is finer than the previous one, but still significant because it allows for versatility at the biochemical level, i.e., different substrate affinities, different cofactor requirements, inhibitor effects, among others. The third level of catabolic diversity is the genetic diversity, or classical gene redundancy: the same biochemical step may be performed by very similar enzymes encoded by different genes. It is assumed that the main point of diversity here is at the regulatory level. Although a gross measure of catabolic versatility, in the following three sections the pathway diversity will be used as a diversity unit for aromatic catabolism properties of a taxonomic group. This is especially relevant to account for the diversity of central pathways as defined in Table 2.

5.1

Metabolism by Bacteria Outside the Actinobacterial and Proteobacterial Phyla

When the genome database is searched for the aromatic catabolic pathways listed in Table 2, using the corresponding representative gene sequences, an unequal distribution of these markers among phyla and genera is easily noticed. Only members of 8 out of 17 phyla where representatives have been sequenced show the presence of the catabolic gene markers described above. However, it should be also noted that among the phyla showing absence of aromatic catabolic pathway markers, often only a few representatives have been sequenced, such as one Verrucomicrobia, two Aquificae, Fusobacteria, or Lentispharea strains, three Planctomycetes, six Thermotogae, or nine Spirochaetes. Specifically in case the phylum contains aerobic species, only further genome analysis will reveal if such capabilities are in fact absent.

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

27

Aromatic metabolic pathways were also absent from Chlamydiaea (11 genomes) where cultured representatives are obligate intracellular parasites of eukaryotic cells, the typically strict anaerobic Chlorobi (10 genomes), but also from Cyanobacteria (40 genomes), even though, for example, phenol degradation by the cyanobacterium Phormidium valderianum has been reported (Shashirekha et al. 1997). Most of the catabolic markers analyzed here are exclusively observed in Proteobacteria and Actinobacteria. This may be due to the fact that an immense amount of work has been invested specifically on elucidation of aromatic degradation in easy to culture members of these phyla. It thus cannot be excluded that novel groups of catabolic enzymes will be identified from other phyla. However, members of certain catabolic gene families can be observed in some representatives of other genera, such that the genome survey performed here is valid to get a reasonable overview of metabolic properties also from other phyla. For example, members of the cupin family, i.e., gentisate 1,2-dioxygenase, homogentisate 1,2-dioxygenase, and 3-hydroxyanthranilate 3,4-dioxygenase are all observed in other phyla, with homogentisate 1,2-dioxygenase being observed in Bacteroidetes, Chloroflexi, and Firmicutes (Bacilli). Bacilli and Bacteroidetes were also indicated not only to encode gentisate 1,2-dioxygenase and 3-hydroxyanthranilate 3,4-dioxygenase but also a phenylacetate degradative pathway. In contrast to ring-cleavage pathways mediated by members of the cupin family, pathways mediated by other extradiol dioxygenases or intradiol dioxygenases are scarce outside of the Actinobacterial and Proteobacterial phyla. Intradiol cleavage dioxygenases are observed in Acidobacteria and the Thermus/Deinococcus phylum, among the LigB-type extradiol dioxygenases only homoprotocatechuate 2,3-dioxygenases is observed in Bacilli and out of EXDO I proteins only homoprotocatechuate 2,3-dioxygenase is observed in Bacilli and Thermus/Deinococcus. Exceptional is the detection of distinct EXDO I proteins in Chloroflexi. Even though only two Acidobacteria and four Deinococcus/Thermus strains have been sequenced, the genomic survey indicates aromatic metabolic properties to be spread among those phyla. It can be suggested that S. usitatus Ellin6076 is capable to degrade 4-hydroxybenzoate via protocatechuate followed by intradiol cleavage and 4-hydroxyphenylpyruvate via the homogentisate pathway. Further capabilities of Acidobacteria thus remain to be discovered. All four members of the phylum Deinococcus/Thermus obviously share the capability to degrade 4-hydroxyphenylacetate via homoprotocatechuate and D. geothermalis DSM 11300 seems to harbor the capability to degrade 4-hydroxybenoate via protocatechuate and intradiol cleavage. Intradiol cleavage seems to be absent from Chloroflexi, Bacteroidetes, and Firmicutes. Interestingly, Chloroflexi can be proposed to be phenol degraders catabolizing it via catechol and meta-cleavage. Among Bacteroidetes, the homogentisate pathway and astonishingly the 3-hydroxyanthranilate pathway, in addition to the phenylacetate degradative pathway, seem to be spread among members of the orders Flavobacteriales and Sphingobacteriales. Out of the Firmicutes, only Bacillaceae (members of the genera Bacillus, Exiguobacterium, Geobacillus, and Oceanobacillus have been sequenced) seem to harbor aromatic metabolic properties. Unfortunately, no Paenibacillus genome sequence is available so far. Bacillus strains

3-Hydroxybenzoate 4-hydroxylase

Enzyme group Protocatechuate 3,4-dioxygenase Catechol 1,2-dioxygenase Hydroxybenzoquinol 1,2-dioxygenase Chlorocatechol 1,2-dioxygenase Catechol 2,3-dioxygenase 2,3-Dihydroxybiphenyl 1,2-dioxygenase Homoprotocatechuate 2,3-dioxygenase Protocatechuate 4,5-dioxygenase Gallate 4,5-dioxygenase Homoprotocatechuate 2,3-dioxygenase 2,3-Dihydroxyphenylpropionate 1,2-dioxygenase 2-Aminophenol 1,6-dioxygenase Gentisate 1,2-dioxygenase Homogentisate 1,2-dioxygenase 3-Hydroxyanthranilate 3,4-dioxygenase Benzoquinol 1,2-dioxygenase Benzoyl-CoA oxygenase Phenylacetyl-CoA oxygenase 2-Aminobenzoyl-CoA monooxygenase/ reductase 4-Hydroxybenzoate 3-hydroxylase LigB-type dioxygenase Cupin superfamily dioxygenase Cupin superfamily dioxygenase Cupin superfamily dioxygenase Type IV extradiol dioxygenase Soluble diiron oxygenase Soluble diiron oxygenase Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase

Family Intradiol dioxygenase Intradiol dioxygenase Intradiol dioxygenase Intradiol dioxygenase Type I extradiol dioxygenase Type I extradiol dioxygenase Type I extradiol dioxygenase LigB-type dioxygenase LigB-type dioxygenase LigB-type dioxygenase LigB-type dioxygenase + ++ ++ ++ + ++ ++ ++

Pathway marker ++ ++ ++ + ++ # ++ ++ ++ ++ ++

Table 2 Key groups of catabolic enzymes discussed in the metabolic diversity section

Mhb4H

Phb3H

Forming protocatechuate Forming protocatechuate

Amn Gen Hge Han Bqu Box Paa Abc

Abbreviation Pca34 Cat12 Hqu Cca Cat23 Dhb HpcEXDOI Pca45 Gal HpcLigB Dhp

Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Dearomatization Dearomatization Dearomatization

Enzyme function Intradiol cleavage Intradiol cleavage Intradiol cleavage Intradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage Extradiol cleavage

28 D. Pérez-Pantoja et al.

Terephthalate 1,2-dioxygenase Phthalate 3,4-dioxygenase Anthranilate 1,2-dioxygenase (2 component) Anthranilate 1,2-dioxygenase (3 component) Benzoate 1,2-dioxygenase Salicylate 5-hydroxylase Phenylpropionate 2,3-dioxygenase

Resorcinol 4-hydroxylase

4-Hydroxyphenylacetate 3-hydroxylases

Chlorophenol 4-hydroxylase

Phenol 2-hydroxylase

4-Hydroxyphenylacetate 3-hydroxylases

Phenol/benzoquinol hydroxylase

3-Hydroxyphenylpropionate 2-hydroxylase 3-Hydroxyphenylacetate 6-hydroxylase

3-Hydroxybenzoate 6-hydroxylase

Salicylate 1-hydroxylase

Channeling to protocatechuate Channeling to protocatechuate Channeling to catechol Channeling to catechol Channeling to catechol Channeling to gentisate Channeling to 2,3-dihydroxyphenylpropionate

Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase

Forming hydroxybenzoquinol

Forming homoprotocatechuate

Forming chlorobenzoquinol

Forming catechol

Forming homoprotocatechuate

Forming catechol/hydroxybenzoquinol

Forming 2,3-dihydroxyphenylpropionate Forming homogentisate

Forming gentisate

Forming catechol

Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase Class A flavoprotein monooxygenase Class D flavoprotein monooxygenase Class D flavoprotein monooxygenase Class D flavoprotein monooxygenase Class D* flavoprotein monooxygenase Class D* flavoprotein monooxygenase Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase

(continued)

BenDO Sal5H PhpDO

AntDO

TphDO Pht34DO AntDO

Res4H

Pha3H

Ph4H

Ph2H

Pha3H

Pbq2H

Mha6H

Mhp2H

Mhb6H

Ohb1H

Phylogenomics of Aerobic Bacterial Degradation of Aromatics 29

Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase Soluble diiron oxygenase Soluble diiron oxygenase

Family Rieske nonheme iron oxygenase Rieske nonheme iron oxygenase

Pathway marker

Channeling to protocatechuate Channeling to protocatechuate Forming phenol Forming catechol

Enzyme function Activation of hydrophobic aromatics Polycyclic aromatic degradation PhtDO IphDO Tmo Pmo

Abbreviation BphDO NidDO

Only enzymes where a function could be assigned with high probability are included in the list Eleven groups of aromatic ring-cleavage activities (homoprotocatechuate 2,3-dioxygenases, even though belonging to different enzyme families were defined as one activity) and all groups of enzymes catalyzing dearomatization of aromatic CoA derivatives were defined as abundant, as they are observed in more than ten sequenced genomes and are marked as ++ Three groups of aromatic ring-cleavage activities were defined as less abundant, as they were observed in ten or less sequenced genomes and are marked as + 2,3-Dihydroxybiphenyl 1,2-dioxygenases (marked #) are not included in the list of aromatic catabolic pathway markers discussed in the proteobacterial section, as they are assumed to have their function in the metabolism of bi- and polycyclic aromatics rather than monocyclic aromatics Class D* flavoprotein monooxygenases refers to enzymes capable of using FMN, FAD, and riboflavin for hydroxylation

Enzyme group Biphenyl 2,3-dioxygenase type Naphthalene inducible dioxygenase (NidA) type Phthalate dioxygenase 4,5-dioxygenase Isophthalate dioxygenase Toluene/benzene monooxygenase Phenol monooxygenase

Table 2 (continued)

30 D. Pérez-Pantoja et al.

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

31

such as Bacillus sp. JF8 (Shimura et al. 1999), B. subtilis IS13 (Shimura et al. 1999), and others have been shown to be capable of degrading aromatics such as biphenyl, guaiacol, cinnamate, coumarate, or ferulate (Peng et al. 2003), and Paenibacilli such as P. naphthalenovorans, Paenibacillus sp. strain YK5, or Paenibacillus sp. KBC101 (Daane et al. 2002; Iida et al. 2006; Sakai et al. 2005) are shown to be capable of degrading naphthalene, dibenzofuran, or biphenyl. Thus, the metabolic diversity of Bacillaceae is clearly underrepresented by the currently sequenced 28 genomes, which indicate metabolic properties similar to those of Bacteroidetes, such as a spread of the homogentisate pathway in Bacillus and the presence of the 3-hydroxyanthranilate and the gentisate pathway in addition to the phenylacetate degradative pathway in members of different genera. In addition, 4-hydroxyphenylacetate degradation via homoprotocatechuate seems to be also a capability spread among Bacillaceae.

5.2

Actinobacteria

Aromatic metabolic routes can be observed in 12 out of 20 families from the phylum Actinobacteria and pathways analyzed here are absent in Actinomycetaceae, Cellulomonadaceae, Kineosporiaceae, Microbacteriaceae, Nocardiopsaceae, Propionibacteriaceae, Bifidobacteriaceae, and Coriobacteriaceae. Within the Corynebacterium genus, C. diphteriae and C. jeijekum, a nocosomial pathogen have no aromatic catabolic pathways. Interestingly, they have the smaller genomes of this group. A similar situation is observed within the Mycobacteria, as M. leprae, M. bovis, and M. tuberculosum also have no aromatic catabolic pathways and the smaller genomes of this group. In contrast, environmental Mycobacteria are characterized by an enormous metabolic potential, however, it should be noted that M. vanbaalenii Pyr1, M. gilvum PYR-GCK, as well as strains JLS, KMS, and MCS have been sequenced due to their capability to degrade various polycyclic aromatics reflected in the presence of up to four NidA-type Rieske non-heme iron oxygenases for initiating metabolism of PAHs and up to six BphC type I extradiol dioxygenases per genome. However, not only Mycobacteria are endowed with a high metabolic potential. In contrast to members of all phyla described above, Actinobacteria not only often comprise a homogentisate pathway, which is observed in seven families, but also a protocatechuate intradiol cleavage pathway observed in eight families and more than one third of sequenced strains. Typically, actinobacterial strains endowed with a protocatechuate pathway also harbor a protocatechuate forming 4-hydroxybenzoate 3-hydroxylase such as both Micrococcaceae or Streptomycetaceae (see Table 3) and often a 3-hydroxybenzoate 4-hydroxylase (such as both Micrococcaceae), indicating protocatechuate to be a central intermediate of various metabolic routes. Interestingly, Mycobacteria harboring a protocatechuate intradiol cleavage do not contain any of the aforementioned genes, but typically a phthalate dioxygenase. Table 3 shows an overview of catabolic markers observed at least twice in genomes of actinobacterial families, from which at least two genomes have been sequenced. Two observations are evident from the table. First, Corynebacteriaceae,

32

D. Pérez-Pantoja et al.

Nocardiaceae, and specifically Micrococcaceae are endowed with a broad metabolic potential. However, it should be noted that among the three Nocardiaceae, R. jostii RHA1 has a metabolic potential much broader than Nocardia farcinica IFM 10152 or Nocardioides sp. JS614. Unfortunately, no more sequences of the reported highly versatile Rhodococcus genus (van der Geize and Dijkhuizen 2004) are available thus far. Also various reports on the metabolic versatility of Arthrobacter strains are known (Nordin et al. 2005). In contrast, Corynebacteria just recently have become the focus of more intense metabolic investigations (Huang et al. 2006). Second, the table shows a clear cooccurrence of ring-cleavage activity markers as well as of markers for peripheral activities, supporting that our annotation efforts are appropriate to deduce metabolic potential.

5.3

Proteobacteria

Three of the five classes of Proteobacteria (α, β, and γ) concentrate the vast majority of the reported catabolic pathways towards aromatic compounds that can be traced in the current genome databases (Table 4). Only a couple of aromatic catabolic pathways (Pca34, Hge, and Han) are found in some strains of the Myxococcales order of δ proteobacteria and none in the ε proteobacterial class. The α class of Proteobacteria has an uneven distribution of aromatic catabolic gene markers. None of the members of the three families of the order Rickettsiales have such catabolic properties. The small genome size of these members may be related to this trait. Aromatic ring-cleavage pathways are also absent from all members of the Parvularculaceae, Bartonellaceae, and Erythrobacteraceae families and some members of the Aurantimonadaceae, Bradyrhizobiaceae, Methylobacteriaceae, Phyllobacteraceae, Rhodobacteraceae, Acetobacteracea, Rhodospirillaceae, and Sphingomonadaceae families. In contrast, four α-proteobacterial strains (Bradyrhizobium sp. BTAi1, S. wittichii RW1, Sagittula stellata E-37, and Silicibacter pomeroyii DSS-3) have 8–9 out of the 14 main pathways and another three strains ( (Bradyrhizobium japonicum USDA110, Bradyrhizobium sp. ORS278, and Jannaschia sp. CCS1) have seven main aromatic catabolic pathways suggesting Bradyrhizobium strains to be metabolically highly versatile. The most broadly distributed pathways in the α class of proteobacteria are Pca34 and Hge being observed in 30–40% of the sequenced genomes and in 11 and 9 families, respectively. Some catabolic pathways are only seldomly found in members of this proteobacterial class, and only N. aromaticivorans DSM 12444 has the Cat23 pathway and only X. autotrophicus Py2 has the Dhp pathway. The Cca, Amn, and Bqu pathways are not found in any α proteobacterial genome. Regarding peripheral pathways, α proteobacterial strains endowed with a protocatechuate pathway also harbor a Phb3H and with lower frecuency a Mhb4H. Isomers of phthalate seems not to be typical substrates for α proteobacteria, since with the exception of IphDO in B. japonicum USDA110, phthalate, isophthalate, or terephthalate dioxygenases are not found. BenDO are usually observed in strains

+

+

+

++

+

+

+ +

++

+

+ ++

++

++

+

+

+

+

+

+

+

(+)

+

+

(+)

+

+

++

++

++

+

++

++

+

++

+

++; More than 60% of the sequenced genomes of these proteobacterial families comprise a gene encoding the mentioned activity (number of sequenced representatives is given in parentheses); +, between 20 and 60%; (+), less than 20%. For abbreviations, see Table 2. Only families where at least two members have been sequenced are included in the analysis

Bifidobacteriaceae (4)

++

+

+

++

++

+

Streptomycetaceae (2) ++

+

+

++

Nocardioidaceae (3)

+

++

+

+

+

++

+

++

+

Mycobacteriaceae (17)

++

++

++

++

++

Pca34 Phb3H Mhb4H Pht34DO TphDO Cat12 BenDO Gen Mhb6H Hge HppDO Mha6H Dhp Mhp2H Hpcexdoi Pha3H NidDO Dhb Paa Abc

Micromonosporaceae (2)

Micrococcaceae (2)

Microbacteriaceae (2)

Frankiaceae (3)

Corynebacteriaceae (5)

Cellulomonadaceae (2)

Actinobacterial families

Table 3 Catabolic gene markers of Proteobacteria

Phylogenomics of Aerobic Bacterial Degradation of Aromatics 33

(+)

+

Pht34DO

(+)

TphDO

(+)

Cat23 (+)

Ohb1H (+)

AntDO ++

(+)

PMO (+)

(+)

Ph2H (+)

(+)

+

(+)

(+)

(+)

+

(+)

+

++

++

+

++

(+)

(+)

(+)

+

(+)

+

Neisseriaceae (5)

++

++

(+)

+

(+)

(+)

Comamonadaceae (8)

Oxalobacteraceae (2)

+

(+)

++

Burkholderiaceae (43)

(+)

++

Alcaligenaceae (3)

β Proteobacterial families

(+)

+

++ +

(+) (+)

++ (+)

(+)

+

+

++

+

+

+

(+)

(+)

++

++

++

++

++

++ +

+

++

++

++

+

++

(+)

+

++

+

+

+

++

Sphingomonadaceae (5)

(+)

++

(+)

+

(+)

++

++

HppDO

Erythrobacteraceae (3)

SAR11 (2)

Rickettsiaceae (16)

Anaplasmataceae (10)

Rhodospirillaceae (3)

(+)

+

BenDO +

(+)

Pbq2H

+

++

++

+

TMO (+)

Hqu

++

++

Xanthobacteraceae (2)

++

+

IphDO

(+)

Res4H

+

++

Rhizobiaceae (5)

+

++

+

Bqu

Acetobacteraceae (3)

++

Phyllobacteriaceae (3)

++

++

Gen +

Mhb6H

Rhodobacteraceae (24)

+

Methylobacteriaceae (3)

Pca45

+

Mhb4H

+

Ohb5H +

++

Brucellaceae (6)

++

Phb3H

+

Hge

Hyphomonadaceae (3)

(+)

Bradyrhizobiaceae (11)

Bartonellaceae (3)

++

+

Aurantimonadaceae (2)

Pca34

Caulobacteraceae (2)

α Proteobacterial families

Cat12

Table 4 Catabolic gene markers of Actinobacteria Mha6H +

(+)

(+)

(+)

Han (+)

(+)

(+)

(+)

(+)

Dhp (+)

+

Mhp2H (+)

Gal (+)

Hpc LigB ++

+

(+)

++

(+)

+

Hpc EXDOI +

+

+

+

Pha3H (+)

+

Paa +

++

++

(+)

+

+

+

+

++

Box ++

(+)

++

(+)

(+)

Abc ++

+

+

(+)

+

+

+

+

+

+

34 D. Pérez-Pantoja et al.

PhpDO

+

(+)

(+)

+

(+)

(+)

(+)

+

+

+

+

(+)

+ (+)

+

+

+ +

(+)

(+) +

++

++

++

++

++

++

in the analysis

given in parentheses); +, between 20 and 60%; (+), less than 20%. For abbreviations, see Table 2. Only families where at least two members have been sequenced are included

++; More than 60% of the sequenced genomes of these Actinobacterial families comprise a gene encoding the mentioned activity (number of sequenced representatives is

++

+

++

++

Xanthomonadaceae (11)

+

++

Vibrionaceae (30)

Thiotrichaceae (2)

+

Pseudomonadaceae (19)

Francisellaceae (7)

+

++

Moraxellaceae (5)

Pasteurellaceae (21)

++ +

++

+

Oceanospirillaceae (3)

+

Legionellaceae (4)

Coxiellaceae (5)

Enterobacteriaceae (61)

(+)

+

+

+

(+)

+

+

+

++

++

(+)

(+)

+

+

(+)

+

(+)

+

(+)

+

(+)

+

+

(+)

++

(+)

+

(+)

(+)

++

+

+

++

+

+

Ectothiorhodospiraceae (3)

(+)

(+)

+

+

(+)

(+)

+

++

(+)

(+)

(+)

+

Shewanellaceae (18)

(+)

(+)

+

Psychromonadaceae (2)

(+)

+

+

++

+

(+)

+

Pseudoalteromonadaceae (3)

+

+

++

(+)

+

Idiomarinaceae (2)

Aeromonadaceae (7)

γ Proteobacterial families

Rhodocyclaceae (3)

Nitrosomonadaceae (3) ++

++

Phylogenomics of Aerobic Bacterial Degradation of Aromatics 35

36

D. Pérez-Pantoja et al.

endowed with a Cat12 pathway and strains endowed with Hge usually also harbor HppDO encoding genes. The β class of proteobacteria harbors all major central aromatic catabolic pathways listed in Table 2. The distribution of these catabolic pathways among β-proteobacterial strains has some points to be noted. Except for the presence of the Hge catabolic pathway in C. violaceum, the families Oxalobacteraceae, Neisseriaceae, and Nitrosomonadaceae are devoid of the investigated aromatic catabolic properties. Specifically, members of the Burkholderiaceae and Comamonadaceae show a high metabolic potential and usually harbor a broad set of aromatic pathways and members of the Burkholderia, Cupriavidus, Ralstonia, Delftia, and Polaromonas genera comprise up to 11 out of the 14 major central aromatic pathways (Pérez-Pantoja et al. 2008). Polynucleobacter sp. QLW-P1DMWA-1 is the only member of the Burkholderiaceae family that has no such catabolic pathway (the smallest genome among them); and Limnobacter sp. MED105 (the second smallest genome) has only Cat23. Members of the Alcaligenaceae and Rhodocyclaceae are obviously relatively limited in their aromatic catabolic potential. It should be noted that Rhodocyclaceae comprise genera such as Azoarcus, Thauera, or “Aromatoleum,” nitrate-reducing bacteria that contribute significantly to the biodegradation of aromatic compounds in anoxic waters and soils and that are endowed with several pathways for anaerobic catabolism of aromatics. It has, however, also been shown that aerobic aromatic pathway are functional in these bacteria (Rabus 2005). The most abundant pathways in the β class are Paa, Hge, Cat12, and Pca34, which are found in 60% or more of the sequenced genomes available. In contrast, Gal, Dhb, and Han are only observed in