Characterization of chlorophenol 4-monooxygenase (TftD) and NADH:FAD oxidoreductase (TftC) of burkholderia cepacia AC1100

THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 285, NO. 3, pp. 2014–2027, January 15, 2010

Characterization of Chlorophenol 4-Monooxygenase (TftD)

and NADH:FAD Oxidoreductase (TftC) of Burkholderia

□

S

cepacia AC1100^*

Received for publication, August 14, 2009, and in revised form, November 11, 2009 Published, JBC Papers in Press, November 13, 2009, DOI 10.1074/jbc.M109.056135

Brian N. Webb^‡1, Jordan W. Ballinger^§1,2, Eunjung Kim^§3, Sara M. Belchik^§, Ka-Sum Lam^§, Buhyun Youn^§4,

Mark S. Nissen^§, Luying Xun^§, and ChulHee Kang^‡§5

Fromthe^‡DepartmentofChemistryand^§SchoolofMolecularBiosciences, WashingtonStateUniversity, Pullman, Washington 99164-4660

2,4,5-Trichlorophenol (2,4,5-TCP)⁶and 2,4,6-trichlorophe-

nol (2,4,6-TCP) belong to an environmentally persistent class of

Burkholderia cepacia AC1100 completely degrades 2,4,5-tri-

chlorophenol, in which an FADH₂-dependent monooxygenase

(TftD) and an NADH:FAD oxidoreductase (TftC) catalyze the contaminants known as polychlorinated phenols (1). They

have been used extensively as pesticides and preservatives,

particularly for lumber and leather preservation, and as gen-

eral herbicides and biocides. Unfortunately, human expo-

sure likely causes various adverse health effects (2). The con-

cern has prompted governments worldwide to classify

several polychlorinated phenols as priority pollutants.

Recently, two microorganisms that degrade trichlorophenols

have been characterized. Burkholderia cepacia AC1100 miner-

alizes 2,4,5-TCP (3, 4), and Cupriavidus necator JMP134 com-

pletely degrades 2,4,6-TCP (5). 2,4,5-TCP 4-monooxygenase

(TftD) of B. cepacia AC1100, an FADH₂-dependent monooxy-

genase, oxidizes 2,4,5-TCP to 2,5-dichloro-p-benzoquinone

(2,5-DiCBQ), which is reduced to 2,5-dichloro-p-hydroqui-

none (2,5-DiCHQ) by NADH (Fig. 1A). TftD further oxidizes

2,5-DiCHQ to 5-chloro-2-hydroxy-p-benzoquinone, which is

then reduced to 5-chloro-2-hydroxy-p-hydroquinone (5-Cl-

2H-HQ) (6, 7). TftD can also oxidize 2,4,6-TCP to 2,6-DiCBQ

but cannot further transform it or its reduced form 2,6-DiCHQ.

In these reactions, TftD uses molecular oxygen and FADH₂as

co-substrates (8), and FADH₂is supplied by TftC, a flavin

reductase.

initial steps. TftD oxidizes 2,4,5-trichlorophenol (2,4,5-TCP)

to 2,5-dichloro-p-benzoquinone, which is chemically reduced

to 2,5-dichloro-p-hydroquinone (2,5-DiCHQ). Then, TftD oxi-

dizes the latter to 5-chloro-2-hydroxy-p-benzoquinone. In

those processes, TftC provides all the required FADH₂. We have

determined the crystal structures of dimeric TftC and tet-

˚

rameric TftD at 2.0 and 2.5 A resolution, respectively. The struc-

ture of TftC was similar to those of related flavin reductases. The

stacked nicotinamide:isoalloxazine rings in TftC and sequential

reaction kinetics suggest that the reduced FAD leaves TftC after

NADH oxidation. The structure of TftD was also similar to the

known structures of FADH₂-dependent monooxygenases. Its

His-289 residue in the re-side of the isoalloxazine ring is within

hydrogen bonding distance with a hydroxyl group of 2,5-Di-

CHQ. An H289A mutation resulted in the complete loss of activ-

ity toward 2,5-DiCHQ and a significant decrease in catalytic

efficiency toward 2,4,5-TCP. Thus, His-289 plays different roles

in the catalysis of 2,4,5-TCP and 2,5-DiCHQ. The results sup-

port that free FADH₂is generated by TftC, and TftD uses

FADH₂to separately transform 2,4,5-TCP and 2,5-DiCHQ.

Additional experimental data also support the diffusion of

FADH₂between TftC and TftD without direct physical interac-

tion between the two enzymes.

Several FADH₂-dependent monooxygenases, such as TftD,

have been discovered in the biodegradation pathways of various

aromatic compounds (9–12). The formation of a flavin-perox-

ide intermediate during catalysis has been proposed, and thus

protection or stabilization of the FAD-peroxide intermediate

from rapid hydrolysis seems crucial for those flavin-dependent

monooxygenases (8, 13–16). All these monooxygenases have a

small component as a partner, from which reduced flavins are

provided. Those smaller reductases have a flavin molecule

either as a tightly bound substrate or as a prosthetic group (10,

17–19). TftC and TftD belong to this two-component flavin-

diffusible monooxygenase (TC-FDM) family and share se-

* This work was supported in part by National Science Foundation Grant

MCB-0323167, the United States Department of Agriculture, Agricultural

Research Service/Cooperative State Research, Education, and Extension

Service, American Heart Association Grant 0850084Z, and The Murdock

Charitable Trust.

□S

The on-line version of this article (available at http://www.jbc.org) contains

supplemental Figs. 1 –4.

The atomic coordinates and structure factors (codes 3K86, 3K87 , 3K88, and

3HWC )havebeendepositedintheProteinDataBank, ResearchCollaboratory

for Structural Bioinformatics, Rutgers University, New Brunswick, NJ

(http://www.rcsb.org/).

⁶The abbreviations used are: 2,4,5-TCP, 2,4,5-trichlorophenol; 2,4,6-TCP,

2,4,6-trichlorophenol; TftD, chlorophenol 4-monooxygenase; TftC, NADH:

FAD oxidoreductase; 2,5-DiCBQ, 2,5-dichloro-p-benzoquinone; 2,5-DiCHQ,

2,5-dichloro-p-hydroquinone; HpaB, 4-hydroxylphenylacetate 3-mono-

oxygenase; MCAD, medium chain-specific acyl-CoA dehydrogenase;

4-BUDH, 4-hydroxybutyryl-CoA dehydratase; HPLC, high performance liq-

uid chromatography; Mops, 4-morpholinepropanesulfonic acid; TCP,

trichlorophenol; TC-FDM, two-component flavin-diffusible monooxy-

genase; MALLS, multiangle laser light scattering; ITC, isothermal titra-

tion calorimetry.

¹Both authors contributed equally to this work.

²Present address: Hollister-Stier Laboratories, Spokane, WA 99207-5788.

³Present address: The Catholic University of Korea, Daejeon St. Mary’s Hospi-

tal, Clinical Research Institute, Daejeon 300-824, Korea.

⁴Present address: Dept. of Biological Sciences, Pusan National University,

Pusan 609-735, Korea.

⁵To whom correspondence should be addressed: School of Molecular Bio-

sciences, Washington State University, Pullman, WA 99164-4660. Tel.: 509-

335-1409; Fax: 509-335-9688; E-mail: chkang@wsu.edu.

2014 JOURNAL OF BIOLOGICAL CHEMISTRY

VOLUME 285•NUMBER 3•JANUARY 15, 2010

Crystal Structures of TftC and TftD

pET30TftD in BL21(DE3) cells.

This was allowed to grow overnight

at 37 °C with constant shaking, after

which this culture was used to inoc-

ulate 1.5 liters of LB medium. In

addition, expression of selenome-

thionine-incorporated TftD in E.

coli B834(DE3) (Novagen) was

carried out using minimal media

containing 30 mg/liter selenomethi-

onine. Protein expression was

induced by addition of isopropyl

␤-D-thiogalactopyranoside to a final

concentration of 0.5 m^Mat mid-log

phase growth (A₆₀₀ϭ ϳ0.6). Fol-

FIGURE1. 2,4,5-TCPand2,4,6-TCPoxidationbyTftCandTftD. A, TftDoxidizes2,4,5-TCPto2,5-DiCBQ, which

lowing induction, the cells were

incubated at 20 °C for 12 h with

shaking at 250 rpm. Cells were har-

vested by centrifugation at 3,000 ϫ

g, and pellets were frozen to pro-

is chemically reduced to 2,5-DiCHQ. Then, TftD oxidizes the latter to 5-chloro-2-hydroxy-p-benzoquinone,

whichcanbenonenzymaticallyreducedto5-chloro-2-hydroxy-p-hydroquinone. B, TftDalsooxidizes2,4,6-TCP

to 2,6-DiCBQ but cannot further metabolize it. The latter is released and reduced to 2,6-DiCHQ. In these

oxidation processes, TftC supplies FADH₂.

quence similarity with other flavin reductases and FADH₂-de- mote lysis. The cell pellet was then thawed at room tempera-

pendent monooxygenases that are involved in the degradation

of various phenolic compounds.

ture, resuspended in a minimal volume of lysis buffer (50 m^M

Tris (pH 8.0), 300 mM NaCl, and 20 mM imidazole for TftC, 20

m^MTris (pH 8.5), and 3 mM dithiothreitol in the case of TftD),

sonicated 10 times for 10 s each using a model 450 Sonifierꢀ

(Branson Ultrasonics), and the resulting lysate cleared by cen-

trifugation (20,000 ϫ g for 40 min).

So far, little is known about the interaction between TftC and

TftD for FADH₂transfer. Particularly reduced flavins can be

readily oxidized by O₂(20), and thus flavin dynamics is one of

the critical features in the catalytic activity of monooxygenases.

So far, several plausible mechanisms have been reported, e.g. (i)

simple flavin diffusion between two components as in the cases

of 4-hydroxyphenylacetate 3-monooxygenase of Escherichia

coli and Acinetobacter baumannii (HpaB and HpaC) and phe-

nol monooxygenase of Bacillus thermoglucosidasius (Phe-1(A)

and Phe-2(A)) (21, 22); (ii) protein-protein interaction for

direct substrate channeling as in the cases of bacterial luciferase

(luciferase and flavin reductase P), alkane sulfonate monooxy-

genase (SsuE and SsuD), and arylamine oxygenase (PrnF and

PrnD) (18, 23–26); and (iii) a probable combination of diffusion

and complex formation for styrene monooxygenase (SMOA

and SMOB) (27).

For the TftC purification, lysate was applied to a nickel-ni-

trilotriacetate column and washed with several column vol-

umes of lysis buffer. Elution took place with the lysis buffer

containing 300 m^Mimidazole. Eluted fractions containing TftC

were combined, concentrated, and buffer-exchanged into 20

m^MTris (pH 8.5) containing 50 mM NaCl by ultrafiltration in an

Amicon 8050 cell with a 30-kDa cutoff membrane (Millipore).

TftC was then applied to an ion-exchange column (Bio-Rad

Uno Q12), and the protein of interest was collected in the flow-

through. TftC was then concentrated and added to several vol-

umes of 5 m^Msodium phosphate (pH 6.8). Precipitates formed

during this step were removed by centrifugation, after which

the remaining solution, containing only TftC, was concentrated

and exchanged into 20 m^MTris (pH 7.5).

In addition, the dechlorination mechanism catalyzed by TftD

is not well understood. An in-depth understanding of both TftC

and TftD is important for elucidating the catalytic mechanisms,

especially the mechanisms of sequential hydroxylations of

2,4,5-TCP. Here, we present a mechanistic analysis, including

thermodynamic properties and crystal structures of TftC and

TftD and propose a model for TCP oxidation. In addition, the

plausible mechanism of flavin transfer between TftC and TftD

was investigated through dynamic light scattering. This collec-

tive information could lead to the improvement of existing

technologies for polychlorinated phenol bioremediation (28).

For the TftD purification, lysate was loaded onto an anion-

exchange column (Toyopearlꢀ DEAE 650 ^M) equilibrated with

buffer A (flow rate 6.0 ml min^Ϫ1). TftD was eluted from the

column using a linear NaCl gradient between 0 and 200 m^M

NaCl. TftD-containing fractions were desalted and concen-

trated into buffer B (5 m^Msodium phosphate (pH 7.0)). Con-

centrated TftD was applied to a CHT-II hydroxyapatite column

(Bio-Rad), equilibrated with buffer B (flow rate 2.0 ml min^Ϫ1),

and eluted in the column flow-through. Fractions were then

concentrated into buffer C (20 m^MTris (pH 7.0), 3 mM dithio-

threitol), loaded onto a Mono Q^TMGL10/100 anion-exchange

column (GE Healthcare), and eluted with a linear NaCl gradient

at ϳ300 mM NaCl. Fractions containing TftD were pooled, con-

centrated, and exchanged into buffer D (20 m^MTris (pH 7.0)),

after which the protein was loaded onto a Sephacryl S-200 col-

EXPERIMENTAL PROCEDURES

Plasmid Construction and Cloning—Both tftC and tftD were

individually cloned into the overexpression plasmid pET30 LIC

(Novagen) as described previously (8).

Expression and Purification—TftC or TftD expression was

carried out by inoculating 100 ml of LB supplemented with 30

␮g/ml kanamycin from a freezer stock of pET30TftC or umn (GE Healthcare) and separated from remaining proteins.

JANUARY 15, 2010•VOLUME 285•NUMBER 3

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Crystal Structures of TftC and TftD

With the exception of the nickel-nitrilotriacetate column, all the substrates dissolved in the same buffer. The apo-form TftC,

purification steps were carried out using an Amersham Bio- TftD, and substrate solutions were degassed prior to titration.

sciences/BioCad 700E preparative HPLC (Applied Biosystems). The experiment consisted of 29 injections of 10 ␮l each of

All purification steps were monitored, and final homogeneity ligand into the protein solution at 25 °C with constant stirring

(Ͼ99%) was estimated using SDS-PAGE and Coomassie Blue at 300 rpm employing a 300-s equilibration interval between

staining.

injections. Heats of dilution of each ligand were determined by

Site-directed Mutagenesis and Enzyme Assay—Site-directed titration of the ligand into buffer without protein and were used

mutagenesis was performed using the QuikChange kit from to correct the protein titration data. The Origin software pack-

Stratagene (La Jolla, CA). The primers used for the conversion age (OriginLab Corp, Northampton, MA) was used to fit the

of H289A of TftD are H289AF (5Ј-CGTATCTTCGACT- data to an n-sites equivalent binding model using nonlinear

GGGTGGCTTACCACATTTTGATCCG-3Ј) and H289AR least squares regression. Fitting the data provides the affinity,

(5Ј-CGGATCAAAATGTGGTAAGCCACCCAGTCGAAG- enthalpy, and entropy change for the binding reaction.

ATACG-3Ј). The mutations were confirmed by sequencing,

Titrations of TftC with FAD, FMN, and NADH and titrations

and the correct clones were transformed into E. coli BL21(DE3) of TftD with FAD and FMN were performed under aerobic

for protein production. The mutant protein (TftD H289A) was conditions. For TftD titration with FADH₂and FMNH₂, the

purified analogous to the wild-type protein for characteriza- titration experiment was performed in a glovebox under anaer-

tion. The enzyme activity was assayed as described previously obic conditions. For anaerobic titrations, protein was diluted

(5). Briefly, 40 ␮l of assay mixture contained 20 mM potassium into buffer followed by addition of 1 m^MNADH, 1 mM glucose,

phosphate buffer (pH 7.0), 100 ␮M 2,4,6-trichlorophenol, 10 ␮M and 1 ␮g/ml catalase. The titrant was prepared under the same

FAD, 300 ␮M NADH, 1 mM ascorbic acid, 0.1 ␮g/ml of E. coli conditions but without TftD and with the addition of 100 ␮M

flavin reductase (Fre), and various amounts of wild-type TftD flavin (FAD or FMN). The samples were degassed under vac-

or H289A mutant. The reaction was initiated by adding NADH uum and then placed in the glovebox. Glucose oxidase was

to the reaction mixture and terminated by adding 40 ␮l of ace- added to 1 ␮g/ml to scavenge oxygen. The flavins were reduced

tonitrile/acetic acid mixture (9:1 (v/v)). The samples were cen- by incubation with 1 ␮g/ml of E. coli flavin reductase (Fre), and

trifuged at 13,000 ϫ g for 2 min, and the supernatants were the progress of the reaction was followed visually by the color

analyzed by HPLC (8). TftC enzyme kinetics were performed by change of the flavin.

adding FAD and NADH to the enzyme at the concentrations

Circular Dichroism—Experimental conditions for all CD

described and then monitoring the change in absorbance of measurements were 0.013 mg/ml protein in 20 m^Msodium

NADH at 340 nm. K_mand V_maxvalues were determined by phosphate buffer (pH 6.8) at 25 °C. CD spectra were recorded

fitting the data to the Michaelis-Menten equation using Origin from 200 to 300 nm using an AVIV 202SF spectrophotometer

5.0.

Molecular Mass Determination—The weight-average molec-

(AVIV Biomedical, Inc.).

Crystallization and Data Collection—Apo-form crystals of

ular masses of TftC and TftD were measured by combined size TftC were grown at 4 °C using the hanging drop vapor diffusion

exclusion chromatography and multiangle laser light scattering method. 1.5 ␮l of TftC in 20 mM Tris (pH 7.5) were mixed with

as described previously (29). Briefly, 100 ␮g of TftC (or TftD) 1.5 ␮l of reservoir solution (20% (w/v) polyethylene glycol 4000,

was loaded onto a BioSep-SEC-S 2000 column (Phenomenex) 0.2 ^Mpotassium formate) and equilibrated against the reservoir.

and eluted isocratically with a flow rate of 0.5 ml min^Ϫ1. The Crystals grew to a maximum size in 5 days and were of diffrac-

eluate was passed through tandem UV detector (Gilson), Opti- tion quality. For complex crystals of TftC with FAD and

lab DSP interferometric refractometer (Wyatt Technology), FADꢁNADH, apo-form crystals of TftC were soaked for 6 and

and a Dawn EOS laser light scattering detector (Wyatt Tech- 3 h, respectively, by adding substrate directly to the drop con-

nology). Scattering data were analyzed using the Zimm fitting taining the crystals.

method with software (ASTRA) provided by the instrument

manufacturer.

Crystals of TftD were made by mixing 1.5 ␮l of protein solu-

tion (19 mg ml^Ϫ1) with an equal volume of crystallization solu-

Dynamic Light Scattering—To test for the formation of a tion (20% (w/v) polyethylene glycol 4000, 0.2 ^MLiNO₃(pH 7.1)).

complex between TftC and TftD, we used a DynaPro Titan The purified TftD protein lost its FAD during purification, and

(Wyatt Technology Corp.). A scan was taken of each protein attempts to make a complex crystal were unsuccessful. Addi-

individually as well as a scan containing a mixture of TftC, TftD, tion of the reducing agent prevented growth of TftD crystals

FAD, and NADH. NADH was added just before each experi- and therefore was eliminated from the final (Sephacryl) purifi-

ment, and FAD reduction was confirmed by a change in color of cation step. Large single crystals grew slowly (over 1 month).

the mixture. Measurements consisted of 10 consecutive 5-s Both crystals of TftC and TftD were flash-frozen by transferring

scans, and average molecular radius was measured using a Ray- to the cryoprotection solutions containing the corresponding

leigh sphere approximation. Data were analyzed using the man- reservoir solutions plus 20% glycerol. The intensity data were

ufacturer-supplied software, Dynamics 6.7.3.

collected at the Advanced Light Source (ALS, BL 8.2.1) at 100 K

Isothermal Titration Calorimetry—The heats of binding of and reduced and scaled using HKL2000 (30) and CrystalClear

TftC and TftD with various cofactors were measured with a 1.3.6 (Rigaku/MSC).

VP-ITC Microcalorimeter (Microcal, Northampton, MA). For

Structure Determination and Refinement—Initial phasing of

calorimetric measurements, TftC or TftD in 20 m^MMops (pH TftC data were done by the program AMoRe (31) through the

7.2), 100 m^MNaCl, 1 mM dithiothreitol was titrated with one of coordinates of CobR (Protein Data Bank code 3CB0). Amino

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Crystal Structures of TftC and TftD

TABLE 1

Summary of crystallographic data

TftD

TftC

TftC-FAD

TftC-FAD-NADH

Space group

C222₁

R3

Unit cell parameters

a

b

c

148.20 Å

149.87 Å

212.01 Å

Energy (␭)

111.346 Å

111.529 Å

113.094 Å

102.997 Å

Resolution (Å)

102.523 Å

Completeness (%)

101.682 Å

Data

R_merge^a(%)

TftD phasing

Edge

0.97956

0.91840

0.97934

25 to 2.5

99.8

99.2

99.5

5.5

6.5

6.7

Remote

Peak

Refinement

TftD

TftC

TftC-FAD

TftC-FAD-NADH

Resolution range

No. of reflections (Ͼ0)

Completeness

20 to 2.5 Å

20 to 2.0 Å

79,750

31,822

25,673

27,350

98.0% (97.0%)

0.17 (0.18)

0.22 (0.25)

98.9% (93.0%)

0.15 (0.17)

0.18 (0.21)

79.9% (72.0%)

0.16 (0.25)

0.21 (0.30)

83.6% (70.0%)

0.16 (0.25)

0.20 (0.30)

b

R_work

c

R_free

No. of atoms

Nonsolvent

Solvent

15,336

1180

2,446

435

2,548

378

2,640

257

Wilson B-factor

Average B-value (Å²)

Nonsolvent

Solvent

22.6

23.2

25.2

27.8

33.6

33.8

26.0

40.7

29.3

42.7

30.8

41.6

r.m.s.d.^d

Bond lengths

Bond angles

Ramachandran plot^e

Favored

0.004 Å

0.757°

0.018 Å

1.763°

0.016 Å

1.737°

0.022 Å

2.504°

95.9%

99.1%

98.5%

100%

97.5%

100%

97.5%

100%

Allowed

^aR_mergeϭ (⌺_h⌺_i͉͗F_h͘ Ϫ F_hi͉)/⌺_hF_h, where ͗F_h͘ is the mean structure factor magnitude of i observations of symmetry-related reflections with Bragg index h.

^bR_crystϭ ⌺ʈF_obs͉ Ϫ ͉F_calʈ/͉F_obs͉.

^cR_freeis calculated with removal of Х5% of the data as the test set at the beginning of refinement.

^dRoot mean square deviations (r.m.s.d.) for main chain atoms are the root mean squared deviations of the bond lengths and bond angles from their respective ideal values as

implemented in PHENIX.

^eRamachandran plots were created using the program MolProbity (51).

acid mutations were then performed using the graphics pro- metric unit. A summary of the crystallographic data for both

gram O. Iterative model building and refinement took place

using the programs O, X-PLOR, and PHENIX.

TftC and TftD is given in Table 1.

Structure of TftC—An asymmetric unit of the TftC crystal

lattice contains a tightly associated homodimer, in which two

subunits are arranged by the noncrystallographic C₂axis. Each

TftC subunit is composed of 4 ␣-helices and 12 ␤-strands. Most

of those ␤-strands constitute a central twisted ␤-barrel, consist-

ing of ␤2, ␤3, ␤4, ␤5, ␤6, ␤9, and ␤10, which is the core of each

subunit. Two subunits are tightly assembled into a dimer by

several interactions as follows: (i) the hydrophobic interaction

between the 2-fold related side of each barrel; (ii) the ␣1 of each

subunit is deeply embedded into the opposite subunit, corking

its ␤-barrel; and (iii) a part of each ␤12 wraps around the ␤12 of

the other subunit in an inter-subunit anti-parallel ␤-sheet. This

dimeric nature of TftC in solution was confirmed by MALLS

data (supplemental Fig. S1). The other side of the ␤-barrel is

also corked by ␣2. Both ␣1 and ␣2 have an amphiphilic nature,

and the hydrophobic sides of those two ␣-helices face toward

the cavity of ␤-barrel, which is filled with hydrophobic residues.

The overall structures of the apo-form and two complex struc-

tures showed no major differences in terms of their backbone

structures. The C␣ carbons among the apo-form and the com-

plexes were superimposable with root mean square deviation

values of 0.13 and 0.17 Å for the FAD complex and FADꢁNADH

complex, respectively.

For TftD, location of the selenium heavy atom sites, phase

calculation, and parameter refinement were performed using

SOLVE (32) using a resolution range of 25 to 2.5 Å. This was

followed by maximum-likelihood density modification using

RESOLVE (32). After density modifications, well defined heli-

cal structures could be seen in the density, and the clear elec-

tron densities corresponding to most of the side chains allowed

model building to proceed. The model was built into the elec-

tron density using the graphics program O (33) and refined

using X-PLOR (34) and PHENIX (35). During the refinement

processes for both TftD and TftC, the noncrystallographic sym-

metry restraint has not been applied. The final coordinates have

been deposited in the Protein Data Bank as follows: apo-form

TftC (3K86), FADꢁTftC complex (3K87), FADꢁNADHꢁTftC

complex (3K88), and TftD (3HWC).

RESULTS

The apo-form of TftC was crystallized in the space group R3

with unit cell dimensions a ϭ b ϭ 111.35, c ϭ 103.00 Å, and

there were two TftC molecules in the asymmetric unit. Cell

dimensions for the complex crystals of TftC were a ϭ b ϭ

113.09, c ϭ 101.68 Å, and a ϭ b ϭ 111.53, c ϭ 102.52 Å for

FAD-NADH and FAD, respectively. TftD was crystallized in

the space group C222₁, with cell dimensions of a ϭ 148.20 Å,

As shown in supplemental Fig. S2 , in addition to the N and C

b ϭ 149.87 Å, c ϭ 212.01 Å, and four molecules in the asym- termini, three loop regions had elevated temperature factors as

JANUARY 15, 2010•VOLUME 285•NUMBER 3

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Crystal Structures of TftC and TftD

extended conformation and the

other a folded-back conformation.

The corresponding quality and cov-

erage of electron density for the

extended conformation was similar

to that of the folded-back site, but

the extended confirmation can be

superimposed well with that of FAD

in the FADꢁNADH complex struc-

ture. In addition, there is crystal

packing potentially hindering

a

proper diffusion for the folded-back

site. Therefore, only the extended

form of FAD will be discussed. The

O2 and O4 atoms of the isoallox-

azine ring are hydrogen-bonded to

backbone nitrogen atoms of Ala-51

and Asn-67. The N5 atom of the

isoalloxazine ring is also hydrogen-

bonded to the hydroxyl side chain of

Ser-50. Among those, only the ala-

nine residue is relatively conserved

among seven related flavin reducta-

ses (Fig. 3). The dimethyl benzene

moiety of the isoalloxazine ring is

located in a pocket formed by resi-

dues Tyr-171, Tyr-166, and Val-33,

all of which are highly conserved in

all flavin reductases (Fig. 3). The

pyrophosphate group of FAD is

located between ␣2 and ␣3, which

are in a perpendicular orientation

with respect to each other. Its phos-

phate and ribose moieties form

hydrogen bonds with Thr-48, Tyr-

71, Ala-99, and Arg-109, among

which only Arg-109 provides a side

chain interaction.

FIGURE2. RibbondiagramrepresentingthecrystalstructuresofTftCanditsbindingsite. A, stereoviewof

the TftC dimer with associated FAD molecule with one subunit shown in green and the other in red. Secondary

structural elements are labeled for one subunit and the noncrystallographic 2-fold axis is indicated by a half-

arrow. B, difference map (F_oϪ F_cmap contoured at ␴ ϭ 1.0) showed FAD molecule bound to TftC. Residues

mentioned in the text as binding to the substrate are shown in yellow. C, difference map (F_oϪ F_cmap con-

toured at ␴ ϭ 1.0) ternary complex of TftC showed both FAD and NADH. Residues important to substrate

binding are shown in yellow. Several secondary structural elements near the FAD and NADH are labeled.

Features marked with a prime are from the symmetry-related subunit. These figures were generated using

Open-Source PyMOL^TM(version 1.2).

The F_oϪ F_cmaps generated with

the data collected from the crystal

soaked with both FAD and NADH

follows: (i) between ␤5 and ␣2, (ii) between ␣3 and ␣4, and (iii) also clearly showed the corresponding electron density for both

␤10 and ␤11. As mentioned in detail later, the first and second FAD and NAD(H) molecules stacked via the isoalloxazine and

loops among those three participate in coordinating an FAD nicotinamide rings (Fig. 2C). In our FADꢁNADH ternary com-

molecule. Accordingly, the corresponding temperature factors plex of TftC, it is likely that the FAD molecules were in their

for Asn-67, Tyr-71, Ala-99, and Val-104, which are located in reduced state judged by the pale yellow color of the soaked

those two loops (Fig. 3), were reduced significantly upon com- crystal in contrast to the intense yellow color of the FAD-con-

plex formation with FAD.

taining crystal. The stacked isoalloxazine and nicotinamide

Substrate-binding Site of TftC—The F_oϪ F_cmaps of the rings were properly oriented at a distance of 3.5 Å between the

FAD-soaked crystal data of TftC showed the corresponding C4 of NAD(H) and the N5 of FAD(H₂). The position of the FAD

electron density for the bound FAD molecule (Fig. 2B), with its molecule in this FADꢁNADH complex was superimposable

isoalloxazine ring in a wide groove located at the dimer inter- with the FAD molecule in the FAD complex structure. The side

face. The boundary of the binding pocket is made of ␣1, ␤1, ␤3, chain of His-145 was within hydrogen bond distance with the

and ␤5 from one subunit and ␤4 of the other subunit. The amide group of nicotinamide, whose re-face (A-side) was facing

isoalloxazine ring sits on top of antiparallel sheet made of ␤2, the re-face of the isoalloxazine ring. The pyrophosphate group

␤3, and ␤6, facing its re-side toward the bulk solvent. The two of NAD(H) was hydrogen-bonded through the backbone of

FAD molecules in the homodimer have somewhat different Tyr-166 as well as the backbone and side chain of Arg-169. The

conformations of their ribityl moieties, one adopting an N3A atom of the adenine ring is within hydrogen bonding dis-

2018 JOURNAL OF BIOLOGICAL CHEMISTRY

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Crystal Structures of TftC and TftD

FIGURE 3. Multiple sequence alignment between TftC and other flavin reductases. The secondary structure of TftC is depicted using arrows, with violet

indicating ␤-strands and cyan indicating ␣-helices. Secondary structure elements for all other sequences are highlighted using the same color scheme.

Conserved residues are shown in boldface. CobR, corrin reductase from B. melitensis; 2R0X, putative flavins reductase from H. somnus 129pt; PheA2, phenol

2-hydroxylase component B from G. thermoglucosidasius; HpaC_Tt, flavin reductase component of 4-hydroxyphenylacetate 3-monooxygenase from T. ther-

mophilus hb8; HpaC_St, flavin reductase component of 4-hydroxyphenylacetate 3-monooxygenase from S. tokodaii str. 7; 2QCK, flavin reductase domain

protein from Arthrobacter sp. fb24.

tance from the hydroxyl group of Ser-54 from the other sub- and ␣16, which are all from the C-terminal segment, form a

unit. Significantly, all residues in the NAD(H)-binding site are sizeable helix bundle together with two short ␣-helices from

highly conserved among all compared flavin reductases. Con- the first segment, ␣4 and ␣5. The surfaces of most of those

sequently, the coordination mechanism observed in TftC is ␣-helices are mostly hydrophobic, establishing a hydrophobic

almost identical to those of previously reported flavin reducta- interaction among them.

ses, including the folded NAD(H) confirmation, first reported

The corresponding electron densities for the C-terminal res-

for Phe-2(A), with a distance between the adenine C6 and nic- idues 442–445 and 483–515 are not visible, probably due to

otinamide C2 atoms of 4.4 Å.

being disordered. In addition to the immediate neighbors of

Structure of TftD—Four tightly associated subunits in the these two regions, the temperature factors for residues 158–

crystal asymmetric unit are arranged in a noncrystallographic 167, which form an exposed loop connecting ␤5 and ␤6,

D₂symmetric assembly (Fig. 4B). The tetrameric nature of TftD showed higher temperature factors than the rest of the

in solution was confirmed by the elution profile and molecular molecule.

weight estimates from a MALLS experiment (supplemental Fig.

The observed tetramer interface had an extensive network of

S3 ). The individual TftD molecules consist of 14 ␤-strands and inter-subunit interaction among the C-terminal ␣-helices. In

17 ␣-helices (Fig. 4A) and can be divided into three sequential particular, ␣16 and ␣17 extend out of the individual subunit to

segments based on their constituent secondary structural ele- embed themselves into the hydrophobic surface made by ␣9,

ments. The first area from residues 1 to 146 is composed of ␣10, and ␣14 of the adjacent subunit. In addition, a large por-

seven ␣-helices (␣1–␣7) and four very short ␤-strands (␤1–␤4). tion of two anti-parallel C-terminal helices, ␣10 and ␣11, from

The second area, which is composed of residues 147–273, has two neighboring subunits contact closely in a perpendicular

eight ␤-strands (␤5–␤12). The third area, C-terminal segment orientation generating a substantial hydrophobic interface

is composed of 10 (-helices (␣8-␣17) and 2 short ␤-strands (␤13 between subunits. These hydrophobic interfaces involve resi-

and ␤14). Those three sequential segments are tightly intercon- dues such as Val-326, Leu-334, Leu-360, Phe-364, Leu-450,

nected in the three-dimensional structure. Noticeably, all of the Phe-454, and Met-458. The remaining three ␣-helices at the

10 helices in the C-terminal segment, ␣8–␣17, are involved in C-terminal segment also form another type of subunit interac-

an inter-subunit interaction as discussed in detail later. All the tion together with the above-mentioned ␤-barrel-like motif, i.e.

eight ␤-strands in the second segment, ␤5–␤12, are arranged in ␣12, ␣13, ␣15, their connecting loops, and the surface-exposed

such a way as to appear almost like a ␤-barrel. At one side of this residues in one side of the barrel form a tightly associated non-

barrel, the ␤1 and ␤2 strands from the first segment fold back crystallographic 2-fold related interface. This interface also

and together form a continuous ␤-sheet in the order of ␤2-␤1- possesses tightly packed hydrophobic residues as follows: Val-

␤10-␤11-␤7-␤6 (Fig. 4A). The remaining four strands in the 187, Val-190, Phe-230, Val-257, Leu-389, Met-398, Trp-405,

segment cover the other side of the same barrel in the order of and Phe-406. However, contrary to the above-mentioned

␤5-␤8-␤9-␤12. The amino acid side chains of those 10 hydrophobic interface involving ␣16 and ␣17, a substantial

␤-strands and the connecting loops between them form a amount of polar inter-subunit interaction was observed, which

hydrophobic core inside of the barrel.

includes Asp-159, Glu-233, Lys-237, Asp-250, Asp-259, Arg-

Another noticeable structural feature is that three long 386, Gln-396, Gln-401, Asn-410, Arg-420, Arg-428, Asp-434,

␣-helices, ␣9, ␣10, and ␣11, and three short helices, ␣14, ␣15, and Arg-438.

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Crystal Structures of TftC and TftD

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Crystal Structures of TftC and TftD

FIGURE 5. Measurement of substrate binding through ITC experiments. A, trend of heat released by serial injections of FAD (square), FMN (inverted

triangle), or NADH (circle) into apo-form TftC was monitored. Only FAD showed the typical heat-release pattern indicative of binding. Solid lines

represent the least square fits of the data using a single-site binding model. B, injections of FAD (diamond), FMN (triangle), FADH₂(circle), and FMNH₂

(inverted triangle) into apo-form TftD.

Substrate-binding Site of TftD—In the apo-structure of TftD, ␣15 of one subunit and portions of ␣9, ␣10, and ␣11 of

a unique entry site and binding pocket for FADH₂was clearly another subunit.

distinguishable (Fig. 4C). The perimeter of this widely open entry

Isothermal Titration Calorimetry (ITC) for TftC and TftD—

site is made with three flexible loops as follows: (i) between ␤5 and ITC was used to characterize the binding of FAD, FMN, and

␤6, (ii) between ␤8 and ␤9, and (iii) between ␣15 and ␣16. Among NADH to TftC. As shown in Fig. 5A, a significant amount of

those three, the first and third loops have high temperature factors heat is released when TftC is titrated with FAD. However, a

and show the most heterogeneous conformations among the four negligible amount of heat was released when TftC was titrated

subunits, reflecting their high level of flexibility. The residues lin- with FMN and NADH, indicating a lack of binding. A calcu-

ing this entry site are predominantly hydrophilic, and a few lated K_dof FAD is 1.8 Ϯ 0.1 ␮M (mean Ϯ S.D.), with a large

ordered solvent molecules were located in the entry site of the enthalpic contribution (⌬H ϭ Ϫ11.5 kcal/mol). The data also

determined apo-form structure of TftD.

indicated an unfavorable entropic contribution, with a calcu-

The position of the riboflavin moiety of the FADH₂was lated ⌬S of Ϫ12.29 cal/mol/degree, possibly indicating that the

readily obtained through the combination of superposition TftC structure was slightly stabilized upon binding of FAD, and

with the 4-hydroxyphenylacetate 3-monooxygenase (HpaB) very few solvent molecules were freed from the pocket. These

from Thermus thermophilus HB8 (2YYJ) (36) and the solid thermodynamic data were consistent with structural data, as

docking module on Quanta (BioSYM/MSI, Inc.), which is based indicated by the significant reduction of the B-values for two

on conformational space, followed by a quick energy minimi- loops constituting the binding pocket upon formation of the

zation by CNS version 1.1 (37). The wall of this binding pocket FAD binary complex. There were few solvent molecules in the

is made with small portions from ␤5, ␤7, ␤8, ␤11, ␣9, and FAD binding pocket.

FIGURE4. A, ribbondiagramrepresentingthe crystal structure of TftD subunit. Secondary structureelementshavebeen numbered sequentially as ␣1–␣17 and

␤1–␤14 following the convention. N and C refer to N- and C-terminal regions, respectively. ␤-Strands are shown in magenta, and ␣-helices are shown in cyan.

The subunit orientation is analogous to that of the green subunit in B for reference. B, arrangements of the tetrameric TftD in a noncrystallographic D₂

symmetry. A few secondary structural elements have been labeled as a guide. C, structure of the substrate-binding pocket of TftD with FAD and 2,5-DiCHQ. The

participating residues in binding 2,5-DiCHQ and FAD are shown with their residue numbers and lines representing hydrogen bonds. The chlorine atoms in

2,5-DiCHQ has been depicted in green. These figures were generated using Open-Source PyMOL^TM(version 1.2).

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Crystal Structures of TftC and TftD

TABLE 2

Kinetic properties of TftD and TftD H289A

K_m

␮M

k_cat

s

k

_cat/K_m

Ϫ1

M

^Ϫ1s

2,4,5-TCP

TftD^a

35.8 Ϯ 3.9

36.6 Ϯ 3.4

0.67 Ϯ 0.03

1.9 ϫ 10⁴

6.6 ϫ 10²

TftD H289A

0.024 Ϯ 0.003

2,4,6-TCP

(28 times)

TftD^a

39.9 Ϯ 7.6

18.5 Ϯ 2.4

0.41 Ϯ 0.03

1.0 ϫ 10⁴

4.2 ϫ 10²

TftD H289A

0.0078 Ϯ 0.0006

2,5-DiCHQ

TftD H289A^b

(53 times)

0.10 Ϯ 0.01

0

TftD^a

4.3 Ϯ 1.1

2.3 ϫ 10⁴

^aExperiments were done in 40 mM KP_ibuffer (pH 7.0) at 24 °C. Values are means of

triplicate experiments with standard deviations. The kinetic properties of TftD

were reported previously (8).

^bNo activities.

As discussed under “Discussion,” the structure of TftD indi-

cated that one of the residues in the active site, His-289, might

play a critical role in its catalysis, and thus TftD H289A mutant

enzyme was generated, and its structural integrity was com-

pared with that of the wild type by measuring their CD spectra.

As shown in the supplemental Fig. S4 A , the spectral pattern of

H289A mutant in a buffer containing 20 m^Msodium phosphate

(pH 6.8) was superimposable to that of the wild type. Both have

two negative bands around 208 and 222 nm, which are the char-

acteristics of CD spectra of typical ␣-rich or ␣␤ proteins. The

tetrameric nature of the H289A mutant in solution was also

confirmed by a MALLS experiment (supplemental Fig. S4 B).

The enzyme activities for the mutant were assayed using 2,4,6-

TCP, 2,4,5-TCP, or 2,5-DiCHQ as the substrate. TftD H289A

mutant still used 2,4,5-TCP and 2,4,6-TCP as substrates with

about 28- and 53-fold decreases in k_cat, respectively, and the K_m

values were not significantly altered (Table 2). Most signifi-

cantly, the mutant enzyme completely lost activity toward

2,5-DiCHQ.

FIGURE6. KineticanalysisofTftC. Double-reciprocalplotsofinitialvelocities

of TftC versus substrate concentration. A, each line represents an experiment

with a constant concentration of FAD, whereas the concentration of NADH is

varied. B, each line is an experiment in which NADH was held constant while

varying FAD concentration. The double-reciprocal plot yielded lines inter-

secting to the left of the y axis, indicating a sequential reaction mechanism.

Error bars indicate 1 S.D.

Dynamic Light Scattering—To probe the dynamics of flavin

transfer between the small component flavin reductase and the

large component monooxygenase, chemical cross-linking and

dynamic light scattering were employed. The average Rayleigh

sphere radius of TftC and TftD was measured independently

and in combination with TftC substrates. When a solution of

TftC and TftD (also containing FAD and NADH) was mea-

sured, FAD was reduced to FADH₂. As was confirmed by the

color change and the FADHOOH spectrum (8), the bound

FADH₂in TftD was immediately oxidized to FADHOOH.

Thus, most added FAD is in the form of the FADHOOHꢁTftD

complex. If a higher order complex were formed between the

two enzymes to transfer FADH₂, the height of the TftC peak

would either decrease or disappear, whereas the TftD peak

would appear to shift to a larger radius. As Fig. 7 indicates,

addition of substrates (FAD and NADH) causes no significant

shift in intensity or any shift in average particle size, indicating

a lack of interaction between the two enzymes. Similar

experiments were carried out using chemical cross-linking,

and the results indicated the same outcome (data not

shown). This supports the growing body of evidence that

As in the case of TftC, ITC was used to confirm the differen-

tial binding affinities of FAD and FMN for TftD and their cor-

responding reduced forms. As shown in Fig. 5B, FAD, FMN,

and FMNH₂molecules did not show any significant binding to

TftD. However, a significant amount of heat was released when

FADH₂associated with TftD, indicating that the binding inter-

actions had significant enthalpic contributions (⌬H ϭ Ϫ8.33

kcal/mol). The data also showed a slightly unfavorable entropic

contribution (⌬S ϭ Ϫ0.78 cal/mol/degree); however, it is less

significant than ⌬S of TftC. ITC data analysis yielded a K_dfor

FADH₂binding to TftD of 1.2 Ϯ 0.2 ␮M.

Enzyme Kinetics and Site-directed Mutagenesis—Steady-

state kinetics experiments were performed by holding the con-

centration of TftC constant while varying the FAD concentra-

tion at 1, 2, 4, 6, and 8 ␮M. The NADH concentrations in the

corresponding four experiments were varied at 25, 50, 100, and

200 ␮M, and each point was performed in triplicate. As shown

in Fig. 6, the double-reciprocal plot of the inverse rate versus

inverse substrate concentration yielded lines intersecting to the

left of the y axis, indicating a sequential reaction mechanism.

The calculated K_mvalues for FAD and NADH were 2.0 (Ϯ0.3)

and 45.3 (Ϯ4.5) ␮M, respectively. The V_maxvalue was 120.3 flavin transfer in TC-FDM systems occurs via a purely diffu-

Ϫ1

(Ϯ7.7) ␮mol min^Ϫ1mg

.

sive mechanism (21, 38, 39).

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Crystal Structures of TftC and TftD

revealed a structural heterogeneity

in terms of the number and size of

helices in the peripheral regions.

Additionally, the C terminus of each

TftC subunit forms interlocking

anti-parallel ␤-strands with each

other, which has not been observed

in other flavin reductases. Notice-

ably the structure of TftC is quite

different from that of EmoB, which

is also a small component of

TC-FDM (19). A search for similar

amino acid sequences in the Protein

Data Bank using BLAST (42)

revealed that putative flavin reduc-

tase from H. somnus (2R0X) and

CobR (3CB0) showed the highest

score (119 bits) with 40 and 38%

identity among matched amino

acids to TftC, respectively, followed

by Phe-2(A) (1RZ0, 70.5 bits, 31%

identity), HpaC_St(2D36; 57.4 bits,

FIGURE 7. Dynamic light scattering analysis. Dynamic light scattering was used to probe for physical inter-

25% identity), and HpaC_Tt(2ECR;

56.6 bits, 26% identity). The con-

served residues were distributed

sporadically through the entire sec-

actionbetweenTftCandTftDduringreducedflavintransfer. TftCisthedashedline;TftDisthedash/dotline, and

a mixture of the two proteins with FAD and NADH is shown as a solid line. The plot indicates a lack of physical

interaction between the two proteins, indicating the reduced flavin could transfer via a diffusive mechanism.

DISCUSSION

ondary structural elements. The ␣2 and its neighboring loops

show the least similarity among the above reductases (Fig. 3). In

addition, there are insertions or deletions between ␣3 and ␤7

producing heterogeneity in the secondary structural elements

among those compared flavin reductases. Significantly, this

area of heterogeneity happens to be involved in binding of the

adenine moiety of flavin in those reductases. In Phe-2(A) and

HpaC_Tt, binding for the adenine moiety is through multiple

hydrogen bonds with this region referred to as Loop 7. How-

ever, as noted previously in the structure of CobR (43), Loop 7 is

replaced by an additional ␣-helix, ␣4, in TftC, which alters the

binding interactions with AMP. It should also be noted that

HpaC of S. tokodaii, which prefers FMN to FAD, contains a 3₁₀

helix in the same position as ␣4 in TftC.

We have determined the crystal structures of both TftC and

TftD to shed light on their reaction mechanism and structural

relationship to other flavin-dependent monooxygenases and

flavin reductases in TC-FDM. In particular, TftC and TftD

together perform two sequential oxidative dechlorination steps

in the degradation process of 2,4,5-TCP (Fig. 1) (8), even

though most monooxygenases hydroxylate their substrate only

once (40, 41). For TftC, in addition to its apo-form, structures of

the binary complex with FAD and the ternary complex with

FAD and NADH were also determined. However, we were

unable to produce a complex crystal of TftD with FAD under

aerobic environments, likely due to the negligible affinity of

TftD for FAD as shown by our ITC data (Fig. 5B). Conse-

quently, the position of flavin was determined through super-

imposition with structural homologues and a substrate docking

approach.

The residues constituting the hydrophobic binding pocket

for the dimethyl benzene moiety of the isoalloxazine ring are

highly conserved among those enzymes. However, the corre-

sponding sites facing the 2,4-pyridine side and the ribose group

of the flavin molecule are not well conserved among the struc-

Structural Classification for TftC

The primary and tertiary structural comparison of TftC turally related proteins (Fig. 3). Despite this poor sequence sim-

revealed its high similarity with other flavin reductases. A Dali ilarity, the backbones of the corresponding regions are similarly

search showed that the most similar structure was corrin engaged in coordination in all the compared reductases whose

reductase (CobR) from Brucella melitensis (3CB0) with a high complex structures are available.

Z-score of 33.5, followed by putative flavin reductase from Hae-

Structural Classification for TftD

mophilus somnus (2R0X) with a Z-score of 26.3, Phe-2(A) from

Geobacillus thermoglucosidasius (1RZ1) with 23.3, HpaC from

A Dali search was also used for identifying structural homo-

Sulfolobus tokodaii str. 7 (2D37) with 21.8, and HpaC from T. logues for TftD (45) and showed that the highest match was to

thermophilus (2ECR) with 21.0. In general, the ␤-barrel-like the 4-hydroxyphenylacetate 3-monooxygenase (HpaB) of T.

core structure and dimeric nature of TftC are similar to the thermophilus HB8 (2YYL) (36) with a Z-score of 43.7. It was

structures of high similarity scores, which include other small followed by 4-hydroxybutyryl-CoA dehydratase (4-BUDH) of

component members of the TC-FDM family, such as Phe-2(A), Clostridium aminobutyricum (1U8V) (46) with a Z-score of

HpaC_St, and HpaC_Tt. However, a detailed visual inspection 36.5 and a couple of other structural homologues with much

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Crystal Structures of TftC and TftD

HpaB share 28% identity and 48%

similarity in their amino acid

sequences. However, there were

also significant structural differ-

ences among them. For example,

in HpaB, the ␣6 is missing due to a

major deletion near that region

(Fig. 8).

Substrate-binding Site and

Reaction Mechanism

TftC—The observed conforma-

tion of the AMP moiety of the

bound FAD in TftC is quite differ-

ent from that in the complex crystal

of the Phe-2(A) and HpaC; likewise,

the local conformation of TftC

shows difference from those, i.e. the

exposed adenine ring of the FAD

stacks with the phenyl side chain of

tyrosine 71 in the TftC structure, in

contrast to the definite binding

pocket offering multiple hydrogen

bonds to the adenine ring observed

in both Phe-2(A) and HpaC. This

apparent difference in binding of

the AMP moiety of FAD might

result in a lower affinity for FAD

(K_dϭ 1.8 ␮M) and loss of FAD dur-

ing the purification. In addition,

there was no flavin occupancy in the

apo-form crystal structure. This is

in contrast with what has seen in

other flavin reductases, such as Phe-

2(A), which has a nanomolar affinity

for flavin.

The NADH:FAD oxidoreductase

activity of TftC was through oxida-

tion of an NADH molecule by a

bound FAD molecule, which caused

reduction of the flavin. The ternary

complex structure of TftC showed

that the stacked nicotinamide:

isoalloxazine rings were at a proper

distance for hydride transfer. Com-

pared with the FAD molecule, the

NAD(H) molecule in the ternary

complex of TftC had less interaction

with the enzyme despite its interac-

tions with highly conserved residues

FIGURE 8. Amino acid sequence and secondary structure alignment of TftD with other flavin-dependent

monooxygenases. Secondary structure elements for TftD are depicted as violet arrows and cyan cylinders for

␤-strandsand␣-helices, respectively. SecondarystructuralelementsforHpaBand4-BUDHarealsohighlighted

in colors (violet for ␤-strands and cyan for ␣-helices). Conserved residues among those monooxygenases are

shown in boldface. TcpA, 2,4,6-trichlorophenol monooxygenase from Cupriavidus necator JMP134; HpaB, 4-hy-

droxyphenylacetate3-monooxygenasefromT. thermophilusHB8;4-BUDH, 4-hydroxybutyryl-CoAdehydratase

from C. aminobutyricum.

and many water-mediated interactions, which are very similar

to that of Phe-2(A). The NAD(H) molecule had a negligible

affinity with TftC that had no bound FAD in it, as indicated by

the ITC results (Fig. 5A). Therefore, the apparent affinity for

NADH existed only after the FAD occupies its site in TftC. In

lower Z-scores such as medium chain-specific acyl-CoA dehy-

drogenase (MCAD) from pig mitochondria (3MDE) (47). Care-

ful manual comparisons of these crystal structures show that

they share similarity with TftD in terms of the location of sec-

ondary structural elements and topological connectivity (Fig.

8). Consistent with a structural similarity among TftD, MACD, addition, in the crystal structure for the FADꢁNADH ternary

HpaB, and 4-BUDH, there is significant sequence similarity complex form, the adenine ring is folded back in a stacking

among these structural homologues. For example, TftD and position on top of the nicotinamide ring, probably shielding the

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Crystal Structures of TftC and TftD

tion in TftD. Therefore, upon

FADH₂binding, a conformational

change might follow in that local

area to accommodate binding of the

pyrophosphate and adenosine moi-

eties. However, considering

a

noticeably weaker affinity of TftD

(micromolar range) for FADH₂

compared with that of HpaB (nano-

molar range) (5) and negligible

affinity for both FAD and FMN,

TftD might have intrinsically poor

coordination for the ADP portion of

the FAD.

The geometry and several polar

residues in the si-face of the mod-

eled isoalloxazine ring are well

conserved among TftD, MCAD

(3MDE), HpaB (2YYJ), and

4-BUDH (1U8V). Thr-192 is com-

pletely conserved among all four

FADH₂-dependent enzymes, and

their corresponding hydroxyl group

side chains are in ϳ3.0 Å distance

from the N5 atom of the isoallox-

azine ring (Fig. 4C). In addition,

three other nearby residues,

Arg-100, Asp-253, and Arg-438, are

conserved among TftD, TcpA, and

HpaB (Fig. 8). The side chain of Arg-

100 has been proposed to form a

hydrogen bond with the peroxy

FIGURE 9. Proposed reaction mechanism in the active site of TftD. A, 2,4,5-TCP as a substrate; B, 2,5-DiCHQ

as a substrate. In both cases, His-289 acts as a general base. The arrows indicate the movement of an electron.

The figures were made using ChemBioDraw Ultra 11.0 (Cambridge Corp.).

C4 atom of nicotinamide from bulk solvent (Fig. 2C). This is moiety of the C4a-hydroperoxy flavin intermediate and to con-

also observed in both HpaC and Phe-2(A) structures (48), tribute to the formation and/or stabilization of that transient

allowing the efficient hydride transfer to FAD. Once FAD in intermediate (36, 50). As observed in the HpaBꢁFAD complex

TftC is reduced by NADH, the FAD molecule will leave as structure, the N⑀ atom of Arg-100 of TftD is located 4.8 Å from

FADH₂. Based on the results from the double-reciprocal plot, it the C4a atom of the modeled isoalloxazine ring.

is obvious that the production of FADH₂by TftC is through a

The K_dof HpaB for FADH₂is in a nanomolar range com-

sequential reaction mechanism (Fig. 6). In the catalytic cycle pared with the micromolar range for FAD binding (21). This

ϩ

NADH reduces the associated FAD and NAD leaves TftC. substantially higher affinity of HpaB for FADH₂over FAD has

The diffusible FADH₂is then used by TftD for TCP metabo- been attributed to the immediate peroxidation of FADH₂and

lism. In this process, the NADH molecule delivers its hydride its subsequent stabilization through the guanidinium side chain

through transiently stacked interactions of its nicotinamide of Arg-100 (36). In contrast to micromolar range affinity of

ring with the isoalloxazine ring of FAD as similarly observed in HpaB for FAD, our ITC data show that TftD does not have

the complex structures of EmoB, Phe-2(A), and HpaC (19, 44, significant affinity to FAD (Fig. 5B). However, in a completely

48, 49).

anaerobic setting, which does not allow the formation of FAD

TftD—The secondary structural elements of TftD around the or FAD-peroxide, TftD shows a K_dof 1.2 ␮M for FADH₂. There-

putative substrate-binding pocket were superimposable with fore, there is a clear difference in flavin affinity between TftD

those in available holo-form structures of the above-mentioned and HpaB.

enzymes, MCAD (3MDE), HpaB (2YYJ), and 4-BUDH (1U8V).

In the si-side of TftD, there exists a significant difference

Therefore, a reliable position for the riboflavin moiety of among apolar residues in the surroundings of the bound

FADH₂could be established. However, the residues for coordi- isoalloxazine ring compared with the same side of the above-

nating the pyrophosphate or adenosine group of the FAD(H₂) mentioned flavoenzymes. Noticeably, Val-154 is unique for

could not be pinpointed due to the conformational differences TftD; the amino acid at the equivalent position is a threonine in

among the four subunits of apo-form TftD. The two conserved HpaB, MCAD, and 4-BUDH. In particular, the side chain of

residues (Fig. 8), Gln-157 and Arg-160, which reside in the flex- that threonine residue in MCAD and 4-BUDH is in a position to

ible loop between ␤5 and ␤6 and coordinate the phosphate hydrogen bond with the N1 and O2 atoms of the isoalloxazine

group of FADH₂in HpaB (36), are not in the coordination posi- ring. Therefore, the substitution of this residue to an apolar

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Crystal Structures of TftC and TftD

amino acid in TftD can be another reason for its negligible His-289 residue of TftD plays critical bifunctional roles in its

unique sequential catalysis of 2,4,5-TCP and 2,5-DiCHQ.

affinity for FAD in addition to the above-mentioned poor coor-

dination of its AMP moiety. In addition, the corresponding res-

idue for Val-190 is threonine, histidine, and tryptophan in

HpaB, 4-BUDH, and MACD, respectively. Both histidine and

tryptophan are located in proximity to the riboflavin part of the

Closing Remark

Characterizing the participating key enzymes is the prereq-

uisite for informed and successful bioremediation of major pol-

attached FAD, which could reflect its tighter affinity as a cofac- lutants, which will improve our understanding of the reaction

mechanisms and increase our ability to remove these pollutants

from the environment. Our data offer the mechanistic under-

standing of TftC and TftD activities. The results support that

free FADH₂is generated by TftC and that TftD uses this dif-

fused FADH₂to transform 2,4,5-TCP and 2,5-DiCHQ. Fur-

thermore, the different roles of His-289 of TftD in the catalysis

of 2,4,5-TCP and 2,5-DiCHQ are proposed, explaining how a

monooxygenase attacks two substrates. Besides contributions

to basic biochemistry, the results may facilitate bioremediation

of polychlorinated phenols.

tor in 4-BUDH and MACD. In both TftD and HpaB, FADH₂is

a substrate instead of a cofactor.

The residues facing the re-side of the isoalloxazine ring,

which is presumably a 2,5-DiCHQ or TCP-binding site, are

quite different among the compared enzymes, as expected from

their substrate specificities. In comparison with the corre-

sponding residues of HpaB, the residues of TftD in the re-side of

the isoalloxazine ring are much more apolar. For example, the

corresponding residues for Leu-151, Ile-205, and Val-288 are

polar in HpaB. In addition there are several other noticeable

differences between these two closely related flavin-dependent

monooxygenases. Previously, Tyr-104 and His-142 residues in

HpaB have been noticed to coordinate the hydroxyl group of its

substrate, 4-hydroxyphenyl acetate, and to abstract the proton

from it, respectively. The corresponding amino acids in TftD

are Ala-104 and Val-151; thus, neither of these TftD residues is

able to coordinate the hydroxyl group of 2,5-DiCHQ or TCP.

His-289 in the putative substrate-binding site, which is located

on the re-side of the isoalloxazine ring, is the only candidate

that could coordinate the hydroxyl group or proton abstraction

from the bound substrate. Establishing a hydrogen bond

between the imidazole side chain of His-289 and one of the

hydroxyl groups of the modeled 2,5-DiCHQ resulted in the

other hydroxyl group facing toward the solvent-exposed side of

the pocket (Fig. 4C). Our results from site-directed mutagene-

sis, MALLS and CD spectra confirmed that the His-289 residue

is involved in catalysis. Despite the completely abolished activ-

ity toward 2,5-DiCHQ, the H289A mutant still maintains the

same CD and MALLS profile and some activity but significantly

decreased the k_catfor the oxidation of both 2,4,5-TCP and 2,4,6-

TCP (about 28- and 53-fold less, respectively), and the K_mval-

ues were only slightly altered (less than 2-fold) (Table 2). There-

fore, based on these enzymatic data, the hydroxyl group of both

2,4,5- and 2,4,6-TCP may face toward the solvent area in an

orientation similar to the acetate group of 4-HPA, instead of

being hydrogen-bonded to the imidazole of His-289 (Fig. 9A).

In this orientation, 4-chlorine faces into the hydrophobic side,

which agrees with the hydroxylation reactions occurring in the

para-position for TCP versus the ortho-position (to the

hydroxyl group) for 4-HPA. The His-289 residue is not likely

involved in abstracting a proton from the phenol group of TCP;

instead, it is involved in abstraction of the proton from the

added hydroxyl group in the reaction intermediate, facilitating

the chlorine removal and catalytic turnover (Fig. 9A). Conse-

quently, the H289A mutant slows down the catalysis of TCP.

However, the His-289 residue is involved in abstracting a pro-

ton from a hydroxyl group of 2,5-DiCHQ for the hydroxylation

at the ortho-position as noticed for 4-HPA oxidation by HpaB

(Fig. 9B). Thus, the mutant completely loses the dechlorination

Acknowledgments—We thank C. Ralston (Berkeley Advanced Light

Source, beamline 8.2.1) and T. Terwilliger (Los Alamos National

Laboratory).

REFERENCES

1. Czaplicka, M. (2004) Sci. Total Environ. 322, 21–39

2. McCollister, D. D., Lockwood, D. T., and Rowe, V. K. (1961) Toxicol. Appl.

Pharmacol. 3, 63–70

3. Kilbane, J. J., Chatterjee, D. K., Karns, J. S., Kellogg, S. T., and Chakrabarty,

A. M. (1982) Appl. Environ. Microbiol. 44, 72–78

4. Kellogg, S. T., Chatterjee, D. K., and Chakrabarty, A. M. (1981) Science

214, 1133–1135

5. Louie, T. M., Webster, C. M., and Xun, L. (2002) J. Bacteriol. 184,

3492–3500

6. Xun, L. (1996) J. Bacteriol 178, 2645–2649

7. Hu¨bner, A., Danganan, C. E., Xun, L., Chakrabarty, A. M., and Hendrick-

son, W. (1998) Appl. Environ. Microbiol. 64, 2086–2093

8. Gisi, M. R., and Xun, L. (2003) J. Bacteriol. 185, 2786–2792

9. Perry, L. L., and Zylstra, G. J. (2007) J. Bacteriol. 189, 7563–7572

10. Otto, K., Hofstetter, K., Ro¨thlisberger, M., Witholt, B., and Schmid, A.

(2004) J. Bacteriol. 186, 5292–5302

11. Gala´n, B., Díaz, E., Prieto, M. A., and García, J. L. (2000) J. Bacteriol. 182,

627–636

12. Kirchner, U., Westphal, A. H., Mu¨ller, R., and van Berkel, W. J. (2003)

J. Biol. Chem. 278, 47545–47553

13. Malito, E., Alfieri, A., Fraaije, M. W., and Mattevi, A. (2004) Proc. Natl.

Acad. Sci. U.S.A. 101, 13157–13162

14. Li, L., Liu, X., Yang, W., Xu, F., Wang, W., Feng, L., Bartlam, M., Wang, L.,

and Rao, Z. (2008) J. Mol. Biol. 376, 453–465

15. Valton, J., Fontecave, M., Douki, T., Kendrew, S. G., and Nivie`re, V. (2006)

J. Biol. Chem. 281, 27–35

16. Massey, V. (1994) J. Biol. Chem. 269, 22459–22462

17. Filisetti, L., Fontecave, M., and Niviere, V. (2003) J. Biol. Chem. 278,

296–303

18. Gao, B., and Ellis, H. R. (2005) Biochem. Biophys. Res. Commun. 331,

1137–1145

19. Nissen, M. S., Youn, B., Knowles, B. D., Ballinger, J. W., Jun, S. Y., Belchik,

S. M., Xun, L., and Kang, C. (2008) J. Biol. Chem. 283, 28710–28720

20. Gibson, Q. H., and Hastings, J. W. (1962) Biochem. J. 83, 368–377

21. Louie, T. M., Xie, X. S., and Xun, L. (2003) Biochemistry 42, 7509–7517

22. Sucharitakul, J., Phongsak, T., Entsch, B., Svasti, J., Chaiyen, P., and Ballou,

D. P. (2007) Biochemistry 46, 8611–8623

23. Jeffers, C. E., Nichols, J. C., and Tu, S. C. (2003) Biochemistry 42, 529–534

24. Low, J. C., and Tu, S. C. (2003) Photochem. Photobiol. 77, 446–452

activity toward 2,5-DiCHQ. Therefore, we propose that the 25. Abdurachim, K., and Ellis, H. R. (2006) J. Bacteriol. 188, 8153–8159

2026 JOURNAL OF BIOLOGICAL CHEMISTRY

VOLUME 285•NUMBER 3•JANUARY 15, 2010

Crystal Structures of TftC and TftD

26. Lee, J. K., and Zhao, H. (2007) J. Bacteriol. 189, 8556–8563

27. Kantz, A., Chin, F., Nallamothu, N., Nguyen, T., and Gassner, G. T. (2005)

Arch. Biochem. Biophys. 442, 102–116

38. Valton, J., Mathevon, C., Fontecave, M., Nivie`re, V., and Ballou, D. P.

(2008) J. Biol. Chem. 283, 10287–10296

39. Campbell, Z. T., and Baldwin, T. O. (2009) J. Biol. Chem. 284, 8322–8328

40. Newman, L. M., and Wackett, L. P. (1995) Biochemistry 34, 14066–14076

41. Kadiyala, V., and Spain, J. C. (1998) Appl. Environ. Microbiol. 64,

2479–2484

28. Tiirola, M. A., Ma¨nnisto¨, M. K., Puhakka, J. A., and Kulomaa, M. S. (2002)

Appl. Environ. Microbiol. 68, 173–180

29. Youn, B., Moinuddin, S. G., Davin, L. B., Lewis, N. G., and Kang, C. (2005)

J. Biol. Chem. 280, 12917–12926

42. Altschul, S. F., Madden, T. L., Scha¨ffer, A. A., Zhang, J., Zhang, Z., Miller,

W., and Lipman, D. J. (1997) Nucleic Acids Res. 25, 3389–3402

43. Lawrence, A. D., Deery, E., McLean, K. J., Munro, A. W., Pickersgill, R. W.,

Rigby, S. E., and Warren, M. J. (2008) J. Biol. Chem. 283, 10813–10821

44. van den Heuvel, R. H., Westphal, A. H., Heck, A. J., Walsh, M. A., Rovida,

S., van Berkel, W. J., and Mattevi, A. (2004) J. Biol. Chem. 279,

12860–12867

30. Otwinowski, Z., and Minor, W. (1997) Methods Enzymol. 276, 307–326

31. Navaza, J. (2001) Acta Crystallogr. D. Biol. Crystallogr. 57, 1367–1372

32. Terwilliger, T. C. (2001) Acta Crystallogr. D. Biol. Crystallogr. 57,

1755–1762

33. Jones, T. A., Zou, J. Y., Cowan, S. W., and Kjeldgaard, M. (1991) Acta

Crystallogr. A 47, 110–119

34. Bru¨nger, A., Adams, P. D., Clore, G. M., DeLano, W. L., Gros, P., Grosse- 45. Holm, L., and Sander, C. (1993) J. Mol. Biol. 233, 123–138

Kunstleve, R. W., Jiang, J. S., Kuszewski, J., Nilges, M., Pannu, N. S., Read, 46. Martins, B. M., Dobbek, H., Cinkaya, I., Buckel, W., and Messerschmidt,

R. J., Rice, L. M., Simonson, T., and Warren, G. L. (1998) Acta Crystallogr.

Sect. D. Biol. Crystallogr. 54, 905–921

A. (2004) Proc. Natl. Acad. Sci. U.S.A. 101, 15645–15649

47. Kim, J. J., Wang, M., and Paschke, R. (1993) Proc. Natl. Acad. Sci. U.S.A. 90,

7523–7527

35. Adams, P. D., Grosse-Kunstleve, R. W., Hung, L. W., Ioerger, T. R., Mc-

Coy, A. J., Moriarty, N. W., Read, R. J., Sacchettini, J. C., Sauter, N. K., and

Terwilliger, T. C. (2002) Acta Crystallogr. D Biol. Crystallogr. 58,

1948–1954

48. Okai, M., Kudo, N., Lee, W. C., Kamo, M., Nagata, K., and Tanokura, M.

(2006) Biochemistry 45, 5103–5110

49. Kim, S. H., Hisano, T., Iwasaki, W., Ebihara, A., and Miki, K. (2008) Pro-

teins 70, 718–730

36. Kim, S. H., Hisano, T., Takeda, K., Iwasaki, W., Ebihara, A., and Miki, K.

(2007) J. Biol. Chem. 282, 33107–33117

50. Torres Pazmin˜o, D. E., Baas, B. J., Janssen, D. B., and Fraaije, M. W. (2008)

Biochemistry 47, 4082–4093

37. Bru¨nger, A. T., Adams, P. D., Clore, G. M., DeLano, W. L., Gros, P.,

Grosse-Kunstleve, R. W., Jiang, J. S., Kuszewski, J., Nilges, M., Pannu, N. S., 51. Davis, I. W., Leaver-Fay, A., Chen, V. B., Block, J. N., Kapral, G. J., Wang,

Read, R. J., Rice, L. M., Simonson, T., and Warren, G. L. (1998) Acta

Crystallogr. D Biol. Crystallogr. 54, 905–921

X., Murray, L. W., Arendall, W. B., 3rd, Snoeyink, J., Richardson, J. S., and

Richardson, D. C. (2007) Nucleic Acids Res. 35, W375–W383

JANUARY 15, 2010•VOLUME 285•NUMBER 3

JOURNAL OF BIOLOGICAL CHEMISTRY 2027