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3 1293 02079 9395

This is to certify that the

dissertation entitled

Characterization and Function of
2,4—Dichlorophenoxyacetic Acid/ alpha—Keto-
glutarate Dioxygenase and Related Enzymes

presented by

Deborah Ann Hogan

has been accepted towards fulfillment
of the requirements for

Ph . D . degree in Microbiology

 

 

\

Major professor /

 

Date 12113/99

MSU is an Affirmative Action/Equal Opportunity Institution 0-12771

 

 

_-.——— ——.— ‘21-).

mat-mm .
Michigan State
University

 

 

CHARACTERIZATION AND FUNCTION OF 2,4-DICHLORO-
PHENOXYACETIC ACID/a-KETOGLUTARATE-DEPENDENT DIOXYGENASE
AND RELATED ENZYMES
By

Deborah Ann Hogan

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY
Department of Microbiology

1999

ABSTRACT
CHARACTERIZATION AND FUNCTION OF 2,4-DICHLORO-
PHENOXYACETIC ACHJ/a-KETOGLUTARATE-DEPENDENT DIOXYGENA SE
AND RELATED ENZYMES
By

Deborah Ann Hogan

The rapid degradation of the herbicide 2,4-dichlorophenoxyacetic acid (2,4-D) in
soils is mediated by bacteria containing the enzyme 2,4-D/Ot-ketoglutarate (cc—KG)
dioxygenase, or deA. deA is among the more than forty Fe(II)—dependent enzymes that
comprise the OL-KG-dependent dioxygenase superfamily. Most of these enzymes couple
the oxidative decarboxylation of OL-KG to diverse oxidations including hydroxylations,
cyclizations, desaturations, and ring expansion reactions. A small group of related
enzymes within the superfamily have been characterized through various biochemical,
spectroscopic and structural studies, but none of these sequences exhibit any overall
identity to deA or its relatives.

To better understand the mechanism of deA, site-directed mutagenesis
experiments were performed to identify the ligands to the Fe(II) metallocenter as well as
other residues that may be critical for activity. Nine histidines and one aspartate were
individually substituted with alanines, and the effects of the mutations were assessed by
kinetic and spectroscopic analyses. Electron paramagnetic resonance (EPR) and electron
spin-echo envelope modulation (ESEEM) spectroscopies were used to determine that
alteration of His 113 and Asp 115 caused dramatic changes to the metal binding site of

deA. A second histidine ligand was tentatively identified as His 262. Two residues, His

213 and His 216, were implicated in catalysis and the former may play a role in substrate
binding. i

In addition to serving as a model enzyme for mechanistic studies of Ot-KG-
dependent dioxygenases, deA performs a novel reaction in the degradation of aromatic
compounds. A search for other enzymes with deA-like activity found a hypothetical
protein, encoded by YLL057c, in Saccharomyces cerevisiae with 27% identity to deA.
All of the residues involved in deA metallocenter formation are conserved in this
sequence. To determine if this protein catalyzed a reaction similar to that of deA,
YLL057c was cloned and found to encode a sulfonate/OL-KG dioxygenase with activity
towards a number of different sulfonates including isethionate and taurocholate, a
conjugated bile salt. Studies with a S. cerevisiae strain which lacked this gene indicated
that sulfonate/Ot-KG dioxygenase is involved in utilization of sulfonates as alternative
sources of sulfur.

The presence of oc-KG-dependent dioxygenases with activities similar to deA
was also assessed in a collection of diverse soil bacteria. Thirty seven percent of the
seventy-seven isolates tested appeared to contain a tfdA-like element as determined by a
PCR-based method. Results from three different assays suggested that none of the
isolates could degrade 2,4-D. The high identity between tfdA-like genes from distantly
related bacteria suggests that extensive horizontal transfer was responsible for its
distribution. Although the function of {fdA-like elements in these bacterial isolates is not
known, the widespread presence of this gene implies its importance in the environment. It
is proposed that these genes are involved in the degradation of the aromatic compounds

prevalent in soils such as those released from lignin degradation.

 

To my family for their love and support

 

ACKNOWLEDGIVIENTS

My appreciation and thanks first go to my advisor, Bob Hausinger. His tireless
enthusiasm for being a both scientist and a mentor creates a wonderful environment for a
graduate student. Iparticularly value his dedication, directness, friendship and sense of
humor and I look forward to being his colleague. I would also like to thank my guidance
committee members, John Breznak, Pat Oriel, Cindy Nakatsu, and C. A. Reddy, for their
support. Each committee member had a unique influence on my graduate education. I
owe special thanks Cindy Nakatsu for sharing her expertise in molecular biology and her
interest in my project. I would also like to acknowledge the invaluable experience I
gained during my eight-month internship at Parke—Davis with Ruth VanBogelen, who is a
true bacterial physiologist and a great role model.

The supportive environment in the Hausinger Lab has been valuable to me. I
would particularly like to thank Gerry Colpas and Matt Ryle for their willingness to be
my spectroscopy gurus. I am very appreciative of the help given to me by Julie Dunning-
Hotopp, Linda Michel, and Ruth Saari. I can’t express enough gratitude to Tommy
Auchtung, who made major contributions to the yeast dioxygenase project, while
curating the lab aquarium and playing the Beatles. In addition, I would like to thank past
and present lab members (Tim Brayman, Aileen Soriano, Mary Beth Moncrief,
Padmakumar Raghavakaimal, Ruth Schaller and Vladimir Romanov) for their advice and

conversation.

Thanks to all of the members of my class who have been great friends ever since
our first post-exam celebration at the Peanut Barrel as students in Biochemistry 801.
Among the friends I have made here, I owe particular thanks to Mary Ellen Davey, Dan
Buckley, Joe Graber, Mark Johnson, Brad Stevenson, and Joel Klappenbach. I would also
like to mention my family and friends from home whose support from afar was essential.

I would like to thank the ROME Lab and John McCracken for use of space and
equipment. I have enjoyed support from the College of Natural Science, the
Biotechnology Training Program, and the Center for Microbial Ecology for which I am
grateful. Finally, I would like to express my gratitude to the departmental staff and

faculty whose assistance was important and appreciated.

vi

TABLE OF CONTENTS

List of Tables ...................................................................................................................... x
List of Figures .................................................................................................................... xi
List of Schemes ................................................................................................................ xiii
List of Equations .............................................................................................................. xiv
Key to Abbreviations ........................................................................................................ xv
CHAPTER 1 INTRODUCTION ....................................................................................... 1
Sequence analysis of a—KG-dependent dioxygenases ..................................................... 3
IPNS-like a-KG-dependent dioxygenases (Group I). .................................................. 5
Enzymes with similarity to deA (Group II) ............................................................... 7
Group HI oc-KG-dependent dioxygenases .................................................................. 10
Summary of oc-KG-dependent dioxygenase sequence analyses ................................ 11
Biochemical and biophysical characterization of oc-KG-dependent dioxygenases ....... l2
Biochemistry of deA and related enzymes ............................................................... 12
Biochemical studies of other a-KG-dependent dioxygenases ................................... l6
Site-directed mutagenesis and chemical modification studies of oc-KG-dependent
dioxygenases ......................................................................................................... l 9
Spectroscopic experiments for metallocenter characterization of a-KG-dependent
dioxygenases ......................................................................................................... 2 1
Crystal structures of IPNS and DAOCS .................................................................... 23
Mechanism of oc-KG-dependent dioxygenases .......................................................... 25
Degradation of 2,4-D and Ecology of deA .................................................................. 27
Overview of thesis ......................................................................................................... 33
Literature cited ............................................................................................................... 35

CHAPTER 2 CHARACTERIZATION OF THE ACTIVE SITE OF 2,4-
DICHLOROPHENOXYACETIC ACID/oc-KETOGLUTARATE DIOXYGENASE . 48

Introduction ................................................................................................................... 49
Experimental Procedures ............................................................................................... 51
Recombinant Plasmids ............................................................................................... 51
Protein Purification .................................................................................................... 53
Analysis of Kinetic Parameters .................................................................................. 54
Native Protein Analysis by Gel Filtration .................................................................. 55
Spectroscopic Analysis .............................................................................................. 55
Results ........................................................................................................................... 56
Production of the mutant deAs ................................................................................. 56
Kinetic analyses of altered deAs .............................................................................. 57
Evaluation of the Structural Consequences of the Mutations .................................... 58
EPR Spectroscopic Characterization of Variants with Altered Metal Sites ............... 59
vii

ESEEM Spectroscopic Characterization of Variants with Altered Metal Sites ......... 65
Discussion ...................................................................................................................... 74
Literature Cited .............................................................................................................. 81

CHAPTER 3 CLONING AND CHARACTERIZATION OF A SULFONATE/oc-

KETOGLUTARATE DIOXYGENASE FROM Saccharomyces cerevisiae ............... 85
Introduction ................................................................................................................... 86
Materials and Methods .................................................................................................. 87
Cloning and expression of YLL057c. ........................................................................ 87
Production and purification of YLL057c gene product. ............................................ 88
Characterization of MBP-dioxygenase fusion protein. .............................................. 89
Studies with the YLL057c deletion mutant ................................................................ 89
Results and Discussion .................................................................................................. 90
Additional properties of S. cerevisiae sulfonate dioxygenase .................................... 95
Phenotype of S. cerevisiae YLL05 7c deletion mutant ............................................... 95
Role of sulfonate/OL-KG dioxygenase in S. cerevisiae. .............................................. 97
Literature cited ............................................................................................................... 98
CHAPTER 4 DISTRIBUTION OF tfdA IN THE ENVIRONMENT ........................... 100
Introduction ................................................................................................................. 1 0 1
Materials and Methods ................................................................................................ 102
Bacterial strains ........................................................................................................ 102
16S rDNA partial sequence analyses ....................................................................... 103
PCR Amplification of the tfdA gene ........................................................................ 103
Dot Blot of PCR products ........................................................................................ 104
Analysis of 2,4-D degradation and deA activity .................................................... 105
Incorporation of radiolabel from l4C-2,4-D ............................................................. 105
Results ......................................................................................................................... 106
Composition of the LTER collection ....................................................................... 106
Identification of tfdA in LTER collection ................................................................ 107
2,4-D degradation by LTER isolates ........................................................................ 110
Experiments to determine if the y‘dA-like gene encodes a functional deA ............ 1 10
Discussion .................................................................................................................... 1 13
Acknowledgements. .................................................................................................... 1 16
Literature cited ............................................................................................................. 117
CHAPTER 5 CONCLUSIONS AND FUTURE RESEARCH ..................................... 121
Understanding the mechanism of oc-KG—dependent dioxygenases .......................... 122
Biological significance and potential applications of a-KG-dependent
dioxygenases ....................................................................................................... 1 29
Role of deA in the environment ............................................................................. 131
Literature cited ......................................................................................................... 137

viii

APPENDD( A AlVIINO ACID SEQUENCE ALIGNMENTS ....................................... 142
Literature cited ............................................................................................................. 151

APPENDDC B STEREOSPECIFIC DEGRADATION OF THE PHENOXY-
PROPIONATE HERBICIDE MECPROP .................................................................. 152

ix

 

LIST OF TABLES

Table 1. Characterized enzymes in the a-KG-dependent dioxygenase superfamily. ......... 4

Table 2. Conserved amino acids among subgroups within the oc-KG-dependent
dioxygenase superfamily. ............................................................................................. 6

Table 3. Amino acid sequence comparisons between a-KG-dependent dioxygenases ...... 8
Table 4. Sequences of mutagenic primers used to create altered tfdA genes ................... 52
Table 5. Summary of the kinetic parameters for active deA variants ............................. 58
Table 6. Summary of the EPR spectral parameters for Cu(II)-substituted deA variants.63
Table 7. Sulfonate specificity and kinetic parameters for sulfonate/a-KG dioxygenase. 93

Table 8. Distribution of isolates with tfdA among the different phylogenetic groups within
the LTER collection. ................................................................................................ 112

LIST OF FIGURES

Figure 1. Substrates of Group II enzymes. ....................................................................... 14
Figure 2. Oxyferryl mechanistic model for OL-KG-dependent dioxygenases .................... 26

Figure 3. Pathway for the degradation of 2,4-D by Ralstonia eutropha JMP134(pJ P4).. 28

Figure 4. Restriction maps of the plasmids used in construction of tfdA mutants. ........... 52
Figure 5. X-band CW-EPR spectra of Cu(II)-substituted deA variants. ........................ 60
Figure 6. 3-Pulse ESEEM spectra of Cu—substituted wild-type deA with 2-histidine
simulation data. .......................................................................................................... 67
Figure 7. 3-Pulse ESEEM time-domain spectra of deA variants .................................... 69
Figure 8. Spectral simulation of ESEEM data for D1 15A-Cu—deA ................................ 70
Figure 9. Spectral simulation of ESEEM data for MBP-H167A-Cu-deA. ..................... 71
Figure 10. 3—pulse ESEEM of D1 15A deA (4.2K). ........................................................ 73
Figure 11. Model for the active site of deA. ................................................................... 79
Figure 12. Structures of substrates for S. cerevisiae sulfonate/oc-KG dioxygenase .......... 91

Figure 13. Substrate concentration dependence of sulfonate/oc—KG dioxygenase

activity. ...................................................................................................................... 94
Figure 14. Role of YLL057c in the utilization of different sulfur sources. ...................... 96
Figure 15. 16S rDNA phylogeny of the LTER isolates .................................................. 108
Figure 16. Hybridization of amplified products from LTER isolates to a digoxigenin-

labeled fragment internal to the tfdA gene. .............................................................. 1 1 1
Figure 17. Comparison of the proposed reaction mechanisms for an a—KG-dependent

hydroxylase and IPNS. ............................................................................................. 125
Figure 18. Substrates of deA ......................................................................................... 132
Figure 19. Structures of the major interrnonomeric linkages in lignin. 132

xi

LIST OF FIGURES (CON ’T)

Figure 20. Open reading frames adjacent to a putative oc-KG-dependent' dioxygenase in

Bordetella pertussis. Genbank accession numbers are shown in parentheses. ........ 134
Figure 21. Scheme for lignin degradation in S. paucimobilis ......................................... 135
Figure 22. Alignment of Group H oc-KG—dependent dioxygenases ................................. 143
Figure 23. Alignment of Group III OL-KG-dependent dioxygenases .............................. 147
Figure 24. Alignment of complete and partial deA sequences ..................................... 149

xii

 

LIST OF SCHENIES

Scheme 1. Reaction catalyzed by deA. ............................................................................. 2
Scheme 2. Reactions catalyzed by IPNS, DAOCS and clavaminate synthase. ................ 18
Scheme 3. Reaction mechanism of S. cerevisiae sulfonate/or-KG dioxygenase ............... 91

xiii

LIST OF EQUATIONS

Equation 1 Equation for substrate inhibition .................................................................... 92

xiv

2,4—D
3-CBA

4-HPPD
a-KG
ACC
ACV
DAOCS
EDTA
EPR
ESEEM
GST
IPNS
LTER
MBP
deA

KEY TO ABBREVIATIONS

2,4-dichlorophenoxyacetic acid
3-chlorobenzoate

4-aminoantipyrene
4-hydroxyphenylyruvate dioxygenase
oc-ketoglutarate

aminocyclopropane carboxylate
L-oc-amino-S-adipoyl-L-cysteinyl-D-valine
deacetoxycephalosporin synthase
ethylenediaminetetraacetic acid
electron paramagnetic resonance
electron spin-echo envelope modulation
glutathione S-transferase

isopenicillin N synthase

long-term ecological research

maltose binding protein

2,4-D/oc-KG dioxygenase

XV

CHAPTER 1

INTRODUCTION

2,4-Dichlorophenoxyacetic acid (2,4-D)/cx-ketoglutarate (on-KG) dioxygenase
(deA) encodes the first enzyme in a well—characterized degradative pathway for the
herbicide 2,4-D by bacteria (35). deA is a non-heme F e(II)-dependent enzyme that
couples the oxidative cleavage of OL-KG, yielding C02 and succinate, to the
decomposition of 2,4-D to form 2,4-dichlorophenol and glyoxalate (26) (Scheme 1).
Based on its requirement for Fe(II) and its use of oc-KG as a cosubstrate, deA is included

in the oc-KG-dependent-dioxygenase superfamily (90).

O/\COO' /lOH
‘000
c: 0 \COO' OH coo
O Fez. J\ Cl spontaneous C1 + (IZHO + + C0
+ + 02 ——> I \ —————> 000' 2
Cl / '00C
'000
Cl (:1

Scheme 1. Reaction catalyzed by deA.

In this introduction, 1 first compare the amino acid sequences of members of the
oc-KG-dependent dioxygenase superfamily to indicate relationships and differences
between these enzymes. Next, I summarize previous research on deA and other
biochemically characterized a-KG-dependent dioxygenases. Finally, I review studies on
the occurrence of the tfdA gene, which encodes deA, in various 2,4-D degrading soil
bacterial isolates and communities. The introductory chapter will conclude with an
overview of the research presented in this dissertation.

oc-KG-dependent dioxygenases are common in plants, animals, and the majority

of aerobic microorganisms. Furthermore, data from recent genome sequence projects

1‘.)

 

suggest their presence in nematodes and insects (y-butyrobetaine hydroxylase in
Caenorhabditis elegans, Genbank # 266523 and Drosophila melanogaster, Genbank #
AL109630, respectively). Since 1967, when prolyl hydroxylase was characterized as the
first a-KG-dependent dioxygenase (45), this class of enzymes has come to include more
than forty members (Table 1). The reactions catalyzed by these enzymes range from steps
in the synthesis of hormones (36, 91), antibiotics (58), collagen (54), and compatible
solutes (30, 44) to the decomposition of pyrimidines (1 1 l, 115), amino acids (21) and
other small molecules (21, 27, 40). In light of the diversity of substrates for a-KG
dependent dioxygenases and the different types of oxidative reactions they catalyze, it is

not surprising that their sequences exhibit little overall similarity.

Amino acid sequence analysis of a-KG—dependent dioxygenases

Although there is no overall sequence identity uniting all enzymes in the a-KG-
dependent dioxygenase superfamily, comparison of their primary structures finds
subgroups of related proteins. More than twenty distinct enzymes with significant levels
of sequence identity to isopenicillin-N—synthase (IPNS) (Table 1) have been described (8,
58). The IPNS-like enzymes (Group I) include the majority of characterized oc-KG-
dependent dioxygenases. A second cluster of enzymes related by sequence (Group H) is
comprised of deA, four previously characterized oc-KG-dependent dioxygenases, and

four putative enzymes yet to be studied. The Group II enzymes do not exhibit significant

 

Table 1. Characterized enzymes in the a—KG—dependent dioxygenase superfamily.

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Enzyme Source for purified References
enzyme
Group I IPNS-like enzymes
Iscmenecillin N synthase (IPNS)a Fungi, Bacteria (93)
Deacetoxycephalosporin C synthetase (DAOCS) Fungi, Bacteria (56)
Deacetylcephalosporin C synthase Fungi, Bacteria (102)
Aminocyclopropane carboxylate (ACC) oxidasea Plant (135)
Ethylene-forming enzyme Bacteria (83)
Deacetoxyvindoline 4-hydrmglase Plant (14)
Hyocyamine-6S—hydroxylase Plant (69)
F lavone synthase I Plant (9)
Flavanol synthase Plant (43)
F lavanone 3-hydroxylase Plant (10)
Leucoanthocyanidin hydroxylase Plant (99)
Gibberellin-ZB-hydroxylase Plant (37) for review
Gibberellin 3B-hydroxylase Plant "
Gibberellin-7-oxidase Plant "
Gibberellin l3-oxidase Plant "
Gibberellin-ZO-oxidase Plant "
Procollagen Prolyl-4—Hydroxylase Mammal, Bird, (54) for review, (23)
Virus
Procollagen Aspartyl Hydroxylase Mammal (55, 129)
Procollagen Lysyl Hydroxylase Mammal (74, 81, 89, 125)
Group II deA-like enzymes
deA Bacteria (27)
Taurine/oc-KG dioxygenase (TauD) Bacteria (21)
Sulfonate/a—KG dioxygenase Fungi (40)
Clavaminate synthase (CS) Bacteria (11)
y-Butyrobetaine hydroxylase Bacteria, Mammal (30, 62)
Group III
Proline-4-Hydroxylase Bacteria (1 O6)
Phytanoyl-COA Hydroxylase Mammal (46)
Miscellaneous
Thymine hydroxylaseb Fun& (1 15)
Pyrimidine-deoxynucleoside- 1 dioxygnaseb Fungi (1 l 1 )
Trirnethyl-lysine dioxygenaseb Mammal (44)
Proline 3-hydroxylase° Bacteria (77)

 

 

a Enzyme does not use a—KG as a cosubstrate
bNo sequence is available for this enzyme
° Enzyme does not have any sequence identity to other characterized enzymes

 

similarity to IPNS and other Group I enzymes. Group IH contains four enzymes that,
despite their very disparate substrates, have obvious sequence relatedness. Future studies
on the structure and activity of enzymes in this class may uncover a relationship between
Group III and either of the other two groups of a-KG-dependent dioxygenases; at
present, however, sequence comparisons predict that they form a distinct cluster. Table 2
summarizes the elements common to the enzymes in each group. Finally, a fourth group
contains or-KG-dependent dioxygenases that either exhibit no sequence similarity to

enzymes in the other three classes or their amino acid sequence is not yet published.

IPNS-like a—KG-dependent dioxygenases (Group I)

IPNS is an Fe(II)—dependent enzyme that converts L-oc-amino-B-adipoyl-L-
cysteinyl-D-valine (ACV) to isopenicillin N (Scheme 2) with chemistry that is similar to
the mechanism of a-KG-dependent dioxygenases (although a—KG is not used as a co-
substrate). IPNS has between 25 and 35% amino acid identity to over twenty enzymes
involved in antibiotic synthesis (58) and the formation of plant hormones (91). For
comparison, sequences for the same enzyme from different plant genera have, on
average, more than 60% amino acid identity. Alignment of Group I sequences indicates
that a His-X—Asp-Xss-His motif is found in all of these enzymes (8) (Table 2 A). Crystal
structures of IPNS (94) and the related enzyme deacetoxycephalosporin C synthase
(DAOCS) (122) supported biophysical studies that predicted these residues to be ligands
to the iron metallocenter. These same structures, combined with further sequence
comparisons (58, 63), reveal a conserved Arg-X-Ser arrangement involved in oc-KG

binding in Group I enzymes.

P4
Lys

 

B. Group H

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

H D x DN H x R
deA WHSDSSFQ 147 WNVGDLVMWDNRCVLH 10 R
TauD WHTDVTFI 153 WQPNDIAIWDNRVTQH 10 R
YSD WHTDVVSY 147 WEPHSVVIWDNRRVQH 11 R
AtsK WHTDVTFV 134 WEAGDVAIWDNRATQH 10 R
Pch LHWDGMYL 147 WRSDDLVIADNLTLLH 10 R
CarC FHADGGLL 148 WEDGDLLIMDNRRVIH 11 R
SyrI YHNESSHL 192 WRKGDVVMLDNMLAAH 8 R
y-BBH LHTDLPTR 139 LEAGQLWCFDNRRVLH 11 R
cs FHTEMAYH 123 LEPGDLLIVDNFRTTH 11 K x R
C. III

H N R
HGSGTNMSPHPR
H TIRR

HGSGQNKTQGFR
RPR

 

Table 2. Conserved amino acids among subgroups within the a-KG-dependent
dioxygenase superfamily.

Amino acids in common among the IPNS-like a-KG—dependent dioxygenases (A), the
deA-like enzymes (B) and the proline 4-hydroxylase group (C) in the a—KG—dependent
dioxygenase superfamily are shown. The numbers in columns labeled “X” represent the
number of amino acids intervening between the conserved regions. The unique identifiers
for the protein sequences in Group I are IPNS, isopenicillin N synthase (IPNS_STRJU);
DAOCS, Deacetoxycephalosporin synthase (pir2:A39204); FL3H, F lavanone 3-
hydroxylase (SW:FL3H_HORVU); VanB, vanillate biosynthesis enzyme B (AF 009673);
P4H, human prolyl 4-hydroxylase (P13674); LysH, human lysyl hydroxylase (Q02809).
The sequences in Group H are deA, 2,4-D/oc-KG dioxygenase (M16730); TauD,
taurine/a-KG dioxygenase (P37610); YSD, S. cerevisae sulfonate/oc-KG dioxygenase
(Q12358); AtsK, putative oc-KG dioxygenase from sulfate-ester transport gene cluster
(AF 126201) Pch, pyoverdine biosynthesis enzyme (AF 002222); CarC, carbapenem
biosynthesis enzyme C (AAD38231); SyrI, syringomycin biosynthesis enzyme I
(AAB63253); y—BBH, y-butyrobetaine hydroxylase (P80193); CS, clavaminate synthase
(A44241). Group III sequences are Pro4H, proline 4—hydroxylase (D78338); HtxA,
putative hypophosphite/oc—KG dioxygenase (AF023463); PPHYH, Phytanoyl-CoA
hydroxylase (NP_006205); MmcH, mitomycin C biosynthesis protein (AF 127374).

Several enzymes involved in the hydroxylation of peptidyl amino acids are
grouped together by activity and placed into Group I based on the presence of the above
motif. Although these key active site residues are maintained in prolyl-4—hydroxylase (15,
23), lysyl hydroxylase (74, 81, 125), and aspartyl hydroxylase (47), their sequences lack
detectable levels of similarity to other Group I enzymes. The importance of the amino
acids in the above motif for activity has been confirmed by mutagenesis studies
(described below). 4—Hydroxyphenylpyruvate dioxygenase (4-I-IPPD) is an Fe(H)-
dependent enzyme that catalyzes ring hydroxylation concomitant with decarboxylation of
the a-keto acid side chain. Mechanistic similarities in addition to seeming conservation
of the above motif prompted inclusion of 4-I-IPPD with Group I enzymes of or-KG
dependent dioxygenase superfamily. Upon introduction of less related 4-I-IPPD sequences
into alignments, the H—X—D motif was no longer absolutely conserved, casting doubt on
the relationship of this enzyme to the a-KG-deperrdent dioxygenases. A recently released
crystal structure of 4-HPPD indicates a protein fold more similar to that of extradiol

(ring-cleaving) dioxygenases than H’NS and DAOCS.

Enzymes with similarity to T fdA (Group 11)

As recently as 1995, no enzymes with significant sequence similarity to deA
were known. Over the past five years, however, multiple deA homologs have been
identified due to the dramatic increase in genome sequence data and the development of
more powerful sequence analysis tools, such as BLASTP (l) and Position-Specific

Iterative (PSD-BLAST (2). Now, it is apparent that deA is related to four characterized

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oc-KG-dependent dioxygenases and to sequences of several putative a-KG-dependent
dioxygenases. Amino acid sequence identities from pairwise comparisons range from
fifteen to forty percent (Table 3).

deA is most closely related to taurine/OL-KG dioxygenase (TauD) (21),
Saccharomyces cerevisiae sulfonate/oc-KG dioxygenase (YSD) (40) and three putative Ot-
KG—dependent dioxygenases: Pch, involved in synthesis of the siderophore pyoverdine
(109), CarC, which is necessary for formation of the B—lactamase inhibitor carbapenem
(72), and AtsK, a putative dioxygenase in a sulfate-ester degradation gene cluster in
Pseudomonas putida (127). Although they have not been biochemically characterized,
the roles of Pch, CarC and AtsK were identified through various transposon
mutagenesis studies. The relationships between these six sequences are detected by a
standard BLASTP analysis. More sensitive analyses with PSI-BLAST find relationships
between deA and more distantly related a-KG-dependent dioxygenases. PSI-BLAST
detects lower levels of relatedness by creating a position-specific matrix of conserved
amino acids to rescore and realign sequences that receive low BLAST scores. As this
process is repeated, unrelated sequences with chance high scores are downgraded as the
matrix of positionally-conserved amino acids improves. By these methods, relationships
between deA and two other well-characterized oc-KG-dependent dioxygenases, y-
butyrobetaine hydroxylase and clavaminate synthase (Table 3), become evident. In
addition, the sequence of a third enzyme, SyrB, which is required for synthesis of
syringomycin, also aligns with the deA-like sequences. Of the twenty nine sequences

identified as relatives of deA after six iterations of PSI—BLAST, all were either

 

previously characterized oc-KG dependent dioxygenases or hypothetical proteins with the
characteristic conserved amino acids described below.

Comparison of all nine Group H a-KG-dependent dioxygenase sequences finds
several strictly conserved residues (Table 2B, Appendix A). As in the Group I enzymes,
an H-X-D-like motif is present in all sequences, although in both clavaminate synthase
and SyrB, the aspartate is replaced by a chemically-equivalent glutamate residue. A
stretch of amino acids near the C-terminus with high levels of similarity across these
sequences, contains the only other invariant histidine, suggesting that this residue may be
analogous to the third metal ligand in the Group I motif described above. Conserved
asparagine and arginine residues near this histidine are also apparent and the latter may

play a role similar to the a-KG-binding arginine in IPNS and DAOCS.

Group III a-KG-dependent dioxygenases

A third group within the oc—KG-dependent dioxygenase superfamily contains two
characterized enzymes, proline 4-hydroxylase (73) and phytanoyl-CoA hydroxylase (46),
and two putative a-KG-dioxygenases, hypophosphite oxidase (HtxA) (73) and MmcH
which is involved in mitomycin C biosynthesis (67). Although no homology is detected
between these sequences and either Group I or Group II or-KG-dependent dioxygenases
by BLAST comparisons, the motif common to this group is clearly similar to those
described above (Table 2C). Again, there is a conserved H-X-D motif and two additional
invariant histidines in this small set of sequences (Appendix A) and in uncharacterized
homologs in other bacteria (120). A His-Xlo-R motif (similar to Group I and Group H

enzymes) identifies a good candidate for the third metal ligand; however, biochemical

10

 

studies or structural information is needed to establish the roles of these residues. In
humans, non-functional phytanoyl-CoA hydroxylase results in Refsum disease due to the
inability to break down phytanic acid (3,7,11,15-tetramethylhexadecanoic acid), a
degradation product of chlorophyll (12). It is interesting to note that two of the disease-
causing alleles of phytanoyl-CoA hydroxylase, which encode inactive enzyme, have

mutations of either the conserved arginine or an invariant asparagine nearby.

Summary of a-KG-dependent dioxygenase sequence analyses

Sequence comparisons of members of the or-KG-dependent dioxygenase
superfamily find three distinct subgroups of related enzymes; no overall sequence
identity was detected between enzymes from different subgroups. Comparison of more
closely related sequences allows for the identification of potentially important, conserved
residues. The consensus motifs for the three subgroups are similar (an H-X-D motif
distant from an H-Xlo-R arrangement) and likely identify residues important for active
site structure. As the number of oc-KG-dependent dioxygenase sequences grows, the three
groups may merge into a continuum of sequences. Alternatively, as other sequences are
found, distinctions between the groups may be maintained. Differences in the
arrangement of the conserved residues across the superfamily may be indicative of the

mechanisms that drove the divergent evolution of these enzymes.

11

Biochemical and biophysical characterization of a-KG-dependent dioxygenases

Biochemistry of T fdA and related enzymes

deA was detected in a 2,4—D-degrading Arthrobacter sp. in 1967 (l 16).
Preliminary studies on the degradation of 18O-ether—labeled 2,4—D found the products to
be 18O-2,4—dichlorophenol and glyoxalate. Thus, it was proposed that the 2,4—D
decomposition proceeds through a hemiacetal intermediate that spontaneously breaks
down to yield the reaction products (Scheme 1). Almost twenty—five years later, the
mechanism of purified deA was examined (27), proving this enzyme to be an or-KG-
dependent dioxygenase.

Various characteristics of recombinant deA, purified from E. coli cells
containing the Ralstonia eutropha JMP134 tfdA gene, have been elucidated over the past
five years. deA, with a predicted subunit molecular weight of 31.7 kDa has an apparent
molecular weight of 50 :r 2.5 kDa by gel filtration chromatography suggesting that the
native protein exists as a homodimer. The high affinity for or-KG is demonstrated by a
Km of 3.2 i 0.54 uM. Several different oc-keto—acids, including oc-ketoadipate and
pyruvate, are less efficient substitutes for oc-KG as the cosubstrate. The km and Km for
2,4-D are 529 i 16 min'] and 17.5 uM, respectively. 2,4—D can be replaced with other
aromatic compounds such as phenoxyacetic acids (27), substituted cinnamic acids,
naphthoxyacetic acid and benzofurancarboxylic acid (R. Padmakumar and J. Hotopp,
unpublished data). deA enzymes from both R. eutropha JMP134 and Burkholderia
cepacia RASC (1 12) selectively abstract the pro-R hydrogen atom from the prochiral

side-chain carbon adjacent to the ether bond of 2,4—D (98). Interestingly, there is evidence

of a deA-like oc-KG-dependent dioxygenase that has the opposite stereoselectivity from
the deAs described above (98, 114), however, this enzyme has not been cloned or
characterized (Appendix B). Distinct oc-KG-dependent dioxygenases in Sphingomonas
herbicidovorans act on each enantionmer of mecoprop ((R,S)-2-(4—chloro-2-
methylphenoxy)—propionic acid), a chiral analog of 2,4—D (84).

Three characterized enzymes with similarity to deA include TauD, which
releases sulfite from taurine, y-butyrobetaine hydroxylase, which is responsible for the
final step in camitine biosynthesis, and clavaminate synthase, which catalyzes three
distinct transformations in the biosynthesis of the B-lactamase inhibitor clavulinic acid.
The diverse substrates of these enzymes are compared in Figure 1. While all four
enzymes have approximately the same apparent KD for iron, other kinetic parameters vary
significantly. The Km for oc-KG ranges from 3 uM for deA to 260 uM for clavaminate
synthase. deA and TauD are the only enzymes that can substitute other oc-keto acids for
OL-KG. The kcm/Km (min'l 11M “1) is 31.1, 2.0, 0.3, 0.013 for deA, TauD, y-butyrobetaine
hydroxylase, and clavaminate synthase, respectively. The wide range of catalytic
efficiencies likely reflects the diversity of substrates. In each case, the hydroxylated
carbon is one or two carbons away from either a carboxylate or sulfonate group
indicating that a positively charged amino acid might be involved in substrate binding.
The pro-R hydrogens abstracted by y-butyrobetaine hydroxylase (22) and clavaminate
synthase (5) are in similar relative positions to the carboxylate moiety, which is two
carbons away from the hydroxylation site. The pro-S hydrogen removed by deA
(98) is in the opposite position with respect to the carboxylate. The stereochemistry of

TauD catalysis has not been examined.

13

2,4-D taurine

 

 

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COOH

y-butyrobetaine deoxyguanidinoproclavaminic acid

Figure 1. Substrates of deA-like enzymes

Structures of substrates for deA, TauD, y-butyrobetaine hydroxylase, and clavaminate
synthase are compared. The abstracted proton is indicated by an arrow.

14

 

 

For maximal activity, many a-KG-dependent dioxygenases require ascorbate to
reduce the iron after non-productive decarboxylation of a-KG that is uncoupled from
oxidation of substrate. Although deA activity is stimulated by the presence of ascorbate
- in the reaction mix (27), deA does not appear to undergo uncoupled turnover of oc-KG
in the absence of substrate. Even with the addition of ascorbate, deA is inactivated in a
matter of minutes in the presence of iron under oxic conditions. Inactivation of deA
appears to occur by both turnover—dependent and a tumover-independent mechanisms
(97). TauD activity is also stimulated by 50% in the presence of ascorbate at
concentrations over 200 uM. y-Butyrobetaine hydroxylase activity is stimulated six-fold
by the addition of 100 M potassium ions; the Km for both substrates is also reduced in
the presence of potassium (130). Without potassium present, y—butyrobetaine hydroxylase
performs the uncoupled decarboxylation of oc-KG. Higher rates of uncoupling are
achieved by the addition of product and its enantiorner (42). Clavaminate synthase also
catalyzes the uncoupled decarboxylation of oc-KG, but the rate is greatly reduced (1 in
890 turnovers) in the presence of substrate (60).

Beyond the differences in kinetic parameters and activity towards a-KG in the
absence of substrate, other characteristics of these enzymes can be compared. All four
enzymes have a pH optimum of 6.5-7. TauD (subunit size of 37.4 kDa) and y-
butyrobetaine hydroxylase, (43 kDa) both exist as homodimers (21, 62). In contrast, gel
filtration chromatography of clavaminate synthase gives an estimated mass of 36 kDa
indicating that the enzyme is not rnultimeric. Clavaminate synthase is unique in that it
catalyzes hydroxylation, desaturation, and cyclization reactions (Scheme 2), all coupled

to oc-KG decarboxylation, at the same active site (11, 60). Kinetic studies suggest that

15

there is no release of the substrate between the desaturation and cyclization reaction. It is
interesting to recognize the similarities between the substrates of clavaminate synthase
and those of IPNS and DAOCS. Despite a lack of sequence similarity, and all three
enzymes participate in the synthesis of B-lactam antibiotics. In addition, a poorly
characterized enzyme, CarC, which shows high levels of identity to deA over the entire
sequence, is involved in formation of carbapenem, another B-lactam-containing

compound (Scheme 2).

Biochemical studies of other a-KG-dependent dioxygenases

There is significant diversity in the subunit size and quaternary structure within
the oc-KG-dependent dioxygenase superfamily. Among the enzymes related to IPNS,
most are between 38 and 45 kDa (similar in size to deA-like enzymes) and many
(flavone synthase (9), gibberellin 3B-hydroxylase (3 7), gibberellin 20-oxidase (100),
hyoscyamine (34)), but not all (flavanone 3-hydroxylase (10), ACC oxidase (49),
thymine hydroxylase (115)), of the enzymes in this subgroup exist in a monomeric state.
Crystal structure information suggests that apo-DAOCS exists as a trimer but dissociates
to a monomer upon metal binding (122). The three mammalian enzymes involved in
collagen maturation, aspartyl hydroxylase, lysyl hydroxylase and prolyl hydroxylase,
show additional differences from other a-KG-dependent dioxygenases. Aspartyl
hydroxylase has a subunit size of 85 kDa, but a 52 kDa catalytic domain still has full
catalytic activity (47). Lysyl hydroxylase has been identified as an 87 kDa monomer in
chick embyros and bovine skin, and a dimer of 70 and 110 kDa subunits in porcine skin.

Prolyl hydroxylase, with a subunit size of 61 kDa, forms an 01202 heterodimer in

16

mammals and an 043 dimer in C. elegans (54). In both cases, the [3 subunit is protein
disulfide isomerase. In all three enzymes, the C-terminal catalytic domain appears to be
linked to a separate N-terminal region involved in cellular localization (52). Furthermore,
a recently discovered prolyl 4-hydroxylase from Paramecium bursaria Chorella virus- 1,
an algal virus, that is 20% identical to the C-terminus of mammalian prolyl 4-hydroxylase
lacks the N-terminal localization domain; thus, it’s subunit size is closer to that of
bacterial and fungal a-KG-dependent dioxygenases (23).

Almost all a—KG-dependent dioxygenases have an apparent KD for iron of
approximately 2 uM. The Km for oc-KG ranges from 5 uM to between 140 uM for
different enzymes. The Km for 02 has been determined to be 30-40 uM for prolyl
hydroxylase (82), 60 uM for thymine hydroxylase (41), and 45 uM for
desacetoxyvindoline 4-hydroxylase (14). Not surprisingly, the measured Km values for
the unique substrates vary from nanomolar to near milimolar concentrations.

Substrate interaction kinetics and product inhibition studies in four a-KG-dependent
dioxygenases, desacetoxyvindoline 4-hydroxylase (l4), prolyl hydroxylase (82), lysyl
hydroxylase (92), and thymine hydroxylase (41), all indicate an ordered ter-ter
mechanism for substrate binding. Competitive inhibition studies indicated that in three of
the four enzymes, oc-KG binds first, followed by the appropriate substrate, then 02. In
prolyl hydroxylase, the order of Oz and substrate binding are reversed (82). The oxidized
product, C02, and succinate are then released sequentially in that order. The unproductive

decarboxylation of a-KG in the absence of substrates provides additional evidence that

oc-KG can bind without the substrate present.

17

 

  

 

lPN

S H
Fe(ll) HOOC N
m NH2 0 O N

 

02 21120 ' COOH

H H
HOOC N HOOC N S
DAOCS
Fe(II) O
NH; 0 O N 7:» NH2 0 N /
0; H20 COOH
aKG succinate
(‘03
NH CS
A Fe(ll) HO NH
N A
N NH
O N NHZ
COOH O ' H
2 succmate
aKG (:02 COOH
HO CS ..... ““110
Fe(ll) H
N N
0 NH; 7? o ”“2
COOH 02 H20 COOH
aKG succinate
C02
..... ““110 ,,_....u\\O
H CS
N Fe(ll) -——
0% NH2 7?» N NH?
0
COOH
02 ”20 COOH
uKG succinate
C02
CarC
acetyl-CoA N /
————> ——>
y-glutamyl phosphate 0
COOH

Scheme 2. Reactions catalyzed by IPN S, DAOCS, clavaminate synthase

and CarC.

Although in deA and related enzymes Ot-KG decarboxylation is relatively tightly
coupled to substrate oxidation, prolyl 4—hydroxylase and lysyl hydroxylase (80) have a
near absolute requirement for ascorbate. Early spectroscopic studies with Fe(H)-prolyl
hydroxylase demonstrated that ascorbate serves to reduce the iron metallocenter which
becomes oxidized to an Fe(HI) form upon brief incubation with a—KG. Ascorbate is
consumed stoichiometrically with respect to uncoupled decarboxylation (80). Catalysis is
not as dependent on the presence of ascorbate in bacterial proline hydroxylases (78).
Studies with analogs of oc-KG and ascorbate suggest that both occupy a similar or
overlapping site in prolyl 4—hydroxylase and both compounds chelate the metal (65).

In addition to IPNS, aminocyclopropane carboxylate (ACC) oxidase does not
require or-KG for activity. ACC oxidase, responsible for the generation of ethylene in
plants, requires Fe(II) and C02 (49). The role for CO; has yet to be determined, but it has
been suggested that bicarbonate, a carbamylated lysine, or CO; itself may ligate the iron
(49). Stoichiometric amounts of ascorbate are consumed during the course of the

reaction, perhaps suggesting that ascorbate replaces a-KG.

Site-directed mutagenesis and chemical modification studies of a—KG-dependent
dioxygenases

Mutagenesis studies of several enzymes within Group I of the oc-KG-dependent
dioxygenase superfamily have confirmed the importance of the conserved residues (H-X-
D-Xss-H-Xlo-R). In IPNS, replacement of His 214, Asp 216 or His 272 with leucine
completely inactivates the enzyme (1 13). The crystal structure of IPNS proves these

residues to be metallocenter ligands (94, 95). Although a fourth metal ligand, Gln 330, is

19

evident in the substrate-free structure, this residue is neither conserved nor critical for
activity (101). The importance of the corresponding residues in flavanone 3B-
hydroxylase, which is 23% identical to IPNS over the entire sequence, has also been
demonstrated (63). Alteration of His 220 or Asp 222 to glutamine causes a greater than
99% loss of activity but the resultant enzymes only show a modest increase in the
apparent KD for iron. Mutation of His 278 to glutamine completely abolished activity,
and mutation of a conserved Arg in position 288 to lysine or glutamine corresponded to
decreased affinity for a-KG (63).

Additional studies have been carried out with the enzymes that hydroxylate
peptidyl substrates. Mutation of His 656, Asp 658, and His 708 rendered lysyl
hydroxylase inactive (88). Likewise, alteration of the second conserved histidine in
aspartate hydroxylase (His 675) completely abolishes activity (70) and appears to cause a
decrease in metal and a—KG binding (47). Replacement of His 675 in aspartate
hydroxylase with iron-binding amino acids restores partial activity (70). In prolyl
hydroxlase, alteration His412, Asp 416, His483 and Lys493 in the human-derived
enzyme had significant effects on activity. Prolyl hydroxylase variants in which either of
the histidines was mutated were inactive regardless of the substitution (59). Chemically-
equivalent substitutions of remaining conserved residues, Glu for Asp 414 and Arg for
Lys 493, cause an increase in the Km for iron and oc-KG, respectively. Mutation of a
fourth histidine in prolyl hydroxylase, His 501, leads to a 95% loss of activity, a small
increase in Km for a-KG, and an increase in the rate of uncoupled oc-KG decarboxylation

suggesting that this residue is also present at or near the active site (79).

20

Together, these studies conclusively show the importance of histidines and
carboxylates in the creation of the reactive metallocenter of or-KG-dependent
dioxygenases. In most enzymes, no alternative amino acids (Arg, Asp, Glu, Gln, Ala) can
substitute for a metal-binding histidine, although in certain instances, low levels of
activity can be seen with a His to Gln substitution. Alteration of the conserved aspartate
to even the very similar glutamate cannot always retain activity, yet in clavaminate
synthase and the sequence of SyrB, glutamate replaces aspartate in the metal binding
motif. In the cases presented above, the conserved residues are identified through
sequence comparisons to IPNS and related enzymes. To date, no mutagenesis studies on

enzymes without similarity to IPNS have been published.

Spectroscopic experiments for metallocenter characterization of a—KG-dependent
dioxygenases

Early insight into the metallocenter structure of enzymes in the OL-KG-dependent
dioxygenase superfamily came from spectroscopic studies of IPNS. Electron
paramagnetic resonance (EPR) spectra of inactive Cu(11)- and Co(II)—substituted IPNS,
used to model the EPR silent Fe(H) form of the enzyme, are characteristic of a five— or
six-coordinate tetragonally distorted site (75). Electron spin-echo envelope modulation
(ESEEM) spectroscopic data are indicative of two equatorial histidines to the Cu(II) form
of the enzyme, one of which is displaced upon substrate binding (48). Extended X-ray
absorption fine structure (EXAFS) (105) spectroscopy demonstrates the presence of a

thiolate ligand to the metal upon substrate binding. Site-directed mutagenesis of cysteine

21

 

 

residues ruled out a potential role for endogenous S-containing ligands thus indicating
that the metal-bound sulfur is derived from substrate (85).

Spectroscopic studies of three Group H oc-KG-dependent-dioxygenases,
clavaminate synthase, deA and TauD, demonstrate common features among these three
enzymes as weH as similarity to IPNS. Again, initial EPR characterization was performed
on the inactive Cu(H)-substituted form of the enzyme. Structural information on the
Cu(H)-metallocenter is likely relevant to the Fe(H) conformation as Cu(H) binds
competitively with respect to iron with an approximate K, of 1-3 uM (131). Later studies,

using F e(H)— nitric oxide (NO) complexes to derive an EPR signal, were also performed
on deA (39). In the absence of substrates, EPR spectra of both Fe(H) and Cu(H)-
substituted deA indicate the presence of two histidines bound to the metal. ESEEM
studies of Cu(H)-deA agree with the EPR data, and also identify three metal-bound
waters in the remaining coordination sites. Addition of 2,4-D to the enzyme does not
affect the coordination geometry, but binding of a-KG decreases the rhombicity of the
metallocenter and displaces two solvent molecules indicating direct interaction between
oc-KG and the metal. ESEEM studies of the Cu(H)-oc-KG—2,4—D deA ternary complex
indicated the loss or shift of a histidine ligand from the equatorial plane. Studies with the
F eCH) deA are consistent with the rearrangement of ligands and loss of the third water
upon substrate binding to give a five-coordinate metal center. Nitric oxide, which also
serves as an oxygen analog, binds reversibly to Fe-deA and F e-or-KG-deA complexes,
but shows a greatly increased affinity for the metal when both substrates are present.
The interaction between oc-KG and the metal was studied in detail using UV-

visible, magnetic circular dichroism (MCD) and circular dichroism (CD) spectroscopies

22

with clavaminate synthase. These techniques revealed that upon binding of a—KG to
clavaminate synthase, the six-coordinate geometry is distorted. The formation of a pink
intermediate arising from ligand to metal charge transfer between oc-KG and the iron
indicates that oc-KG binds to the metal through both the C-1 carboxylate and the C-2 keto
group based on comparisons to model compounds (87). A similar mode for binding of Ot-

KG to deA and TauD is predicted from the results of recent experiments (39, 96).

Crystal structures of IPNS and DA OCS

IPNS has been crystallized in the presence of Mn(H) and Fe(H) (94, 95). Both
structures show that the metallocenter is ligated by His 214, Asp 216, His 270, and Gln
330. The active site is buried within the hydrophobic pocket of an eight-strand “jelly-roll”
motif. In the F e(H)-IPNS structure with substrate bound, Gln 330 and one water is
displaced upon addition of the tripeptide substrate, ACV. In the Fe-ACV-IPNS complex,
the iron is five-coordinate which agrees with the geometry observed in the spectroscopic
experiments described above. Furthermore, nitric oxide binds to the open axial
coordination site forming an octahedral complex as revealed by the crystallographic
studies.

The crystal structure of DAOCS is the only published structure of an oc-KG
dependent dioxygenase. The anaerobic addition of F e(H) to crystals of apo-DAOCS leads
to only small structural changes. In DAOCS, ferrous ion is ligated by residues
corresponding to the metal ligands in IPNS, and three water molecules. As predicted by
the spectroscopic experiments described above, a-KG displaces two solvent molecules to

chelate the iron. The C-l carboxylate binds opposite to His 243 (the distal histidine of the

23

H-X-D-Xss-H motif) and the C-2 keto group binds trans to the metal-bound aspartate.
The only accessible site, located in a hydrophobic pocket, is occupied by a solvent
molecule.

The crystal structures of IPNS and DAOCS lend very strong support for the
models of a-KG—dependent dioxygenase metallocenter structure as deduced from
sequence alignments, site-directed mutagenesis studies, and spectroscopic experiments.
Although the substrate-free configuration of the two rnetallocenters are similar, the
differences in the chemistry performed by these two enzymes, particularly the use of or—
KG as a co-substrate in DAOCS and not in IPNS, is reflected in the differences in ligands
and coordination geometries upon substrate binding. There is no available structure of an
a-KG-dependent dioxygenase that performs hydroxylation chemistry.

Crystal structures for two additional a-KG-dependent dioxygenases, clavaminate
synthase (Group H) and proline 4-hydroxylase (Group IH) will soon accompany the
structures of IPNS and DAOCS as described in a seminar by Prof. C. Schofield. Despite
the distinct differences between the three groups of sequences, these enzymes appear to
fold on a similar scaffold. This exciting finding will allow us to better assimilate the large
amount of spectroscopic, stereochemical, and biochemical data into a universal model for
enzymes in this superfamily. Within the subgroups, information from one crystal
structure will greatly aid in the prediction of substrate binding residues and potential
catalytic residues in other related enzymes.

A 2-His-1-carboxylate motif is also found in other non-heme, ferrous ion
dependent enzymes such as lipoxygenase, extradiol dioxygenase, and pterin-dependent

tyrosine hydroxylase (38). These three ligands anchor the iron in the active sites, and, as

24

demonstrated by site-directed mutagenesis studies, are clearly important for establishing
the proper geometry and reactivity for interaction with substrates. Thus, the three
remaining coordination sites provide the opportunity for substrates to interact directly

with the iron to modulate activity.

Mechanism of a-KG-dependent dioxygenases

a-KG-dependent dioxygenases perform a wide range of oxidative reactions
including hydroxylations, cyclizations, ring expansions and desaturations using or—KG as
a reductant of diatomic oxygen. Despite the diversity of substrates and reactions, the
initial catalytic steps likely proceed in a similar fashion in these disparate enzymes.
Kinetic, spectroscopic, and structural data, in addition to mutagenesis data and the studies
on the behavior of model compounds, have lent support to the Hanauske-Abel and
Gt‘rnzler model (33) that was proposed for prolyl hydroxylase. The expanded and updated
catalytic model for this superfamily of enzymes predicts the generation of the ferryl-oxo
species (Fe(IV)=O) via the oxidative decarboxylation of Ot-KG. A summary of the steps
proposed in the oxyferryl mechanism for oc—KG—dependent dioxygenases are summarized
in Figure 2. The iron in the resting enzyme is bound by two histidines and one
carboxylate with three waters occupying the open coordination sites. Spectroscopic
studies indicate that oc-KG displaces at least one water molecule upon binding to the
metal through the C-1 carboxylate and C-2 keto groups. Chelation of the iron by or-KG
alters the coordination geometry and redox environment sufficiently to induce binding of
oxygen in an axial position. In some enzymes, the presence of substrates may be
necessary for oxygen to bind to the metal. During the course of oxidative

decarboxylation, one oxygen atom is incorporated into succinate, the other forming an

25

Resting metallocenter /0 °

 

 

o2 C|>
/ Fe(H>\ (ll—KG Fe(III)-"‘O
His Asp ‘. -
1 "0 COO
9H His2
R-C-R O
hydroxylated x
substrate
0 0\
Fe(lll)—OH R-C-R Fe(fi 0
\\ .O -
\ ,0 coo
H
‘ o
Fe(lV)=O R-C-R
sztrate O
CO
SUCClnate /O\O/“\/\COO' 2
Fe(ll)

Figure 2. Oxyferryl mechanistic model for oc-KG-dependent dioxygenases.

26

Fe(IV)-oxo species. In hydroxylation reactions (shown here), the reactive oxygen species
abstracts a hydrogen atom from the substrate to yield a hydroxide radical that recombines
with the substrate to give the final product. During cyclization, ring expansion, or

desaturation reactions, the iron-bound oxo species is reduced by two electrons to water.

Degradation of 2,4—D and Ecology of deA

deA was initially characterized for its ability to degrade 2,4-D (1 10), one of the
most commonly used herbicides in agriculture (121). Despite its widespread use, 2,4-D
does not accumulate in the environment due to its rapid consumption by soil bacteria.
The estimated half-life for 2,4-D in oxic soils is approximately seven days (132). Over
the past few decades, researchers have been particularly interested the mechanism by
which soil microorganisms come to exploit 2,4-D, a xenobiotic compound, as a source
for carbon and energy.

The best characterized pathway for 2,4—D degradation is encoded by genes found
on plasmid pJP4 isolated from Ralstonia eutropha (formerly Alcaligenes eutrophus)
JlVfP134 (Figure 3) (17, 19). In this bacterium, 2,4—D is converted to 2,4-dichlorophenol
which is then broken down by series of enzymes encoded by zdeCDEF to form [3-
ketoadipate, a Kreb’s cycle intermediate. A thorough review of the enzymes involved in
2,4-D degradation as well as an overview of other 2,4-D degradation pathways can be
found in a recent chapter by Hausinger et al. (35) and will be only briefly summarized
here. After deA catalyzes the initial transformation of 2,4—D, 2,4-dichlorophenol
hydroxylase (deB) hydroxylates the aromatic ring to give dichlorocatechol. deB is a

homotetramer with 63 kDa subunits and is similar in sequence and mechanism to

27

2,4-D 2,4-dichlorophenol 3,5-dichlorocatechol 2,4-dichloro-c1’s, crs-muconate

oAcoo -
OOC Cl
OOC
Cl
deA deB deC
1
Cl C
deD
000
'OOC
rde/ deE
chromosomal
O enzymes
B-ketoadipate 2-chlorom aleyl acetate cis-Z-chlorodienelactone

R A, .5 DC” E11 17!! BH. K T C D; EF. 3..

          
 

Figure 3. Pathway for the degradation of 2,4—D by Ralstonia eutropha
JMP134(pJP4)

The enzymes involved in 2,4-D degradation include deA, 2,4-dichlorophenoxyacetic
acid/oc-KG dioxygenase; deB, 2,4—dichlorophenol hydroxylase; deC, 3,5-
dichlorocatechol dioxygenase; deD, chloromuconate cycloisomerase; deE,
dienelactone hydrolase; and deF, maleylacetate reductase. The region of the pJP4
plasmid encoding these enzymes is also shown. The tfd gene names in italics correspond
to the enzymes defined above. tde encodes 2,4—D perrnease and tde/S encodes a LysR-
type regulator of the tfd genes. The above figure is adapted from van der Meer et al.
(126).

28

 

 

enzymes responsible for the hydroxylation of salicylate and other aromatics. Each subunit
contains a molecule of FAD and activity is dependent on NAD(P)H as a cosubstrate.
deC is an Fe(H)-dependent intradiol dioxygenase that converts 3,5-dich10rocatechol to
2,4-dichloro-cis,cis-muconate. The fourth enzyme involved in 2,4-D degradation,
choromuconate isomerase, or deD, converts 2,4—dichloro-cis,cis-muconate to 2-
chlorodienelactone which is then hydrolyzed to 2-chloromalylacetate by dienelactone
hydrolase or deE. The resultant chloromaleylacetate is then degraded by either a
chromosomally—encoded maleylacetate reductase, which has been purified from R.
eutropha JMPl34, or by deF, an enzyme which, by sequence, appears to have the same
activity. The genes encoding deC, deD, deE, and deF are cotranscribed and are
highly similar to operons involved in the degradation of other chlorinated aromatic
compounds such as chlorobenzoate and trichlorobenzene (126).

As shown in Figure 3, there are several other tfd genes associated with the 2,4—D
degradative pathway in addition to the catabolic enzymes. The transcription of tfdA and
the tdeDEFB operon are regulated by a LysR-type regulator, deR/ S, which activates
transcription in response to the inducer 2,4-dichloro-cis,cis-muconate (24).There are two
identical copies of the gene encoding deR/S. Recently, an energy-dependent protein that
facilitates the uptake of 2,4—D, deK, was described and found to be similar to
transporters for 4-hydroxybenzoate and other aromatic compounds (61). In addition, a
second set of genes similar to tdeDEF B and a truncated regulatory gene, tde, are
located in between the two sets of tfd genes. Because the second set of tfd genes

(zdeHC ”E ”F ”311) is not required for 2,4-D degradation, but is induced in the presence of

29

2,4-D, they may have resulted from a gene duplication or gene transfer event that failed
to produce a functional 2,4-D degradation pathway (126).

Researchers have used the pJP4-encoded 2,4—D degradation pathway as a model
to study the assembly of genes necessary to enable an organism to degrade a xenobiotic
compound. Several features of the zfd operon, including the duplication of the regulatory
element, a second copy of the tdeDEF operon, and the presence of the ISJP4 insertion
element sequences in tde and near tde, suggest recent evolution of the 2,4—D
degradation cassette. The data presented below provide additional evidence that the
necessary genes are widespread in environmental bacteria and that they have been
assembled multiple times to enable single organisms to degrade 2,4—D.

Enrichments from soils ((86) and references therein), aquifers and water samples
(3) readily degrade 2,4-D, although often only after an acclimation period (29, 103, 107).
Soulas (107) modeled the dynamics the 2,4—D degradation as the result of competition
between two populations with different 2,4—D degradation rates. In this simulation, the
acclimation period reflects the time required for competitive 2,4wD degraders to achieve
dominance. In support of this theory, phylogenetically diverse 2,4-D degrading isolates
were obtained by directly plating soil bacteria onto 2,4-D-containing medium, but the
diversity of 2,4-D degraders in liquid. enrichment cultures, a more competitive
enviroMent, was greatly reduced (20). Experiments using pristine soils argue that
genetic exchange may also play a role in 2,4-D degradation. Soil enrichments from six
pristine locations metabolized 3-chloroben20ate (3-CBA) and 2,4-D (‘29). While 3-CBA
was catabolized without a lag period and 3--CBA-degrading cultures were readily isolated

from/these enrichments, 2,4-D was generally not consumed until the second ‘week of

.30

incubation and only a small number 2,4-D degrading bacteria were obtained (29). These
data suggest that 2,4-D is degraded by a consortium in environments without a history of
2,4-D treatment.

Diverse bacteria capable of 2,4-D degradation have been isolated from various
environmental samples and enrichments (76). Survey studies of isolates from soil, water
and sewage sludge have found distinct taxa capable of growth on 2,4-D coexisting in
certain environments (3, 20, 117). Over the past thirty years, many 2,4-D degrading
bacteria from different genera have been characterized including strains of Pseudomonas
(3, 31), Bordetella (31) and Alcaligenes (3, 18, 32), Variovorax (123), Burkholderia
(112), Halomonas (66), Flavobacterium (13) Sphingomonas (50), Acinetobacter (6),
Arthrobacter (116), Corynebacterium (3), Nocardioides (5 7), Azotobacter (4),
Achromobacter (108), and Streptomyces (128). Further, genetic and biochemical studies
of many of these strains demonstrate the pervasiveness of deA in these 2,4-D
degradation pathways. Sphingomonas strains that degrade 2,4-D are commonly found in
Michigan soils, but these isolates do not have a tfdA gene that can be detected by gene
probes or a PCR-based screening method for different alleles of the tfdA gene (50).
Nevertheless, biochemcial characterization of the Sphingomonas isolates indicate that
2,4-D degradation is initiated by an oc-KG-dependent dioxygenase (104).

The tfd genes often reside on mobile genetic elements. Plasmids belonging to at
least two different incompatibility groups have been isolated from Alcaligenes species
(18) and soils (118). Plasmid capture experiments were used to isolate pEMT8, a 75-kb
plasmid that encodes tfdA. This plasmid does not appear to contain tde or tdeDEF and

fails to confer activities of the downstream enzymes to a 2,4-D' Ralstom’a eutropha

31

recipient (119). Nearly identical alleles of tfdA have been detected on plasmids isolated
from soils (68) and on the chromosome of plasmid-cured Burkholderia sp. suggesting
that the gene may travel on transposons as well as plasmids (112). Evidence for a
conjugative plasmid capable of integration into the chromosome of different strains has
also been published (51). The pJP4 plasmid can be transferred to bacteria of different
genera such as Variovorax, Pseudomonas, and Burkholderia in soils (16, 25) and similar
plasmids have been isolated from F lavobacterium sp. (13, 50). Different plasmids (7, 64)
and putative degradative transposons, such as Tn5 53 0 carried on pIJBl (134) have also
been purified from 2,4-D degrading strains. Additional proof of the extensive intergeneric
transfer of tfdA is the distribution of at least three different alleles of tfdA across the [3-
Proteobacteria (7 1).

In addition to studies on the dispersal of 2,4—D degradative ability, much attention
has been paid to the creation or assembly of this pathway. Several experiments have been
designed to determine if there is one common ancestor to tfd-based gene cluster or if
these genes were assembled multiple times from other biodegradative pathways. Xia et
al. (133) reported that DNA from soils with no or low exposure to 2,4-D do not hybridize
to the pJP4 tfdA gene probe and no 2,4-D degrading isolates could be cultured from these
soils. Upon exposure to herbicide, 2,4-D degraders with different tfdA and tde genes
become dominant in the different plots. Because there was not a major shift in the total
community structure in response to 2,4-D application, based on random amplified
polymorphic DNA (RAPD) fingerprint analysis, it seems that either a 2,4-D degrading
organism was created from the resident community or a mobile element carrying the

necessary genes was acquired and transferred among resident organisms. Phylogenetic

32

analyses of different 2,4-D degrading bacteria also indicate independent recruitment of
the {fd genes into different organisms during the assembly of the 2,4-D catabolic pathway
(124). Dendrograms plotting the relationships of each of three tfd alleles (g’dA, tde, and
tde) from a collection of 2,4-D degraders are significantly different indicating that these
three genes have not evolved together (28, 124).

The seemingly ubiquitous nature of 2,4-D degrading organisms from both
agricultural and pristine soils as well as from aquatic environments argues that this
metabolic characteristic is important for competitiveness in the environment. Although
some bacteria can degrade 2,4—D by other means (4, 53, 57), deA appears to play a
major role in the breakdown of 2,4-D in the environment. Of all the enzymes in the pJP4-
encoded pathway, deA is the most unique to 2,4-D degradation. Phenols and catechols
are intermediates in numerous catabolic pathways, thus different enzymes exist to

catalyze their degradation. No alternative role for deA has been identified.

Overview of thesis

In this thesis, I present my research on the characterization of the active site of
deA and a study of related enzymes. Chapter 2 focuses specifically on studies relating to
the structure and activity of deA from Ralstonia eutropha JMP134. I describe the site-
directed mutagenesis of ten residues and the characterization of the resultant enzymes to
determine the role, if any, of these amino acids in deA structure or activity. These
studies are an important contribution to research on the structure and mechanism of deA

and the related enzyme TauD. In Chapter 3, I describe the cloning and characterization of

an a-KG-dependent dioxygenase from Saccharomyces cerevisiae to determine if it

33

 

 

possesses activity similar to deA. Work with a strain of S. cerevisiae in which this gene
was disrupted confirms our findings that this enzyme is a sulfonate/oc-KG dioxygenase.
Chapter 4 describes my research on putative deA-like oc-KG-dependent dioxygenases in
soil bacteria. In the last chapter, I summarize my current thoughts on the role of oc-KG-

dependent dioxygenases in the environment and propose future directions for research on

deA and deA-like enzymes.

34

 

10.

11.

Literature cited

Altschul, S. F., W. Gish, W. Miller, E. W. Myers, and D. J. Lipman. 1990.
Basic local alignment search tool. J Mol Biol 215:403-10.

Altschul, S. F., T. L. Madden, A. A. Schiiffer, J. Zhang, Z. Zhang, W. Miller,
and D. J. Lipman. 1997. Gapped BLAST and PSI-BLAST: a new generation of
protein database search programs. Nucleic Acids Res. 25:3389-3402.

Amy, P. S., J. W. Schulke, L. M. Frazier, and R. J. Seidler. 1985.
Characterization of aquatic bacteria and cloning of genes specifying partial
degradation of 2,4-dichlorophenoxyacetic acid. Appl. Environ. Microbiol.
49:1237-1245.

Balajee, S., and A. Mahadevan. 1990. Dissimilation of 2,4-
dichlorophenoxyacetic acid by Azotobacter chroococcum. Xenobiotica 20:607—
617.

Baldwin, J. E., K. D. Merritt, C. J. Schofield, S. W. Elson, and K. H.
Baggaley. 1993. Studies on the stereospecificity of the clavaminic acid synthase
catalyzed hydroxylation reaction. J. Chem. Soc., Chem. Comm.:l30l-1302.

Bell, G. R. 1957. Some morphological and biochemical characteristics of a soil
bacterium which decomposes 2,4-dichlorophenoxyacetic acid. Can. J. Microbiol.
3: 82 1-840.

Bhat, M. A., M. Tsuda, K. Horiike, M. N ozaki, C. S. Vaidyanathan, and T.
N akazawa. 1994. Identification and characterization of a new plasmid carrying
genes for degradation of 2,4-dichlorophenoxyacetate from Pseudomonas cepacia
CSV90. Appl. Environ. Microbiol. 60:307-312.

Borovok, I., O. Landman, R. Kreisberg-Zakarin, Y. Aharonowitz, and G.
Cohen. 1996. Ferrous active site of isopenicillin N synthase: genetic and
sequence analysis of the endogenous ligands. Biochemistry 35: 1981-1987.

Britsch, L. 1990. Purification and characterization of flavone synthase I, a 2-
oxoglutarate-dependent desaturase. Arch. Biochem. Biophys. 282: 1 52-160.

Britsch, L., and H. Grisebach. 1986. Purification and characterization of (2S)-
flavanone 3-hydroxylase from Petunia hybrida. Eur. J. Biochem. 156:569-577.

Busby, R. W., M. D.-T. Chang, R. C. Busby, J. Wimp, and C. A. Townsend.

1995. Expression and purification of two isozymes of clavaminate synthase and
initial characterization of the iron binding site. J. Biol. Chem. 270:4262—4269.

35

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

Chahal, A., M. Khan, S. G. Pai, E. Barbosa, and 1. Singh. 1998. Restoration of
phytanic acid oxidation in Refsum disease fibroblasts from patients with
mutations in the phytanoyl-CoA hydroxylase gene. FEBS Lett. 429: 1 19-122.

Chaudry, G. R., and G. H. Huang. 1988. Isolation and characterization of a new
plasmid from a F lavobacterium sp. which carries the genes for the degradation of
2,4-dichlorophenoxyacetate. J. Bacteriol. 170:3897-3902.

De Carolis, E., and V. De Luca. 1993. Purification, characterization, and kinetic
analysis of a 2-oxoglutarate-dependent dioxygenase involved in vindoline
biosynthesis of Catharanthus roseus. J. Biol. Chem. 268:5504-5511.

De Long, L., S. P. J. Albracht, and A. Kemp. 1982. Prolyl 4-hydroxylase
activity in relation to the oxidation state of enzyme-bound iron. Biochim.
Biophys. Acta. 704:326-332.

DiGiovanni, G. D., J. W. N eilson, I. L. Pepper, and N. A. Sinclair. 1996. Gene
transfer of Alcaligenes eutrophus JMP134 plasmid pJP4 to indigenous soil
recipients. Appl. Environ. Microbiol. 62 :252 1 —2526.

Don, R. H., and J. M. Pemberton. 1985. Genetic and physical map of the 2,4-
dichlorophenoxyacetic acid-degrading plasmid pJP4. J. Bacteriol. 161:466-468.

Don, R. H., and J. M. Pemberton. 1981. Properties of six pesticide degradation
plasmids isolated from Alcaligenes paradoxus and Alcaligenes eutrophus. J.
Bacteriol. 145:681-686.

Don, R. H., A. J. Weightman, H. J. Knackmuss, and K. N. Timmis. 1985.
Transposon mutagenesis and cloning analysis of the pathways for degradation of
2,4-dichlorophenoxyacetic acid and 3-chorobenzoate in Alcaligenes eutrophus
JMP134 (pJP4). J. Bacteriol. 161:85-90.

Dunbar, J., S. White, and L. Forney. 1997. Genetic diversity through the
looking glass: effect of enrichment bias. Appl. Environ. Microbiol. 63: 1326-133 1.

Eichhorn, E., J. R. van der Ploeg, M. A. Kertesz, and T. Leisinger. 1997.

Characterization of oc—ketoglutarate-dependent taurine dioxygenase from
Escherichia coli. J. Biol. Chem. 272:23031-23036.

Englard, S., J. S. Blanchard, and C. F. Midelfort. 1985. y-Butyrobetaine
hydroxylase: stereochemical course of the hydroxylation reaction. Biochemistry
24:1110-1116.

Eriksson, M., J. Myllyharju, H. Tu, M. Hellman, and K. I. Kivirikko. 1999.

Evidence for 4-hydroxyproline in viral proteins. Characterization of a viral prolyl
4-hydroxylase and its peptide substrates. J Biol Chem 274:2213 1-22134.

36

 

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

34.

Filer, K., and A. R. Harker. 1997. Identification of the inducing agent of the
2,4-dichlorophenoxyacetic acid pathway encoded by plasmid pJ P4. Appl.
Environ. Microbiol. 63:317-320.

Friedrich, B., M. Meyer, and H. G. Schlegel. 1983. Transfer and expression of
the herbicide-degrading plasmid pJP4 in aerobic autotrophic bacteria. Arch.
Microbiol. 134:92-7.

Fukumori, F ., and R. P. Hausinger. 1993. Alcaligenes eutrophus JMP134 "2,4-

dichlorophenoxyacetate monooxygenase" is an oc—ketoglutarate-dependent
dioxygenase. J. Bacteriol. 175:2083-2086.

Fukumori, F., and R. P. Hausinger. 1993. Purification and characterization of

2,4-dichlorophenoxyacetate/OL-ketoglutarate dioxygenase. J. Biol. Chem.
268:24311-24317.

Fulthorpe, R. R., C. McGowan, O. V. Maltseva, W. E. Holben, and J. M.
Tiedje. 1995. 2,4-Dichlorophenoxyacetic acid-degrading bacteria contain mosaics
of catabolic genes. Appl. Environ. Microbiol. 61:3274-3281.

Fulthorpe, R. R., A. N. Rhodes, and J. M. T iedje. 1996. Pristine soils
mineralize 3-chlorobenzoate and 2,4-dichlorophenoxyacetate via different
microbial p0pulations. Appl. Environ. Microbiol. 62:1159-1 166.

Galland, S., F. Le Borgne, D. Guyonnet, P. Clouet, and J. Demarquoy. 1998.
Purification and characterization of the rat liver gamma-butyrobetaine
hydroxylase. Mol. Cell. Biochem. 178:163-8.

Greer, L. E., J. A. Robinson, and D. R. Shelton. 1992. Kinetic comparison of
seven strains of 2,4-dichlorophenoxyacetic acid-degrading bacteria. Appl.
Environ. Microbiol. 58: 1027-1030.

Gunalan, and J. C. Fournier. 1993. Effect of microbial competition on the
survival and activity of 2,4-D degrading Alcaligenes xylosoxidans subsp.
denitrificans added to soil. Lett. Appl. Microbiol. 16: 178-181.

Hanauske—Abel, H. M., and V. Giinzler. 1982. A stereochemical concept for the
catalytic mechanism of prolyl hydroxylase. Applicability to classification and
design of inhibitors. J. Theor. Biol. 94:421-455.

Hashimoto, T., and Y. Yamada. 1987. Purification and characterization of

hyoscyamine 6B—hydroxylase, from root cultures of Hyoscyamus niger L. Eur. J.
Biochem. 164:277-285.

37

 
 

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

46.

Hausinger, R. P., F. Fukumori, D. A. Hogan, T. M. Sassanella, Y. Kamagata,
H. Takami, and R. E. Wallace. 1996. Biochemistry of 2,4-D degradation:

evolutionary implications, p. 35-51. In K. Horikoshi, M. Fukuda, and T. Kudo
(ed.), Microbial Diversity and Genetics of Biodegradation. Japan Scientific Press,

Tokyo.

Hedden, P. 1992. 2-Oxog1utarate-dependent dioxygenases in plants: mechanism
and function. Biochem. Soc. Trans. 20:373-377.

Hedden, P. 1997. The oxidases of giberellin biosynthesis: Their function and
mechanism. Physio]. Planta. 101 :709-71 9.

Hegg, E. L., and L. Que Jr. 1997. The 2-His-1-carboxylate facial triad. Eur. J.
Biochem. 250:625-629.

Hegg, E. L., A. K. Whiting, J. McCracken, R. P. Hausinger, and J. Que, L.
1999. The herbicide-degrading a-keto acid enzyme deA: metal coordination
environment and mechanistic insights. Biochemistry In press.

Hogan, D. A., T. A. Auchtung, and R. P. Hausinger. 1999. Cloning and
characterization of a sulfonate/a—ketoglutarate dioxygenase from Saccharomyces
cerevisiae. J. Bacteriol. 181:5876—5879.

Holme, E. 1975. A kintic study of thymine 7-hydroxylase from Neurosporra
crassa. Biochemistry 14:4999-5003.

Holme, E., S. Lindstedt, and I. Nordin. 1982. Uncoupling in the y—
butyrobetaine hydroxylase reaction by D- and L-carnitine. Biochem. Biophys.
Res. Comm. 107:518-524.

Holton, T. A., F. Brugliera, and Y. Tanaka. 1993. Cloning and expression of
flavonol synthase from Petunia hybrida. Plant J 4:1003-10.

Hulse, J. D., S. R. Ellis, and L. M. Henderson. 1978. Camitine biosynthesis. [3-

Hydroxylation of trimethyllysine by an a-ketoglutarate-dependent mitochondrial
dioxygenase. J. Biol. Chem. 253:1654-9.

Hutton Jr., J. J., A. L. Tappel, and S. Udenfriend. 1967. Cofactor and substrate
requirements of collagen proline hydroxylase. Arch. Biochem. Biophys. 118:231-
240.

Jansen, G. A., S. J. Mihalik, P. A. Watkins, C. Jakobs, H. W. Moser, and R.
J. Wanders. 1998. Characterization of phytanoyl-coenzyme A hydroxylase in
human liver and activity measurements in patients with peroxisomal disorders.
Clin. Chim. Acta 271:203-11.

38

47.

48.

49.

50.

51.

52.

53.

54.

55.

56.

Jia, S., K. McGinnis, W. J. VanDusen, C. J. Burke, A. Kuo, P. R. Griffin, M.
K. Sardana, K. O. Elliston, A. M. Stern, and P. A. Friedman. 1994. A fully
active catalytic domain of bovine aspartyl (asparaginyl)-B-hydroxylase expressed
in Escherichia coli: characterization and evidence for the identification of an

active-site region in vertebrate-OL-ketoglutarate-dependent dioxygenases. Proc.
Natl. Acad. Sci. 91 :7227-723 1.

Jiang, F., J. Peisach, L.-J. Ming, L. Que, Jr., and V. J. Chen. 1991. Electron
spin-echo envelope modulation studies of the Cu(H)-substituted derivative of
isopenicillin N synthase: a structural and spectroscopic model. Biochemistry
30:11437-11445.

John, P. 1997. Ethylene biosynthesis: The role of l-aminocyclopropane-l-
carboxylate (ACC) oxidase, and its possible evolutionary origin. Physiologia
Plantarum 100:583-592.

Ka, J. 0., W. E. Holben, and J. M. Tiedje. 1994. Genetic and phenotypic
diversity of 2,4-dichlorophenoxyacetic acid (2,4-D)-degrading bacteria isolated
from 2,4-D-treated field soils. Appl. Environ. Microbiol. 60:1106-11 15.

Ka, J. 0., and J. M. Tiedje. 1994. Integration and excision of a 2,4-
dichlorophenoxyacetic acid degradative plasmid in Alcaligenes paradoxus and
evidence of its natural intergeneric transfer. J. Bacteriol. 176:5284-5289.

Kellokumpu, S., R. Sormunen, J. Heikkinene, and R. Myllyla. 1994. Lysyl
hydroxylase, a collage processing enzyme, exemplifies a novel class of luminally-
oriented peripheral membrane protein in the endoplasmic reticulum. J. Biol.
Chem. 269:30524-30529.

Kilbane, J. J., D. K. Chatterjee, J. S. Karns, S. T. Kellogg, and A. M.
Chakrabarty. 1982. Biodegradation of 2,4,5-trichlorophenoxyacetic acid by a
pure culture of Pseudomonas cepacia. Appl. Environ. Microbiol. 44:72-78.

Kivirikko, K. I., and T. Pihlajaniemi. 1998. Collagen hydroxylases and the
protein disulfide isomerase subunit of prolyl 4-hydroxylases. Adv. Enzymol.
Relat. Areas Mol. Biol. 72:328-398.

Korioth, F., C. Gieffers, and J. Frey. 1994. Cloning and characterization of the
human gene encoding aspartyl B-hydroxylase. Gene 150:395-399.

Kovacevic, S., B. J. Weigel, M. B. Tobin, T. D. Ingolia, and J. R. Miller. 1989.
Cloning, characterization, and expression in Escherichia coli of the Streptomyces
clavuligerus gene encoding deacetoxycephalosporin C synthetase. J. Bacteriol.
171 :754-60.

39

57.

58.

59.

60.

61.

62.

63.

64.

65.

66.

67.

Kozyreva, L. P., Y. V. Shurukhin, Z. I. Finkelstein, B. P. Baskunov, and L. A.
Golovleva. 1993. Metabolism of the herbicide 2,4—D by a Nocardioides simplex
strain. Mikrobiol. 62:110-119.

Kreisberg-Zakarin, R., I. Borovok, M. Yanko, Y. Aharonowitz, and G.
Cohen. 1999. Recent advances in the structure and function of isopenicillin N
synthase. Antonie Van Leeuwenhoek 7 5:33—39.

Lamberg, A., T. Pihlajaniemi, and K. I. Kivirikko. 1995. Site-directed

mutagenesis of the or subunit of the human prolyl 4-hydroxylase. J. Biol. Chem.
270:9926-9931.

Lawlor, E. J., S. W. Elson, S. Holland, R. Cassels, and J. E. Hodgson. 1994.
Expression in Escherichia coli of a clavaminic acid synthase isozyme: 3
trifuctional oxygenases involved in clavulanic acid biosynthesis. Tetrahedron

50:8737-8748.

Leveau, J. H., A. J. B. Zehnder, and J. R. Van der Meer. 1998. The tde gene
product facilitates the uptake of 2,4-dichlorophenoxyacetate by Ralstonia
eutropha JMP134(pJP4). J. Bacteriol. 180:2237-2243.

Lindstedt, G., and S. Lindstedt. 1970. Cofactor requirements of y-butyrobetaine
hydroxylase from rat liver. J. Biol. Chem. 245:4178-4180.

Lukacin, R., and L. Britsch. 1997. Identification of strictly conserved histidine

and arginine residues as part of the active site in Petunia hybrida flavanone 3B-
hydroxylase. Eur. J. Biochem. 249:748-757.

Mae, A. A., R. O. Martis, N. R. Ausmees, V. M. K6iv, and A. L. Heinaru.
1993. Characterization of a new 2,4-dichlorophenoxyacetic acid degrading
plasmid pEST4011: physical map and localization of catabolic genes. J. Gen.
Microbiol. 139:3165-3170.

Majamaa, K., V. Giinzler, H. M. Hanauske—Abel, R. Myllylii, and K. I.
Kivirikko. 1986. Partial identity of the 2-oxoglutarate and ascorbate binding sites
of prolyl 4-hydroxylase. J. Biol. Chem. 261:7819-7823.

Maltseva, O. V., C. McGowan, R. Fulthorpe, and P. J. Oriel. 1996.
Degradation of 2,4-dichlorophenoxyacetic acid by haloalkaliphilic bacteria.
Microbiol. 142:1115-1122.

Mao, Y. Q., M. Vargolu, and D. H. Sherman. 1999. Molecular characterization

and analysis of the biosynthetic gene cluster for the antiturnor antibiotic
mitomycin C from Streptomyces lavendulae NRRL2564. Chem. Biol. 6:251-263.

40

68.

69.

70.

71.

72.

73.

74.

75.

76.

77.

78.

Matheson, V. G., L. F. F orney, Y. Suwa, C. H. Nakatsu, A. J. Sexstone, and
W. E. Holben. 1996. Evidence for acquisition in nature of a chromosomal 2,4-

dichlorophenoxyacetic acid/oc-ketoglutarate dioxygenase gene by different
Burkholderia spp. Appl. Environ. Microbiol. 62:2457-2463.

Matsuda, J., S. Okabe, T. Hashimoto, and Y. Yamada. 1991. Molecular
cloning of hyoscyamine 6 beta-hydroxylase, a 2-oxoglutarate- dependent
dioxygenase, from cultured roots of Hyoscyamus niger. J. Biol. Chem. 266:9460-
9464.

McGinnis, K., G. M. Ku, W. J. VanDusen, J. Fu, V. Garsky, A. M. Stern, and
P. A. Friedman. 1996. Site-directed mutagenesis of residues in a conserved
region of bovine apartyl (asparaginyl) B-hydroxylase: evidence that histidine 67 5

has a role in binding Fe+2. Biochemistry 35:3957-3962.

McGowan, C., R. Fulthorpe, A. Wright, and J. M. Tiedje. 1998. Evidence for
interspecies gene transfer in the evolution of 2,4-dichlorophenoxyacetic acid
degraders. Appl. Environ. Microbiol. 64:4089-4092.

McGowan, S. J., M. T. Holden, B. W. Bycroft, and G. P. Salmond. 1999.
Molecular genetics of carbapenem antibiotic biosynthesis. Antonie Van
Leeuwenhoek 75:135-141.

Metcalf, W. W., and R. S. Wolfe. 1998. Molecular genetic analysis of phosphite
and hypophosphite oxidation by Pseudomonas stutzeri WM88. J. Bacteriol.
180:5547-5558.

Miller, R. L., and H. H. Varner. 1979. Purification and enzymic properties of
lysyl hydroxylase from fetal porcine skin. Biochemistry 18:5928-5932.

Ming, L.-J., L. Que, Jr., A. Kriauciunas, C. A. Frolik, and V. J. Chen. 1990.
Coordination chemistry of the metal binding site of isopenicillin N synthase.
Inorg. Chem. 29:1111-1112.

Miwa, N ., and S. Kuwatsuka. 1991. Characteristics of 2,4-
dichlorophenoxyacetic acid (2,4-D)-degrading microorganisms isolated from
different soils. Soil Sci. Plant Nutr. 37 :575-58 1.

Mori, H., T. Shiasaki, K. Yano, and A. Ozaki. 1997. Purification and cloning of
a proline 3-hydroxylase, a novel enzyme which hydroxylates fiee L-proline to cis—
3-hydroxyl-L-proline. J. Bacteriol. 179:5677-5683.

Mori, H., T. Shibasaki, Y. Uozaki, K. Ochiai, and A. Ozaki. 1996. Detection of
novel proline 3-hydroxylase activities in Streptomyces and Bacillus spp. by regio-
and stereospecific hydroxylation of L-proline. Appl. Environ. Microbiol.
62:1903-1907.

41

79.

80.

81.

82.

83.

84.

85.

86.

87.

88.

Myllyharju, J., and K. I. Kivirikko. 1997. Characterization of the iron- and 2-
oxoglutarate-binding sites of human prolyl 4-hydroxylase. EIVIBO J. 16:1173-
1 180.

Myllylii, R., K. Majamaa, V. Giinzler, H. N. Hanauske-Abel, and K. I.
Kivirikko. 1984. Ascorbate is consumed stoichiometrically in the uncoupled
reactions catalyzed by prolyl 4-hydroxylase and lysyl hydroxylase. J. Biol. Chem.
259:5403-5405.

Myllyla, R., T. Pihlajaniemi, L. Pajunen, T. Turpeenniemi-Hujanen, and K.
I. Kivirikko. 1991. Molecular cloning of chick lysyl hydroxylase. Little
homology in primary structure to the two types of subunit of prolyl 4-
hydroxylase. J. Biol. Chem. 266:2805-2810.

Myllylii, R., L. Tuderman, and K. I. Kivirikko. 1977. Mechanism of the prolyl
hydroxylase reaction: 2. Kinetic analysis of the reaction sequence. Eur. J.
Biochem. 80:349-357.

 

Nagahama, K., T. Ogawa, T. Fujii, M. Tazaki, S. Tanase, Y. Merino, and H.
Fukuda. 1991. Purification and properties of an ethylene-forming enzyme from
Pseudomonas syringae pv. phaseolicola PK2. J. Gen. Microbiol. 137:2281-2286.

Nickel, K., M. J.-F. Suter, and H.-P. E. Kohler. 1997. Involvement of two oc-
ketoglutarate-dependent dioxygenases in enantioselective degradation of (R)- and
(8)-mecoprop by Sphingomonas herbicidovorans MI-I. J. Bacteriol. 179:6674-
6679.

Orville, A. M., V. J. Chen, A. Kriauciunas, M. R. Harpel, B. G. Fox, E.
Miinck, and J. D. Lipscomb. 1992. Thiolate ligation of the active site F e+2 of
isopenicillin N synthase derived from substrate rather than endogenous cysteine:

spectroscopic studies of site-specific Cys to Ser mutated enzymes. Biochemistry
31:4602-4612.

Ou, L.-T. 1984. 2,4-D degradation and 2,4-D degrading microorganisms in soils.
Soil Sci. 137:100-107.

Pavel, E. G., J. Zhou, R. W. Busby, M. Gunsior, C. A. Townsend, and E. 1.
Solomon. 1998. Circular dichroism and magnetic circular dichroism
spectroscopic studies of the non-heme ferrous active site in clavaminate synthase
and its interaction with ct-ketoglutarate cosubstrate. J. Am. Chem. Soc. 120:743-

754.
Pirskanen, A., A.-M. Kaimio, R. Myllylii, and K. I. Kivirikko. 1996. Site-

directed mutagenesis of human lysyl hydroxylase expressed in insect cells. J.
Biol. Chem. 271:9398-9402.

42

89.

90.

91.

92.

93.

94.

95.

96.

97.

98.

99.

Pirskanen, A., A.-M. Kaimio, R. Myllylii, and K. I. Kivirikko. 1996. Site—
directed mutagenesis of human lysyl hydroxylase expressed in insect cells.

Identification of histidine residues and an aspartic acid residue critical for
catalytic activity. J. Biol. Chem. 271:9398-9402.

Prescott, A. G. 1993. A dilemma of dioxygenases (or where biochemsitry and
molecular biology fail to meet). J. Exp. Bot. 44:849-861.

Prescott, A. G., and P. John. 1996. Dioxygenases: Molecular structure and role
in plant metabolism. Annu. Rev. Plant Physiol. Plant Mol. Biol. 47 :245-271.

Puistola, U., T. M. Turpeenniemi-Hujanen, R. Myllylii, and K. I. Kivirikko.
1980. Studies on the lysyl hydoxylase reaction: H. Inhibition kinetics and the
reaction mechanism. Biochim. Biophys. Acta 611 :5 1-60.

Ramon, D., L. Carramolino, C. Patino, F. Sanchez, and M. A. Penalva. 1987.
Cloning and characterization of the isopenicillin N synthetase gene mediating the
formation of the beta-lactam ring in Aspergillus nidulans. Gene 57:171-81.

Roach, P. L., I. J. Clifton, V. Fiiliip, K. Harlos, G. J. Barton, J. Hajdu, I.
Andersson, C. J. Schofield, and J. E. Baldwin. 1995. Crystal structure of
isopenicillin N synthase is the first from a new structural family of enzymes.
Nature 375:700-704.

Roach, P. L., I. J. Clifton, C. M. H. Hensgens, N. Shibata, C. J. Schofield, J.
Hajdu, and J. E. Baldwin. 1997. Structure of isopenicillin N synthase complexed
with substrate and the mechanism of penicillin formation. Nature 387 :827—830.

Ryle, M. J., R. Padmakumar, and R. P. Hausinger. 1999. Stopped-flow kinetic
analysis of Escherchia coli taurine/OL-ketoglutarate dioxygenase: interactions with
oc-ketoglutarate, taurine, and oxygen. Biochem. In press.

Saari, R. E., and R. P. Hausinger. 1998. Ascorbic acid-dependent turnover and
reactivation of 2,4- dichlorophenoxyacetic acid/alpha-ketoglutarate dioxygenase
using thiophenoxyacetic acid. Biochemistry 37 :3035-3042.

Saari, R. E., D. A. Hogan, and R. P. Hausinger. 1999. Stereospecific
degradation of the phenoxypropionate herbicide dichlorprop. J. Molec. Catalysis
B: Enzymatic 266:421-428.

Saito, K., M. Kobayashi, Z. Gong, Y. Tanaka, and M. Yamazaki. 1999. Direct
evidence for anthocyanidin synthase as a 2-oxoglutarate-dependent oxygenase:
molecular cloning and functional expression of cDNA from a red forma of Perilla
frutescens. Plant Journal 17: 181-1 89.

43

100.

101.

102.

103.

104.

105.

106.

107.

108.

109.

110.

Saito, T., S. S. Kwak, Y. Kamiya, H. Yamane, A. Sakurai, N. Murofushi, and
N. Takahashi. 1991. Plant Cell. Physiol. 32:239-245.

Sami, M., T. J. N. Brown, P. L. Roach, C. J. Schofield, and J. E. Baldwin.
1997. Glutamine--330 is not essential for activity in isopenicillin N synthase from
Aspergillus nidulans. FEBS Lett. 405: 191-194.

Samson, S. M., J. E. Dotzlaf, M. L. Slisz, G. W. Becker, R. M. Van Frank, L.
E. Veal, W. K. Yeh, J. R. Miller, S. W. Queener, and T. D. Ingolia. 1987.
Cloning and expression of the fungal expandase/hydroxylase gene involved in
cephalosporin biosynthesis. Biotechnology 5:1207-1214.

Sandmann, E. R. I. C., M. A. Loos, and L. P. van Dyk. 1988. Microbial
degradation of 2,4-dichlorophenoxyacetic acid in soil. Rev. Environ. Contamin.
Tox. 101:1-53.

Sassanella, T. M., F. Fukumori, M. Bagdasarian, and R. P. Hausinger. 1997.
Use of 4-nitrophenoxyacetic acid for the detection and quantitation of 2,4-
dichlorophenoxyacetic acid (2,4-D)/oc-ketoglutarate dioxygenase activity in 2,4-D
degrading microorganisms. Appl. Environ. Microbiol. 63:1 189-1 191.

Scott, R. A., S. Wang, M. K. Eidsness, A. Kriauciunas, C. A. Frolik, and V. J.
Chen. 1992. X-ray absorption spectroscopic studies of the high-spin iron(II)
active site of isopenicillin N synthase: evidence for F e-S interaction in the
enzyme-substrate complex. Biochemistry 31 :45 96-601 .

Shibasaki, T., H. Mori, S. Chiba, and A. Ozaki. 1999. Microbial proline 4-
hydroxylase screening and gene cloning. Appl. Environ. Microbiol. 65:4028-
4031.

Soulas, G. 1993. Evidence for the existence of different physiological groups in
the microbial community responsible for 2,4-D mineralization in soil. Soil Biol.
Biochem. 25:443-449.

Steenson, T. I., and N. Walker. 1957. The pathway of breakdown of 2,4-
dichloro- and 4-chloro-2-methyl-phenoxyacetic acid by bacteria. J. Gen.
Microbiol. 16:146-155.

Stintzi, A., Z. Johnson, M. Stonehouse, U. Ochsner, J. Meyer, M. Vasil, and
K. Poole. 1999. The pvc gene cluster of Pseudomonas aeruginosa: role in
synthesis of the pyoverdine chromophore and regulation by Pth and PvdS. J.
Bacteriol. 181:4118-4124.

Streber, W. R., K. N. Timmis, and M. H. Zenk. 1987. Analysis, cloning, and

high-level expression of 2,4-dichlorophenoxyacetate monooxygenase gene tfdA of
Alcaligenes eutrophus JMP134. J. Bacteriol. 169:2950-2955.

44

111.

112.

113.

114.

115.

116.

117.

118.

119.

120.

121.

122.

Stubbe, J. 1985. Identification of two alpha—ketoglutarate-dependent
dioxygenases in extracts of Rhodotoru/a glutinis catalyzing deoxyuridine
hydroxylation. J. Biol. Chem. 260:9972—9975.

Suwa, Y., A. D. Wright, F. Fukumori, K. A. Nummy, R. P. Hausinger, W. E.
Holben, and L. J. Forney. 1996. Characterization of a chromosomally encoded
2,4-dichlorophenoxyacetic acid/a-ketoglutarate dioxygenase from Burkhola’eria
sp. strain RASC. Appl. Environ. Microbiol. 62:2464-2469.

Tan, D. S. H., and T.-S. Sim. 1996. Functional analysis of conserved histidine
residues in Cephalosporin acremoniumisopenicillin N synthase by site-directed
mutagenesis. J. Biol. Chem. 271:889-894.

Tett, V. A., A. J. Willetts, and H. M. Lappin-Scott. 1997. Biodegradation ofthe
chlorophenoxy herbicide (R)—(+)—mecoprop by Alcaligenes denitrificans.
Biodegradation 8:43-52.

Thornburg, L. D., M. T. Lai, J. S. Wishnok, and J. Stubbe. 1993. A non-heme
protein with heme tendencies: an investigation of the substrate specificity of
thymine hydroxylase. Biochemistry 32: 14023-14033.

Tiedje, J. M., and M. Alexander. 1969. Enzymatic cleavage of the ether bond
of 2,4-dichlorophenoxyacetate. J. Agr. Food Chem. 17: 1080-1084.

Tonso, N. L., V. G. Matheson, and W. E. Holben. 1995. Polyphasic
characterization of a suite of bacterial isolates capable of degrading 2,4-D.
Microb. Ecol. 3:3-24.

Top, E. M., W. E. Holben, and L. J. Forney. 1995. Characterization of diverse
2,4-dichlorophenoxyacetic acid-degradative plasmids isolated from soil by
complementation. Appl. Environ. Microbiol. 61: 1691-1698.

Top, E. M., O. V. Maltseva, and L. J. Forney. 1996. Capture ofa catabolic
plasmid that encodes only 2,4-D/oc-ketoglutarate dioxygenase (deA) by genetic
complementation. Appl. Environ. Microbiol. 62:2470-2476.

Unfinished Genome Database Website. 11 October 1999.
http://www.ncbi.nlm.nih.gov/BLAST/ unfmishedgenomehtml.

United States Department of Agriculture. 1996. Biologic and economic
benefits from the use of phenoxy herbicides in the United States NAPIAP Report
No. 1-PA-96.

Valegard, K., A. C. T. van Scheltinga, M. D. Lloyd, T. Hara, S. Ramaswamy,
A. Perrakis, A. Thompson, H.-J. Lee, J. E. Baldwin, C. J. Schofield, J. Hajdu,

45

123.

124.

125.

126.

127.

128.

129.

130.

131.

132.

and I. Andersson. 1998. Structure of a cephalosporin synthase. Nature 394:805-
809.

Vallaeys, T., L. Albino, G. Soulas, A. D. Wright, and A. J. Weightman. 1998.
Isolation and characterization of a stable 2,4-dichlorophonoxyacetic acid
degrading bacterium, Variovorax paradoxus, using chemostat culture. Biotechnol.
Lett. 20:1073-6.

Vallaeys, T., L. Courde, C. McGowan, A. D. Wright, and R. R. Fulthorpe.
1999. Phylogenetic analyses indicate independent recruitment of diverse gene

casettes during assemblage of the 2,4-D catabolic pathway. FEMS Microbiol.
Ecology 28:373-382.

Valtavaara, M., H. Papponene, A.-M. Pirttilii, K. Hiltunen, H. Helander, and
R. Myllylii. 1997. Cloning and characterization of a novel human lysyl
hydroxylase isoform highly expressed in pancreas and muscle. J. Biol. Chem.

272:6831-6834.

van der Meer, J. R., W. M. de Vos, S. Harayama, and A. J. B. Zehnder. 1992.
Molecular mechanisms of genetic adaption to xenobiotic compounds. Microbiol.
Rev. 56:677-694.

Vermeij, P., J. Hummerjohann, T. Leisinger, and M. A. Kertesz. 1999.
Unpublished.

Walker, R. L., and A. S. Newman. 1956. Microbial decomposition of 2,4-
dichlorophenoxyacetic acid. Appl. Microbiol 4:201-207.

Wang, Q., W. J. VanDusen, C. J. Petroski, V. M. Garsky, A. M. Stern, and P.

A. Friedman. 1991. Bovine liver aspartyl B—hydroxylase. Purification and
characterization. J. Biol. Chem. 266: 14004-14010.

Wehbie, R. S., N. S. Punekar, and H. A. Lardy. 1988. Rat liver y-butyrobetaine
hydroxylase catalyzed reaction: influence of potassium, substrates and substrate
analogues on hydroxylation and decarboxylation. Biochemistry 27 :2222-2228.

Whiting, A. K., L. Que J r., R. E. Saari, R. P. Hausinger, M. A. Fredrick, and
J. McCracken. 1997. Metal coordination environment of a Cu(H)-substituted Ot-

keto acid-dependent dioxygenase that degrades the herbicide 2,4—D. J. Amer.
Chem. Soc. 119:3413-3414.

Wilson, R. D., J. Geronimo, and J. A. Armbruster. 1997. Dissipation in field

soils after application of 2,4-D dimethylamine salt and 2,4-D 2-ethylhexyl ester.
Environmental Toxicology and Chemistry 16: 1239-1246.

46

133.

134.

135.

Xia, X., J. Bollinger, and A. Ogram. 1995. Molecular genetic analysis of the
response of three soil microbial communties to the application of 2,4-D. Mol.
Ecol. 4:17-28.

Xia, X.-S., S. Aathithan, K. Oswiecimska, A. R. W. Smith, and I. J. Bruce.
1998. A novel plasmid pIJBl possessing a putative 2,4-dichlorophenoxyacetate

degradative transposon TN5530 in Burkholderia cepacia strain 2a. Plasmid
39: 1 54-1 59.

Zhang, Z., C. J. Schofield, J. E. Baldwin, P. Thomas, and P. John. 1995.

Expression, purification and characterization of l-aminocyclopropane-l-
carboxylate oxidase from tomato in Escherichia coli. Biochem. J. 307 :77-85.

47

CHAPTER 2
CHARACTERIZATION OF THE ACTIVE SITE OF 2,4-
DICHLOROPHENOXYACETIC ACID/a-KETOGLUTARATE DIOXYGENASE

Many of the results in this chapter have been submitted for publication to the
Journal of Biological Chemistry in a manuscript entitled “Site-directed mutagenesis of

2,4-dichlorophenoxyacetic acid/a-ketoglutarate dioxygenase” by Hogan, D.A., S. R.
Smith, E. A. Saari, J. McCracken, and R. P. Hausinger.

48

Introduction

2,4-Dichlorophenoxyacetic acid (2,4-D)/oc—ketoglutarate (cc-KG) dioxygenase
(deA) is an Fe(H)— and oc-KG—dependent enzyme that catalyzes the first step in
degradation of the herbicide 2,4-D. This enzyme couples the oxidative decarboxylation of
a-KG to the hydroxylation of a side—chain carbon atom. The resultant hemiacetal
spontaneously decomposes to form 2,4-dichlorophenol and glyoxalate (6).
Mechanistically, deA resembles numerous other OL-KG—dependent dioxygenases from
plants, animals, fimgi, and bacteria that catalyze similar hydroxylation reactions at
unactivated carbon centers (4, 28). The best—studied oc-KG-dependent hydroxylases,
including prolyl and lysyl hydroxylase (14) and flavanone hydroxylase (8, 28), show
sequence identity to isopenicillin N synthase (IPNS) and thus fall into Group I of the oc-
KG-dependent dioxygenase superfamily (defined in Chapter 1). These enzymes have an
H-X-D-Xss-H-Xlo-R motif in common (2, l8) and site-directed mutagenesis studies
have confirmed the importance of these residues (15, 18, 21, 25, 27). Through various
biochemical and biophysical studies of Group I enzymes, these amino acids are
implicated in creating the F e(H) and oc-KG binding sites (17, 29, 30, 33). The sequence of
deA does not have significant overall identity to the sequences of Group I enzymes.

Although deA is not similar in sequence to the Group I oc-KG-dependent
dioxygenases, it is clearly homologous to a number of other enzymes (Group II).
Sequences of Escherichia coli taurine/(x-KG dioxygenase (TauD) (5) and sulfonate/oc-KG
dioxygenase from Saccharomyces cerevisiae (11) have 25-30% overall identity to deA.

Alignment of deA, TauD and sulfonate/oc-KG dioxygenase indicates the conservation of

49

an H-X—D motif (His 113 and Asp 115 in deA) and three other histidines (Appendix A).
One invariant histidine, His 167, is positioned to create a three amino acid arrangement
similar to the H—X—D-Xss-H configuration of metal ligands in Group I enzymes, but
there are no good candidates for a residue corresponding to the conserved arginine in the
Group I motif. In addition, sequence comparisons between deA and more distantly
related Group II enzymes, such as y-butyrobetaine hydroxylase and clavaminate synthase,
do not show conservation of a histidine corresponding to His 167. Alignment of Group II
dioxygenase sequences indicates that His 262 is absolutely conserved and is in close
proximity to an invariant arginine which may be analogous to the OL-KG-binding arginine
in deacetoxycephalosporin synthase (DAOCS) and other Group I enzymes.

In this study, we used site-directed mutagenesis methods to examine the roles of
the all nine histidines and Asp l 15 in deA. Previously published work showed that deA
was inactivated by diethylpyrocarbonate, a histidine-selective reagent, and provided
evidence consistent with the presence of multiple histidines in the active site (7).
Spectroscopic studies of deA showed the presence of two equatorially—bound imidazole
nitrogens as ligands to the active site metal, and indicated that one imidazole ligand may
be displaced or shifted to an axial position upon substrate binding (3, 34). Based on
analyses of different deA variants, I identify several likely metal—binding residues and

provide evidence that another one or two histidines may aid in substrate coordination.

50

Experimental Procedures
Recombinant Plasmids

All plasmids were constructed from pUS311 (7), a pUCl9 derivative that contains
the Ralstonia eutropha Jl\/[P134 tfdA gene (Figure 4). The H8A, Dl 15A, H213A, H216A,
H235A, H245A, and H262A deA variants were created by direct mutation of tfdA in
pUS311 by the Stratagene Quickchange System (Stratagene, La Jolla, CA). All
mutagenic primers are listed in Table 4. Two alternative approaches were used to
construct the three remaining variants. Plasmids encoding H113A and H167A deA
variants were created by Clontech mutagenesis of pXHtfdA, a pUC19 plasmid containing
the 5’-XbaI-HindII fragment of the tfdA gene (Figure 4). To create the complete gene
containing the indicated mutations, the XbaI—HindII fragment was cloned into pHthdA,
which contains the 3’ end of the tfdA gene. pHthdA was constructed in two steps. First,
the 1.4—kb XbaI-Sall fragment from pUS311, containing the complete tfdA gene, was
cloned into pBC KS' (Stratagene) cut with XbaI and XhoI to create pBthdA. This step
had the benefit of eliminating a HindH site that interfered with further cloning steps. The
727-bp HindH-Kpnl fragment of pBthdA was subcloned into pBC KS‘ cut with the same
enzymes to give pHthdA. Similarly, the gene encoding H200A deA was made by
mutagenesis of pXthdA, a pUC19 plasmid containing the 5’-XbaI—Xhol fragment of
tfdA. The altered XbaI-XhoI fragment was then inserted into pHthdA cut with Xba I and

XhoI to give the complete H200A tfdA gene. The identity of

51

Figure 4. Restriction maps of the plasmids used in construction of tfdA mutants.

Abbreviations for the restriction enzyme sites are as follows: H, HindH; X, Xbal; Xh,
XhoI; Sl, SalI; K, Kpnl; So, Sad; N, Nrul. The vector backbone is indicated in the box on
the left. The tfdA open reading frame is shown as a wide, solid line.

 

 

x N H Xh N SIfH so
pUC19 I I I I I I pUS311
x N H Xh N SI/Xh
ch KS I I I I I I I pBthdA
X N H

x N H Xh
@' I '4 pXHtfdA
HthdA
ch KS I I I H p

 

 

Table 4. Sequences of mutagenic primers used to create altered tfdA genes

 

 

Mutant Primer Sequencea

H8A 5’- CGC AAA TCC CCT TGC TCC TCT TTT CGC C —3’
H113A 5’- GTC GCT GGC CCA AGC TGG-3’
D115A 5’—GCA CAG CGC CAG CTC CTT TCA-3’
H167A 5’-GCG TGC CGA GCA GTA CGC ACT G—3’
HZOOA 5’—GGT TCG AAC CGC CGC CGG CTC—3’
H213A 5’-GCT CGC GGC CGC GCC GAT-3’

H216A 5,-CCT TCG ACG GCG CTC GCG—3’

H235A 5,-GCT TCT CGA GGC GAC ACA G—3’
H245A 5’-GTG TAC CGG GCT CGC TGG AAC-37
H262A 5’—CGT TCT TGC ACG CGG ACG CAG—3’

dea—MBPF 5’—TCT CTA GAG TGA GCG TCG TCG CAA ATC C-3’
dea-MBPR 5’-GTC AAG CTT GGT TGC GTA CAT CTT GTG G-3’

21The reverse primer used for the creation of all mutants was the complement of the
forward primer.

52

all final constructs was confirmed by sequence analysis. To insert the genes encoding
H167A and H262A variants of deA into a plasmid that would allow for IPTG-controlled
expression, the XbaI—Sall fragments from the corresponding plasmids described above
were cloned into pET23a (N ovagen) prepared with the same enzymes.

To create the maltose-binding protein (MBP)-deA fusion proteins, the wild-type
tfdA gene was amplified from pUS311 with deA-MBPF and deA-MBPR primers
(Table 4) to create an Xbal site directly upstream of the GTG start codon of the tfdA gene

and a HindIII restriction site 54 bp downstream of the stop codon. The PCR product was

 

cloned directly into the pGEM-T vector according to the manufacturer’s instructions
(Promega, Madison, WI). The XbaI-HindIII fragment was isolated from the resulting
plasmid and cloned into the pMAL-cZ vector that had been digested with the same
enzymes. The identity of the newly created malE—tfdA gene fusion was confirmed by
sequencing. Substitution of the mutation-containing internal NruI fragment for the same
fragment of the wild-type malE-tfdA gene created MBP-fusion forms of altered deAs.
First, pMAL—tfa’A was digested with Nrul and the vector fragment was purified and
religated to create pMAL-t‘fa’AANruI. The resultant plasmid was linearized with NruI and
dephosphorylated with calf intestine alkaline phosphatase prior to ligation with the NruI
fragments isolated from the previously described mutant genes. Constructs were

confirmed by restriction analysis.

Protein Purification

H8A, H1 13A, D1 15A, H216A, H235A and wild—type deA proteins were purified
from E. coli DHSa cells carrying pUS311 and its mutated derivatives according to a

previously described protocol (7). In addition, the non-mutated enzyme and the deA

53

variants H167A, H200A, H213A, H245A and H262A were purified as MBP-deA fusion
proteins from E. coli DH50t by the protocol described in the pMAL Protein Fusion and

Purification System Manual (New England Biolabs, Beverly, MA).

Analysis of Kinetic Parameters

Specific activities of the wild-type and variant deA proteins were determined by
a previously described spectrophotometric assay (7). The typical assay mixture contained
1 mM 2,4-D, 1 mM Ot-KG, 100 uM (NH4)2Fe(SO4)2, and 100 uM ascorbic acid in 10
rnM MOPS buffer (pH 6.75) at 30°C. The reactions were quenched by the addition of
EDTA to a concentration of 5 mM. 2,4-Dichlorophenol was quantified by reaction with
4-aminoantipyrene followed by measurement of the absorbance at 510 nm. One unit of
activity was defined as the amount of enzyme required to produce one umol of
dichlorophenol-min". Protein concentrations were determined using the BioRad Protein
Assay with bovine serum albumin as a standard. For calculation of the kcat values, the
deA variants were assumed to have M, = 31,600 and the MBP-deA variants were
assigned Mr = 74,500.

The low Km for or-KG (~2-5 uM for the wild-type enzyme) precluded use of the
4-aminoantipyrene assay for accurate determination of this value. The alternative method
used to measure the Km for oc-KG quantified the amount of 14C02 liberated from a—[l—

14C]-KG during the course of the reaction (7).

54

Native Protein Analysis by Gel Filtration

Size exclusion chromatography was used to estimate the native molecular weights
of deA, MBP—deA, and mutant proteins. The proteins were chromatographed on a
Superose 6 gel filtration colurrm (1.0 x 30 cm, Pharmacia-Upjohn) in 20 mM Tris buffer
(pH 7.5), 1 mM EDTA, and 200 mM NaCl at a flow rate of 0.2 ml°min‘1. The elution
volumes were compared to those for gel filtration standards (Bio-Rad) including
thyroglobulin, 670 kDa; bovine gamma globulin, 158 kDa; chicken ovalbumin, 44 kDa;

myoglobin, 17 kDa; and vitamin B12, 1350 Da.

Spectroscopic Analysis

Proteins for electron paramagnetic resonance (EPR) and electron spin-echo
envelope modulation (ESEEM) spectroscopic analyses were exchanged into 25 mM
MOPS (pH 6.75) by repeated concentration and dilution in Centricon 30 (Amicon)
centrifugal concentrators. The final subunit concentration was 0.5 mM for the non—fusion
forms of deA and 0.4 mM for the MBP-deA proteins. CuClz was added to a
concentration of 450 uM and 350 uM, respectively. Buffered solutions of oc-KG and 2,4-
D were added to final concentrations of 5 mM. Glycerol was present at 40% in all
samples.

X—band EPR spectra were obtained at 77K on a Bruker ESP-300E spectrometer.
ESEEM data were collected on a home-built spectrometer; the microwave bridge of this
instrument has been previously described in detail (20). Data collection and analyses
were controlled by a Power Computing model 200 Power PC using software written with

LabView v. 5.01 (National Instruments). Electron spin echoes were digitized, averaged,

55

and integrated by a Tektronix model 620B digital oscilloscope interfaced to the
spectrometer computer via an [BEE-488 bus. Two four-channel delay and gate generators
(Stanford Research Systems model DG535), a Bruker BH-15 magnetic field controller,
and a Hewlett-Packard model 8656B radiofrequency synthesizer were also interfaced
using [BEE-488 protocol. Data were collected using a reflection cavity that employed a
folded rnicrostrip resonator (16). A three-pulse stimulated echo sequence (90°-t-90°-T-
90°) was used. ESEEM spectra were generated by Fourier transformation of the time—
domain data using dead time reconstruction (23). Simulations of the experimental data
were performed on a Sun SparcII work station. Simulation programs were written in
FORTRAN and based on the density matrix formalism developed by Mims (22).
Software for the frequency analysis of the experimental and simulated data was written in

Matlab (Mathworks, Natick, MA).

Results
Production of the mutant T fdAs

Initially, all of the mutant genes were expressed from their pUC19-based plasmids
except for tfdA genes encoding H113A, H167A and H200A deA, which were in pBC
KS‘-derived plasmids. Using the standard protocol to produce soluble, wild-type deA
(growth at 30°C to early stationary phase), only the H8A, H113A, D1 15A, H216A, and
H23 5A variants existed as soluble proteins. All of the other deA variants were present
as inclusion bodies even when grown at lower temperatures (22°C), in M9 minimal

medium, or in LB broth containing 660 mM sorbitol and 2.5 mM betaine (1). In addition,

56

IPTG-controlled production of H167A and H262A proteins from mutant genes cloned
into pET23a did not yield soluble samples even when the harvested cell pellets were
suspended in buffer containing 20% glycerol to limit protein aggregation.

To overcome the solubility problems for the five deA variants, MBP-deA
fusion proteins were created. Wild-type deA and the MBP—deA fusion protein had
essentially identical km and very similar Km values for oc-KG and 2,4-D (Table 5). A
slight increase in the apparent KD F e(II) may reflect some metal binding capacity of
MBP. Since the presence of the fusion protein did not appear to greatly affect the kinetic
parameters of wild-type enzyme, similar fusion proteins were created for the H167A,

H200A, H213A, H245A, and H262A deA variants.

Kinetic analyses of altered T fdAs

Results from kinetic analyses of the four active mutant proteins are summarized in
Table 5. H8A deA was soluble and active, but was rapidly proteolyzed to an inactive
form. By electrophoretic comparisons, the cleavage site appeared to be the same as in
wild-type deA (between Arg77 and Phe78) (7). The rate of proteolysis of H8A deA
was enhanced compared to that seen for the wild-type enzyme despite the presence of
EDTA and protease inhibitors in the purification buffer. Because purified H8A deA was
more than 75% degraded, the catalytic rate constant was calculated on the basis of the
estimated amount of intact enzyme. These calculations indicate rates and Km values
similar to those for the wild-type enzyme. Similarly, the kinetic parameters for H23 5A
deA were comparable to the native enzyme. In contrast, two variants exhibited

differences from wild-type enzyme in their kinetic parameters. The H213A MBP-deA

57

variant exhibited a twenty-fold reduction in kcat and a ten-fold increase in Km for 2,4-D. In
addition, H216A deA had a modest (2.5-fold) increase in the Km for 2,4-D and no
change in catalytic rate. The other kinetic parameters for HZ 1 3A MBP-deA and H216A
deA (Km for a—KG and KD for ferrous ion) did not differ significantly from the wild-type
values. Six soluble deA variants (H113A, D115A, MBP-H167A, MBP-H200A, MBP-
H245A and MBP-H262A) exhibited no activity even when assayed with elevated

substrate and cofactor concentrations (10 mM oc-KG, 5 mM 2,4-D, 250 uM Fe(H)).

Table 5. Summary of the kinetic parameters for active deA variants

 

 

deA Sample kcat Km aKG K.,, 2,4-D KD Fe(II)
min" 11M M M

wild-type 442 i 33 4.9 i 0.73 19.3 i 3.7 2.0 i 0.5
MBP-fusion 411 i 23 9.9 i 1.8 33.7 i 2.5 15.2 i 5.4
H8Aa 284i6.9 6.51L 1.8 17.7:38 1.9:r 0.7
MBP-H213A 22.1 i 2.9 8.3 i 2.9 318 i 44 27.7 i 9.8
H216A 474 i 47 6.05 i 2.6 52 i 5 2.4 i 1.7
H235A 284 i 25 9.6 i 1.6 34 i 8 1.4 :1.7

 

a More than 75% of the protein was present as the degradation product. The kcat was
estimated from the active, full-length fraction.
Evaluation of the Structural Consequences of the Mutations

To assess whether the inactive mutant proteins assumed conformations similar to
the wild-type enzyme, their apparent molecular weights were estimated by gel filtration
analysis. The observed size of wild-type deA was found to be 51 kDa by comparison to
protein standards, suggesting that deA forms a compact dimer or an elongated
monomer. The elution volume for both H113A and D115A corresponded exactly to wild-
type deA indicating that these proteins are not significantly altered in their quaternary
structure. MBP-deA eluted both in the void volume (approximately 25 % of the protein)

and at a position corresponding to 216 kDa (roughly 75 % of the protein), suggesting that

58

MBP-deA forms at least a dimer. Because each MBP-deA subunit is comprised of two
domains separated by a thirteen amino acid linker, the resultant protein may migrate with
a larger apparent molecular weight. MBP-H167A and MBP-H200A samples
{demonstrated the same two-peak profile as MBP-deA, but with larger proportions
eluting in the void volume. MBP-H245A and MBP-H262A proteins were soluble,
however, gel filtration analysis indicated the presence of only highly aggregated material
eluting in the void volume. Because these mutant proteins exhibited aberrations in their

folding properties, the catalytic role of His 245 and His 262, if any, could not be assessed.

EPR Spectroscopic Characterization of Variants with Altered Metal Sites

The deA metallocenter properties were examined by EPR spectroscopy. For the
purposes of these experiments, the diamagnetic Fe(II) metallocenter of deA was
substituted with spectroscopically active Cu(H) ion. Although the Cu(H) form of deA is
inactive, Cu(H) binds competitively with respect to F e(H) (K, = l-3 uM) and Cu(H)-
substituted deA has been used previously to study the metal coordination environment
of this enzyme in the presence and absence of substrates (3, 34). Spectral parameters of
wild-type Cu(H)-deA, Cu(H)-deA + oc-KG, and Cu(H)-deA + oc-KG + 2,4-D (Figure
5A and Table 6), agreed well with those reported previously (34). Earlier studies of
Cu(H)-substituted wild-type deA indicated that the metal was bound in a tetragonal
environment with a mixture of O and N ligands in the equatorial plane. Upon addition of
oc-KG and 2,4—D, the spectral parameters were altered to a more rhombic signal with
accompanying resolution of ligand hyperfine coupling (Figure 5A). These results suggest

that binding of the cofactor and substrate to the enzyme leads to a better-defined copper

59

Figure 5. X-band CW-EPR spectra of Cu(H)-substituted deA variants.

EPR spectra were obtained at 77 K for the following samples (substrate concentrations
were 5 leI, when present). A, (Top to bottom) Wild-type Cu-deA in the absence of
substrates, with or-KG, and with a—KG + 2,4-D. B, H113A Cu-deA in the absence of
substrates, with a-KG, and with a-KG + 2,4-D. C, D115A Cu-deA in the absence of
substrates, with oc-KG, and with oc-KG + 2,4-D. D, MBP-H167A Cu—deA in the absence
of substrates, with a-KG, and with oc-KG + 2,4-D. E, MBP-H200A Cu-deA in the
absence of substrates and in the presence of OL-KG + 2,4-D. Spectral parameters are
listed in Table 6.

60

Figure 5. X-band cw-EPR spectra of Cu(II) substituted deA variants

 

 

 

 

 

 

A wt deA ’/ gt Cu only
Cu+aKG
811
I Cu+aKG+2AJD

L__J

A11
2500 2750 3000 3250 3500 3750
Gauss
B H113A deA

 

 

:-

 

2500 2750 3000 3250 3500 3750
Gauss

 

 

 

C D115A deA Cu only

 

    
 
    

Cu+aKG

Cu+aKG+2A{)

2500 2750 3000 3250 3500 3750

Gauss

 

 

61

Figure 5. (con’t)

 

D H167A lVIBP-deA

Cu only

 

Cu + OtKG + 2,4-D

3250 3500 3750

 

 

E H200A IVIBP-deA

 

Cu + ot—KG + 2,4-D

2500 2750 3000 3250 3500 3750
Gauss

 

 

 

62

Table 6. Summary of the EPR spectral parameters for Cu(H)-substituted deA

 

 

 

 

 

 

variants.
811 A11 g1
mT
wild-type deA 2.34 16.3 2.07
wild-type deA/oc—KG 2.36 15.1 2.07
wild—type deA/a—KG/2,4—D 2.38 12.0 2.09
H113A 2.30 16.6 2.07
H113A/0L-KG 2.33 15.8 2.07
H113A/0t-KG/2,4-D 2.35 14.0 2.08
D115A 2.30 17.0 2.06
D115A/0t—KG 2.30 16.2 2.06
2.34 16.8 2.06
D115A/0t-KG/2,4-D 2.30 16.2 2.06
2.34 16.8 2.06
MBP—H167A 2.30 17.1 2.07
lVfBP-Hl67A/ot—KG/2,4-D 2.30 17.1 2.07
MBP-H200A 2.30 17.1 2.07
2.34 16.1 2.07
MBP-H200A/0t-KG/2,4-D 2.30 17.1 2.07
2.34 16.1 2.07

 

63

 

site in the enzyme (34) and the decreased values for A” upon addition of substrates (AH
=12.0 mT) indicated a significant distortion from planarity.

The effects of the amino acid substitutions on metallocenter structure in the deA
variants w1th quaternary structures s1milar to the corresponding wild— type protein
(H113A, D1 15A, MBP-H167A and MBP-H200A deA) were assessed by EPR
spectroscopy. Because there were no significant differences between the spectra of MBP-
deA and the non—fusion wild-type form of the protein (data not shown), H167A and
H200A deA were analyzed as MBP fusion. In all cases, EPR spectra of the Cu(II)—
substituted deA variants showed contributions from multiple copper species. The
mixture of copper centers may have resulted from either copper interactions in other
regions of the protein or the presence of multiple binding modes at the active site.

The EPR spectra for Cu—H113A deA (Figure 5B and Table 6) differed
significantly from spectra of wild-type enzyme. Irnportantly, these spectra showed clear
changes in Cu(II) g-values and hyperfine tensor principal values upon substrate additions
consistent with the binding of copper to the active site pocket. The broadening of the EPR
signal in the gH region upon addition of or-KG suggested a mixture of copper coordination
environments consistent with a less tightly bound metal site.

The EPR spectrum of D1 15A deA (Figure 5C) had parameters similar to those
observed in the copper-substituted wild-type protein (Table 6), although with less
resolution of A“. Addition of oc-KG (in the presence or absence of 2,4-D) had a dramatic
effect on the appearance of the D115A data, enhancing resolution of the Cu hyperfine
peaks at g“ and the ligand hyperfine structure in the g i region (Figure 5C). The

appearance of these superhyperfine interactions may have resulted from a subtle shift in

64

the orientation of the principal g—tensor such that the imidazole ligands occupied an
increasingly equatorial position (31). These results were consistent with the formation of
a tighter or more regular copper binding site upon addition of the cofactor. Again, the
perturbations of the spectrum upon a—KG addition were consistent with copper binding to
the active site.

MBP-H167A deA (Figure 5D) and lVIBP-H200A deA (Figure 5E) EPR spectra
were poorly resolved and showed evidence for multiple modes of copper binding.
Significantly, few changes were seen upon addition of either oc—KG or 2,4-D. The MBP—
H200A EPR spectrum (Figure SE) was more defined than that of H1 13A or MBP—
H167A, and clearly showed a second set of resonances of lower amplitude with
parameters identical to those observed for the copper-substituted wild—type enzyme
(Table 6). The broadness of the resonances in the MBP-H167A sample suggested that a

similar situation may occur in that enzyme as well.

ESEEM Spectroscopic Characterization of Variants with Altered Metal Sites

ESEEM spectroscopy was used to gain additional insight into the number of
nitrogen ligands that create the copper centers in the deA variants. As reported
previously, ESEEM analysis of wild-type Cu(H)—deA indicated the presence of two
equatorial histidines directly coordinated to the metal (34). The Fourier transformed 3-
pulse ESEEM data, collected near g“, for wild-type Cu(II) deA with oc—KG showed sharp
peaks at 0.6, 0.9, and 1.5 MHz and a broad feature at 3.5 MHz (Figure 6). This signature
is typical of copper bound by an equatorial imidazole. The narrow combination bands at

2.1, 2.5, and 3.1 MHz (Figure 6) indicated the presence of more than one histidine per

65

copper site (19, 20, 24). Simulations of the ESEEM data for the wild-type enzyme plus
a-KG, generated with the density matrix approach of Mims (22) and the parameters for
copper coordinated by two identical equatorial histidines, are superimposed on the
experimental data in Figure 6. ESEEM data for Cu(H)—deA in the presence of or—KG
plus 2,4-D were analyzed by a similar method (data not shown). A substantial decrease in
the modulation intensity and disappearance of the combination bands (34) upon addition
of both substrates is consistent with the loss of one histidine from the equatorial plane.

Similar analyses of Cu(H)—substituted deA variants were used to compare their
metallocenter structure to that of the wild—type enzyme. Three pulse ESEEM patterns
were generated for the H113A, MBP-H167A, and D115A variants of deA (Figure 7) as
well as for the MBP-H200A samples (data not shown). To normalize the data, the
amplitudes in the spectra for all four mutants were corrected to account for differences in
baseline frequencies. Samples for each deA variant gave rise to low frequency
modulations indicative of equatorially—bound histidine ligands. The ESEEM spectra from
H113A (Figure 7A) and MBP-H200A (not shown) variants were nearly identical and
show modulations that were considerably weaker than those measured for MBP—H167A
(Figure 7B) and D1 15A (Figure 7C). As for wild-type spectra, the data were simulated
with a program based on Mims' density matrix formalism, assuming a system of spherical
symmetry. In each case, the ratio of modulated signal intensity to unmodulated signal
intensity was suggestive of copper bound in more than one coordination environment.
The differences in strength of copper binding to the active site can be visualized in Figure
7 by comparing of the peak heights of a given spectrum (modulated frequency) to the

background signal that is observed at the end of data collection upon shift to the direct

66

 

 

 

 

 

 

 

 

 

 

0.8
8 0.6’ ,
3
E- 0 4
< ' — _
0.2 i
0 — 1
12
l l l I I I I l l l l
a)
‘C
3
E _
< /\ 7
'_ -
u.
.I \ NINA/WW V‘s/Ivar, BWMi/LMW‘KV
‘-'. . . l . i . 1 i . i l . . . I i _
0 2 4 6 8 10
frequency (MHz)

Figure 6. 3-Pulse ESEEM spectra of Cu-substituted wild-type deA with 2-histidine
simulation data.

ESEEM spectrum of wild-type Cu(H)-deA + or-KG (solid) and computer—simulated data
for Cu(H) bound by two identical histidines (dashed) (A) and the corresponding
frequency domain spectrum (B). Parameters for data collections were magnetic field =
2800 G, t = 375 ns, T = 50 ns, v = 8.98 GHz; 4.2 K. Cu hyperfine parameters: g“: 2.05,
gz=2.34, Axx=Ayy=20 MHz, A2: 487 MHz, Superhyperfine parameters: gN = 0.40347,
A150: 1.7 MHz,radius= 2.75 A, [0,¢]=[2.37,2.07] radians, e2qQ = 1.60 MHZ, n = 0.75,
NQI [0t,[3,y]=[ .50,1.1,0.1] radians. The simulated data were treated with an exponential
((_rs+ T‘)
3500
background decay fimction was applied to allow the amplitudes of the initial 1.5 usec of
the simulations to match the data.

decay fiinction to give y'jj = (y — 0. 17) * exp + 0.17 For each simulation, 3

67

current (unmodulated frequency). Although this ratio was less than the wild-type value
for all of the deA variants, H113A and MBP-HZOOA had a markedly decreased capacity
to constrain copper to one geometry in the absence of substrates.

Fourier transformations of the data shown in Figure 7 yielded major features at
0.7, 1.5, and 4.0 MHz indicative of histidines equatorially coordinated to Cu(H) (24).
Although Cu—bound histidines were visible in the H113A and MBP-H200A spectra, the
ESEEM signals were too weak to be consistent with one dominant Cu(II) conformation,
and thus were omitted from further analyses. The frequency and Fourier transformed
data, with the corresponding simulations, are shown for D115A (Figure 8) and MBP-
H167A deA (Figure 9). In both cases, the large unmodulated component of the data
required a severe decay function in order to simulate the data. In addition, it was
necessary to use the average of several simulation data sets with varying isotropic
hyperfine coupling constants to compensate for the presence of multiple copper binding
environments identified by EPR spectroscopy. Although the combination lines were
weaker in the data for D115A than for wild-type deA, the combination line at
approximately 3.0 MHZ was observed and its frequency was predicted nicely by the 2 His
simulation data. Thus, these data were most consistent with two histidine ligands to the
Cu(II) site in this protein. The broadness of the Fourier transformed spectrum for D115A
deA and the subsequent difficulty in observing the combination lines most likely reflects
a distribution of 14N superhyperfine couplings from multiple Cu(H) centers in D115A
deA which is consistent with our use of an average of several isotropic hyperfine

coupling constants to simulate the data.

68

 

   

 

 

 

 

 

I I I I I I I I I l I I I I l l l I I I I

(”Va ....

M.,, A -

"MW”r-rM»Mimrwm. _

s A -
3

E \ ,va 4

8‘ IIW/W W., W’W'I‘rtw B __

< WWW I

8 ' IIWflIntrIW/sim WWW “twnmwn ‘

.5 ” “W WWW -‘\ ‘

To 0. 4 -- IF WW WWI -

E - c -

o _ _

z - __

0. 2 -— _

O _ l l l i l l l i 1 l l i l l l l l l i L:

0 2 4 6 8 10
r+T (usec)

Figure 7. 3-Pulse ESEEM time-domain spectra of deA variants.

Spectra for H113A Cu-deA (A) (magnetic field = 3050 G, r = 300 ns, T = 40 ns); MBP-
H167A Cu-deA (B) (magnetic field = 3100 G, r = 300 ns, T = 40 ns) and D115A Cu-
deA (C) (magnetic field = 3050 G, r = 385 ns, T = 55 ns, v = 8.8 GHz) are compared.
All were collected at 4.2 K.

69

 

 

 

 

 

 

 

 

_ , I'IT , I I I I l I I l I I I I I I I I _
a) j I I
go.8_Il,III A —
-*—'—' ” 1 1 I: I.‘ -
a i III! I. . —
E 0-6 _— III: 1,// \ ”V” j
'C) _1IfIII1 ‘/ ,\ 143% ““WWMN.W W
a 04 _— I ., -—--wwmm —_
E I
i. 0.2 '— L
O _ 4
Z — I
0 — l l l I I l I I I. L I 1 l L I l 1 1 I
0 2 8 1O
T+T(IlS€C)
1O LI I I I I I I I I I I 1
+51 -
8 3}, B ‘
EI ‘; E
#3 'u III/L ,I, d
a 4 i 2,, i
E " 1 1'. , -
< 2 -_':;\/I I ,11 /~, g
E :I/I '1 I Inc? ,/\., 3
o 3' :. a... / Wwww
_ \. ‘f’.‘ —
-2 _ 1 e 1 I I I ‘1 I J I 1 I i 1 1 I 1 i 1
0 2 4 6 8 10
Frequency (MHz)

Figure 8. Spectral simulation of ESEEM data for D115A-Cu-deA.

Experimental (solid line), computer-simulated l-His (dashed line), and simulated 2-His
(dotted line) (A) and the corresponding time—domain spectra (B) for copper-substituted
D115A deA are shown. Magnetic Field = 3050 G, ‘C = 230 ns, T = 50 ns G, v = 8.8 GHZ,
4.2 K, gN = 0.40347, eZqQ = 1.55 MHz, 11 = 0.87, Axx = 1.6 MHZ, Ayy = 1.9 MHz, An =
2.2 MHz. The simulated data were treated with an exponential decay function to give

. " (1+ T) 0'8
y” = (y, — 0.05) * exp ( 600 — 0.05.

 

70

_\

 

 

 

 

 

 

 

 

 

 

... I I I I I I I I I I I I I l I I I I _
3 O 8 ’ ’IIIIII/i 'I A ‘
—Q. HI". , I IN _
' .' ‘-V..\ i ‘ e ‘
8 — “I -
.E 0.4 _ i _
E : I :
§02 _ I _
Z I I
0 — I l I I I I I I I I I I l I I I I I in ~
0 2 4 6 8 1O
T+T (usec)
I III I I I I I I I I I I I I I I I I I I I I I I Til Ti r1 I—I T I I I r IE
5:? B 3
a) ; J; ' '- . 1
g I III/I: I ,r/IPI‘“\ _f
< . II I r '- 3
I— II Wig .. / \ :
LL 1 I I! . 2 7 w. / \\ “ ”ca/‘JA’m/Vi
II /( II/i \4/ a wwwwf \/ i
l l‘\‘11/- I4 I . I I I . I I I . é . #44} . I . . l I I E
0 1 2 3 4 5 6 7 8

Frequency (MHz)

Figure 9. Spectral simulation of ESEEM data for Cu(II) MBP-H167A deA.

Experimental (solid line), computer-simulated l-His (dashed line), and 2—His (dotted line)
time -domain spectra (A) and the corresponding Fourier transformed data (B) are shown.
Magnetic field = 3100 G, I = 300 ns, T = 40 ns, V = 8.8 GHz, 4.2 K, gN = 0.40347, eZqQ
= 1.60 MHZ, n = 0.82, AXX = 1.55 MHz, Ayy = 1.9 MHz, An = 2.0 MHz. The simulated
data were treated with an exponential decay function to give

(I + D)“

y'” = (yfl — 0.03)*eprW —0.03.

71

Simulations of the MBP-H167A deA also suggested the presence of two
histidines per copper (Figure 9). The peak broadness in the experimental data resulted in
our inability to resolve the features at 0.6 and 0.9 MHz, indicative of a heterogeneous
population of copper sites. Simulations achieved by averaging of multiple isotropic
hyperfine coupling constants, as in those for D1 15A, predicted the coalescence of these
peaks. The number of averaged data sets required to simulate the MBP-H167A spectra
indicated that the copper center in this protein was more disordered than that in D1 15A
deA. The increased number of unconstrained copper binding environments in MBP-
H167A deA may have resulted from non-specific binding to the MBP domain or a small
population of misfolded deA protein, as observed in gel filtration experiments.

The Cu-substituted forms of H1 13A, D115A, MBP-H167A, and MBP-H200A
deA species were also analyzed in the presence of oc-KG and 2,4—D. Although the Cu(H)
site in wild-type deA appeared to undergo significant rearrangement upon binding of
substrates, only slight variations in the lineshape of the higher frequency peak at ~4.0
MHz were observed for H1 13A, MBP-H167A, and MBP-His 200A. The substrate-
dependent changes in the copper site of H1 13A deA upon substrate addition that were
observed by EPR spectroscopy were not well defined in the ESEEM spectra, again
indicating a less constrained copper site. Addition of a-KG and 2,4-D to D115A deA
led to enhanced definition of the D1 15A copper site similar to what was observed by
EPR analysis (Figure 5). The ternary complex of Cu(H)-D1 15A deA exhibited
coherence in the frequency data at longer times with a slightly increased depth of
modulation. These changes were not accompanied by a significant increase in

unmodulated intensity (Figure 10) indicating that the copper was predominantly found in

72

 

 

 

 

 

 

 

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8 — /\ I I
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Z 3 I :
F— l J I I l I J I I l l I l I l I l I L I I-.L
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I I T I I I I I I I l I ' I
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FT Amplitude

 

 

lllllll

 

 

Frequency (MHz)

Figure 10. 3-pulse ESEEM of Cu(II) D115A T fdA (4.2K).

(Top to bottom) ESEEM spectra for Cu(H) D1 15A deA in the absence of substrate, with

5 mM a—KG, and with 5 mM a-KG + 5 mM 2,4-D (A) and the corresponding Fourier
transformed data (B) are shown. Conditions for these measurements were identical to
those of Figure 6.

73

l
...

 

 

a single environment. ESEEM spectra of the D115A deA ternary complex showed
resolution of all three combination lines from the electron spin manifold that gave rise to
the intense low-frequency peaks (Figure 10B). These data provided convincing evidence
for two equatorial histidine ligands to the copper in D1 15A deA in the presence of
substrates and supported the conclusions drawn from substrate—free Cu (H)-Dl 15A deA
ESEEM spectra. The enhanced intensity and definition upon substrate binding suggested

that OI-KG constrained the geometry of the copper at the active site.

Discussion

Our results demonstrate the importance of multiple histidines and one aspartate
for deA activity. EPR, ESEEM, and extended x—ray absorption fine structure
spectroscopies of both the Fe(H)- and Cu(H)-forms of wild-type deA had previously
indicated that the resting enzyme has two imidazoles and a mixture of bound nitrogens
and oxygens as ligands to the metal (3, 34). Changes in the Cu-EPR spectral parameters
upon addition of OL-KG are consistent with an increased number of oxygen ligands to the
metal, but ESEEM and X-ray absorption data indicated that both imidazole ligands are
retained. Addition of 2,4-D, however, leads to the displacement or g-tensor reorientation

of a histidine out of the equatorial plane (3, 34).

Mutation of His 1 13 or Asp 1 15 to alanine both eliminates activity and greatly
alters the structure of Cu(H) metallocenter site. Significantly, however, substrate binding
perturbs the spectra of these sites consistent with the metal binding to the active site
pocket. These residues comprise the H-X-D motif which is conserved in enzymes related

to deA (Appendix A) and common to all other known oc-KG-dependent dioxygenases

74

 

(2) (Chapter 1). Site-directed mutagenesis studies analogous to those described here
demonstrated the importance of this motif in a number of Group I enzymes including
aspartyl, lysyl, prolyl, and flavanone hydroxylases (15, 18, 21, 25, 27). Crystal structures
of IPNS and DAOCS established these residues as ligands to the metal. My studies
support the proposal that residues in the H—X—D motif are ligands to the metal in deA

and, most likely, other Group H enzymes.

Cu(H)-D115A deA ESEEM spectra indicate the presence of two imidazole
ligands, whereas similar analyses of the H1 13A proteins shows evidence of multiple
copper binding environments. Clearly, the absence of the aspartate ligand results in a
significantly altered binding site, producing a less structurally-defined copper geometry,
and contributes to the increased damping of the ESEEM. Addition of oc-KG to this
mutant, however, results in increased resolution in the EPR spectrum and ESEEM
pattern, indicating that the binding of this cofactor locks the geometry of the active site
into place. Together, these data may suggest that the histidines are primarily responsible
for Cu(H) binding, and Asp 115 may contribute more to the orientation of the metal. The
properties of copper predict a much stronger interaction with nitrogen rather than oxygen
ligands; in fact, there are no mononuclear copper enzymes known to have a carboxylate
metal ligands (13). Because of differences in the preferred environments of Fe(H) and
Cu(H), it is difficult to draw conclusions about the specific electronic role of Asp 115 in

the native Fe(H) deA based on these observations.

The identity of the second metal binding histidine, predicted by various
spectroscopic studies, is less clear. A previously described motif (H-X-D—Xss-H-XIO-R-

X-S) common to IPNS and related enzymes (2, 18) would best match His 167 in deA

75

except that the terminal arginine is displaced. Spectral simulations of H167A, however,
are consistent with the presence of two imidazole nitrogens per copper in this protein.
Furthermore, His 167 is replaced with an arginine in the deA sequence from pH B in
Burkholderia cepacia (Appendix A). Thus, it is unlikely that His 167 is one of the three
metal ligands necessary to create the metallocenter. EPR and ESEEM analyses of H200A
deA suggest that His 200 is important for creation of the metallocenter. Although His
200 is conserved among the enzymes with high identities to deA, such as TauD from E.
coli and sulfonate/a-KG dioxygenase from yeast, a corresponding histidine is not found
in more distantly related Group H enzymes such as y-butyrobetaine hydroxylase or
clavaminate synthase (Appendix A). Mutational analysis of prolyl 4-hydroxylase (25)
provided evidence for a non-ligand histidine at active site that is important for a—KG
binding and controlling the decarboxylation of OI-KG in the absence of substrate. Perhaps
His 200 serves a similar function in deA providing an explanation for the changes in
active site structure upon its alteration. Sequence comparisons of all Group H enzymes
reveal that His 262 in deA is the only other invariant histidine and I suggest that this is
most likely the third metal—binding residue. Supporting my assessment of His 262 as a
metal ligand, this residue lies ten amino acids away from Arg 273 creating an H—XIo-R
motif similar to that in DAOCS and related enzymes (18). It is surprising, however, that
the H262A variant cannot fold properly and forms inclusion bodies when overproduced
in E. coli. For comparison, wild-type deA is routinely purified in its apo—enzyme form
and deA activity is immediately detected upon addition of Fe (11). Together, these data
suggest a more general motif for a—KG-dependent dioxygenases, H-X-D/E—Xn-H-XIO_13-

R (the carboxylate occurs as a glutamate in clavaminate synthase, see Appendix A),

76

 

 

where Xn varies with the different subgroups within the superfamily (Group I, n=44-65;

Group [I n=123-153; Group III, n=74—104).

His 213 and His 216 may also be present at or near the deA active site. MBP—
H213A and H216A deA variants had increased Km values for 2,4-D and the former
enzyme also demonstrated a decreased catalytic rate. These data may indicate that His
213 (perhaps also assisted by His 216) facilitates 2,4-D binding by electrostatic
interactions between the imidazole nitrogen and the substrate carboxylate group;
however, additional studies are required to test this hypothesis. The importance of the
2.4-D carboxylate for binding to deA is indicated by the inability of phenoxyethanol to
serve as a substrate (7). Neither His 213 nor His 216 are conserved among other related
oc-KG—dependent dioxygenases. The divergence of sequences in this region may reflect

differences in the residues required to bind substrates unique to each enzyme.

Our results provide no evidence to indicate that His 8, His 235, or His 245 are
critical for catalysis. The H8A variant exhibits enhanced protease sensitivity, but the full-
length protein possesses near wild-type activity. Similarly, the H23 5A protein is nearly
unaffected in its activity. Neither of these residues is conserved in homologous
sequences. As described for the H262A deA variant, His 245 appears to be structurally
important. Because H262A and H245A derivatives of deA exist in non—native, highly
aggregated conformations, we were unable to assess whether these residues may also
play a catalytic role. Although His 245 is not conserved in these sequences, it is in the C-
terminal half of the protein which in other a—KG—dependent dioxygenases show a higher
level of similarity than the overall sequence comparisons (12). This region might be more

constrained for structural reasons.

77

The H—X—D/E-Xn-H motif, which is found in all known oc-KG dependent
dioxygenases, is an example of the more general structural arrangement (2- His-1-
carboxylate) found in non-heme Fe (H) enzymes including extradiol dioxygenases and
pterin-dependent hydroxylases (9). It is proposed that these three ligands anchor the iron
in the active site of the enzyme while leaving three open coordination sites available for
the binding of substrate, cofactor and oxygen. Several studies suggest that 02 reacts much
more readily with such iron centers after the displacement of solvent molecules by more
electronegative substrate molecules (9). In addition, it is the diversity of exogenous
ligands that enable non-heme Fe(H)-dependent oxygenases to catalyze such a wide range

of reactions.

Our proposed model of the active site of deA is shown in Figure 11. His 113,
Asp l 15 and a third histidine tentatively identified as His 262 are shown as ligands to the
metal. As in the crystal structures of both DAOC S and IPNS, water molecules are
thought to occupy the three remaining coordination sites in substrate-free enzyme. The
model includes a likely binding mode for oc-KG based on the DAOCS crystal structure
(3 3). Recent spectroscopic studies of clavaminate synthase (26), TauD (32) and deA
(10) are consistent with a similar mode of oc—KG binding in these enzymes. We have no
evidence to suggest direct binding of 2,4-D to the metal center, and indicate the presence
of a separate 2,4-D binding site adjacent to the metal in the model. Mutation of His 213
affects both substrate binding and the rate of catalysis so I show this residue at the active
site to indicate potential substrate interaction, but its precise role is still to be defined. In
the absence of a crystal structure, the site—directed mutagenesis, metallocenter

spectroscopy, and kinetic analysis studies described here provide the most complete

78

/ NH
U
+
\\\ )0 O
\‘ O
2000 2 >7Asp115
-ooc l /‘0

o—Fe(n)——N

/ \ His113

000- N» n
:k NH
His 262

Figure 11. Model for the active site of deA.

The three residues proposed to ligate the ferrous ion of deA are shown along with a
possible coordination mode for the substrates, oc-KG and 2,4-D. His 213, implicated in
binding 2,4-D, is also illustrated.

79

 

 

working model of the deA active site. This model begins to define the necessary
elements for catalysis in Group H oc-KG-dependent dioxygenases and finds similarities

between these enzymes and other well-studied representatives of this superfamily.

80

Literature Cited

Blackwell, J. R., and R. Horgan. 1991. A novel strategy for the production of a
highly expressed recombinant protein in an active form. FEBS 295:1-12.

Borovok, I., O. Landman, R. Kreisberg-Zakarin, Y. Aharonowitz, and G.
Cohen. 1996. Ferrous active site of isopenicillin N synthase: genetic and
sequence analysis of the endogenous ligands. Biochemistry 35:1981—1987.

Cosper, N. J., C. M. V. Stfilhandske, R. E. Saari, R. P. Hausinger, and R. A.
Scott. 1999. X—Ray absorption analysis of Fe(H) and Cu(II) forms of an
herbicide-degrading OL-ketoglutarate dioxygenase. J. Biol. Inorg. Chem. 4:122—

De Carolis, E., and V. De Luca. 1994. 2—Oxoglutarate—dependent dioxygenases
and related enzymes: biochemical characterization. Phytochem. 36:1093—1 107.

Eichhorn, E., J. R. van der Ploeg, M. A. Kertesz, and T. Leisinger. 1997.
Characterization of oc—ketoglutarate-dependent taurine dioxygenase from
Escherichia coli. J. Biol. Chem. 272:23031-23036.

Fukumori, F., and R. P. Hausinger. 1993. Alcaligenes eutrophus JMP134 "2,4-
dichlorophenoxyacetate monooxygenase" is an oc-ketoglutarate-dependent
dioxygenase. J. Bacteriol. 175:2083-2086.

Fukumori, F., and R. P. Hausinger. 1993. Purification and characterization of
2,4-dichlorophenoxyacetate/oc-ketoglutarate dioxygenase. J. Biol. Chem.
268:24311-24317.

Hedden, P. 1992. 2—Oxoglutarate-dependent dioxygenases in plants: mechanism
and function. Biochem. Soc. Trans. 20:373-377.

Hegg, E. L., and L. Que Jr. 1997. The 2—His-l—carboxylate facial triad. Eur. J.
Biochem. 250:625-629.

Hegg, E. L., A. K. Whiting, J. McCracken, R. P. Hausinger, and J. Que, L.
1999. The herbicide-degrading oc-keto acid enzyme deA: metal coordination
environment and mechanistic insights. Biochemistry In press.

Hogan, D. A., T. A. Auchtung, and R. P. Hausinger. 1999. Cloning and
characterization of a sulfonate/Obketoglutarate dioxygenase from Saccharomyces
cerevisiae. J. Bacteriol. 181:5876-5879.

Jia, S., K. McGinnis, W. J. VanDusen, C. J. Burke, A. Kuo, P. R. Griffin, M.
K. Sardana, K. O. Elliston, A. M. Stern, and P. A. Friedman. 1994. A fully

81

20.

21.

22.

active catalytic domain of bovine aspartyl (asparaginyl)—B-hydroxylase expressed
in Escherichia coli: characterization and evidence for the identification of an
active-site region in vertebrate—OL-ketoglutarate-dependent dioxygenases. Proc.
Natl. Acad. Sci. 91:7227—7231.

Karlin, 8., Z. Y. Zhu, and K. D. Karlin. 1997. The extended environment of
mononuclear metal centers in protein structures. Proc. Natl. Acad. Sci. U S A
94: 14225—14230.

Kivirikko, K. I., and T. Pihlajaniemi. 1998. Collagen hydroxylases and the
protein disulfide isomerase subunit of prolyl 4—hydroxylases. Adv. Enzymol.
Relat. Areas Mol. Biol. 72:328-398.

Lamberg, A., T. Pihlajaniemi, and K. I. Kivirikko. 1995. Site-directed
mutagenesis of the on subunit of the human prolyl 4-hydroxylase. J. Biol. Chem.
270:9926-9931.

Lin, C. P., M. K. Bowman, and J. R. Norris. 1985. J. Mag. Res. 65:369-374.

Lloyd, M. D., H. J. Lee, K. Harlos, Z. H. Zhang, J. E. Baldwin, C. J.
Schofield, J. M. Charnock, C. D. Garner, T. Hara, A. C. Terwisscha van
Scheltinga, K. Valegard, J. A. Viklund, J. Hajdu, I. Andersson, A.
Danielsson, and R. Bhikhabhai. 1999. Studies on the active site of
deacetoxycephalosporin C synthase. J. Mol. Biol. 287:943—960.

Lukacin, R., and L. Britsch. 1997. Identification of strictly conserved histidine

and arginine residues as part of the active site in Petunia hybrida flavanone 3B-
hydroxylase. Eur. J. Biochem. 249:748-757.

McCracken, J., J. B. Cornelius, and J. Peisach. 1989. Quantitative aspects of
nitrogen-14 electron spin-echo envelope modulation in Cu(H)-proteins, p. 155-
161. In C. P. Keijzers, E. J. Riejerse, and J. Schmidt (ed.), Pulsed EPR: A New
Field of Applications. North Holland Publishers, Amsterdam.

McCracken, J., D.-H. Shin, and J. L. Dye. 1992. Pulsed EPR studies of
polycrystalline cesium hexamethyl hexacyclen sodide. Appl. Mag. Res. 3:305—
316.

McGinnis, K., G. M. Ku, W. J. VanDusen, J. Fu, V. Garsky, A. M. Stern, and
P. A. Friedman. 1996. Site-directed mutagenesis of residues in a conserved

region of bovine apartyl (asparaginyl) B-hydroxylase: evidence that histidine 675
has a role in binding Fe+2. Biochemistry 35:3957-3962.

Mims, W. B. 1972. Phys. Rev. B: Solid State 5:2409-2419.

82

 

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

Mims, W. B. 1984. Elimination of the dead-time altifact in electron spin—echo
envelop specta. J. Magn. Res. 59:291-306.

Mims, W. B., and J. Peisach. 1978. The nuclear modulation affect in electron

spin echoes for complexes of Cu2+ and imidazole with 1‘4N and 15N. J. Chem.
Phys. 69:4921-4930.

Myllyharju, J ., and K. I. Kivirikko. 1997. Characterization of the iron- and 2-
oxoglutarate-binding sites of human prolyl 4—hydroxylase. EMBO J. 16:1 173—
1180.

Pavel, E. G., J. Zhou, R. W. Busby, M. Gunsior, C. A. Townsend, and E. 1.
Solomon. 1998. Circular dichroism and magnetic circular dichroism
spectroscopic studies of the non-heme ferrous active site in clavaminate synthase
and its interaction with a—ketoglutarate cosubstrate. J. Am. Chem. Soc. 120:743—
754.

Pirskanen, A., A.-M. Kaimio, R. Myllylfi, and K. I. Kivirikko. 1996. Site-
directed mutagenesis of human lysyl hydroxylase expressed in insect cells. J.
Biol. Chem. 271:9398-9402.

Prescott, A. G., and P. John. 1996. Dioxygenases: Molecular structure and role
in plant metabolism. Annu. Rev. Plant Physiol. Plant Mol. Biol. 47:245-271.

Roach, P. L., I. J. Clifton, V. Fiiltip, K. Harlos, G. J. Barton, J. Hajdu, I.
Andersson, C. J. Schofield, and J. E. Baldwin. 1995. Crystal structure of
isopenicillin N synthase is the first from a new structural family of enzymes.
Nature 375:700-704.

Roach, P. L., I. J. Clifton, C. M. H. Hensgens, N. Shibata, C. J. Schofield, J.
Hajdu, and J. E. Baldwin. 1997. Structure ofisopenicillin N synthase complexed
with substrate and the mechanism of penicillin formation. Nature 387:827—830.

Rotilio, G., A. F. Agro, L. Calabrese, F. Bossa, P. Guerrieri, and B. Mondovi.
1971. Studies on the metal sites of copper proteins: ligands of copper in
hemocuprein. Biochemistry 10:616-621.

Ryle, M. J., R. Padmakumar, and R. P. Hausinger. 1999. Stopped-flow kinetic
analysis of Escherchia coli taurine/OL—ketoglutarate dioxygenase: interactions with
a—ketoglutarate, taurine, and oxygen. Biochem. In press.

Valegard, K., A. C. T. van Scheltinga, M. D. Lloyd, T. Hara, S. Ramaswamy,
A. Perrakis, A. Thompson, H.-J. Lee, J. E. Baldwin, C. J. Schofield, J. Hajdu,
and I. Andersson. 1998. Structure of a cephalosporin synthase. Nature 394:805-
809.

83

 

34.

Whiting, A. K., L. Que Jr., R. E. Saari, R. P. Hausinger, M. A. Fredrick, and
J. McCracken. 1997. Metal coordination environment of a Cu(H)-substituted Ot-
keto acid-dependent dioxygenase that degrades the herbicide 2,4-D. J. Amer.
Chem. Soc. 119:3413-3414.

84

 

CHAPTER 3
CLONING AND CHARACTERIZATION OF A SULFONATE/oc-
KETOGLUTARATE DIOXYGEN ASE FROM Saccharomyces cerevisiae

This chapter is based largely on the published article: Hogan, D.A., T.A.

Auchtung, and R. P. Hausinger (1999) “Cloning and characterization of a sulfonate/0L-
ketoglutarate dioxygenase from Saccharomyces cerevisiae.” J. Bacteriol. 181:5876-5879.

85

 

Introduction

The Saccharomyces cerevisiae genome sequence project revealed an open reading
frame, YLL057c, that encodes a protein with 27 % identity to the herbicide-degrading
enzyme deA (5) and 31.5 % identity to taurine/oc—KG dioxygenase, TauD (4). The
product of YLL057c has lower levels of amino acid sequence identity to other a—KG-
dependent dioxygenases such as y-butyrobetaine hydroxylase and clavaminate synthase.
Alignment of the deduced amino acids from YLLOS 7c to these four sequences shows
conservation of the H-X-D and H-X—R motifs that are signatures for the Group H subset
of the a—KG-dependent dioxygenase superfamily that I defined in the Introduction of this
thesis. The presence of these residues, which were proven to be important in the site—
directed mutagenesis studies of deA described in Chapter 2, provides strong evidence
that the product of YLL057c is an OL-KG-dependent dioxygenase.

Given the nearly equivalent levels of identity between deA, TauD, and the
putative yeast dioxygenase, it is impossible to predict the natural substrate for the yeast
enzyme. Although there are no reports of 2,4-D degradation in S. cerevisiae, yeast may
contain an enzyme involved in the decomposition of other naturally occurring aromatic
compounds such as aromatic amino acids or lignin degradation products. Alternatively,
the YLL05 7c product may resemble TauD in its ability to decompose taurine, a naturally
occurring sulfonate (6). A number of yeast can use aliphatic sulfonates, such as taurine,
cysteate and isethionate, as alternative sulfur sources (17). Sulfonate utilization by S.
cerevisiae only occurs under aerobic conditions, is independent of sulfate utilizing
enzymes, and requires sulfite reductase, consistent with the formation of sulfite prior to

assimilation. This pattern of sulfonate utilization is similar to that in Escherichia coli and

86

 

 

other enteric bacteria (16, 18), suggesting the presence of a common pathway in these
diverse organisms. In E. coli, TauD hydroxylates the carbon atom in the C-S bond of
taurine to give an unstable intermediate that spontaneously decomposes to sulfite and
aminoacetaldehyde (4). There are no reports of an analogous enzyme in fungi.

In this chapter, I report both the characterization of the S. cerevisiae YLL05 7c
gene product and studies with the YLL05 7c deletion mutant to determine whether this
enzyme plays a role that is analogous to deA in 2,4-D degrading microorganisms or to

TauD in E. coli.

Materials and Methods

Cloning and expression of YLL05 7c.

Open reading frame YLL057c (Genbank Accession # 273162) was PCR
amplified and cloned into pET23a for expression in E. coli. The 1236-bp YLL057c
sequence, which contains no introns, was amplified from XPM-5392 (ATCC) containing
a 30-kb fragment of S. cerevisiae chromosome XH, with primers 5 ’-CAT ATG TAC
AGA GGA CGT CGT CGA G -3’ and 5’-CTA CAA CAC TTT TCG TCT CCG AGG-
3 ’. The forward primer contained a 5’ extension that introduced an NdeI restriction site

directly upstream of the start ATG codon. Template DNA was added by transferring a
small amount of material from a lPM-5392 plaque, obtained by plating on E. coli Y1090
(13), directly to the PCR reaction mix. The PCR reaction mix included 20 mM Tris-HCl

(pH 8.4), 50 mM KCl, 1.5 mM MgC12, 25 uM dNTPs (Gibco), 1 uM of each primer and

0.05 U Taq polymerase (Gibco). The temperature program was comprised of an initial

denaturation step at 95°C for 2 min 10 sec, 35 cycles of a denaturation step at 95°C for 1

87

min, an annealing step at 55°C for 1 min, a 2 min 10 sec extension step at 68°C, and a
final extension at 68°C for 6 min. The resulting 1734-bp PCR product (which included
additional downstream sequence) was cloned into pCR2.l TOPO vector (Invitrogen)
according to the manufacturer’s instructions to give plasmid pCR2.1—YDO. The cloned
gene was cut from pCR2.1-YDO with NdeI and SacI and inserted into pET23a cut with
the same enzymes to create pBlO. To create an N—terminal fusion of the YLL057c gene
product to the maltose-binding protein (MBP), YLL057c was amplified from pBlO using
the protocol described above with primers 5’-TCT CTA GAA TGT CTC CTG CAG
CAG C —3’ and 5’-GTC AAG CTT AAA GAA GTG TTG TCG CCG—3’. The forward
primer introduced an Xbal site directly upstream of the start site for in—frame insertion
into pMAL-c2 (New England Biolabs). The amplification product was directly cloned
into pGEM-T (Promega) to give pGTYDOlé. YLL057c was cut from pGTYDOl6 with
XbaI and HindHI, and ligated to pMAL-c2 prepared with the same enzymes to create

pMBPYDO, a fusion of the malE gene to YLL057c.

Production and purification of YLL05 7c gene product.

Various efforts to directly express the yeast gene in E. coli DHSOL from pBlO
failed to achieve high level production of the desired protein. In contrast, a MBP fusion
of this product was produced in E. coli DH50L(pMBPYDO), consistent with an increased
stability and decreased toxicity imparted by the N—terminal extension. Cultures were
grown to an A600 of 0.4-0.6 at 30 °C in LB containing 100 ug/ml ampicillin, induced
with 0.3 mM IPTG, and incubated for three hours. Harvested cells were suspended 20

mM Tris buffer (pH 7.5) containing 1 mM EDTA, 200 mM NaCl, and 1 mM

88

dithiothreitol, and lysed by two passages through a French pressure cell at 120 MPa. The
cell lysates were spun at 100,000 x g to yield the clarified cell extracts. The 89.7 kD
fusion protein was readily purified by passing cell extracts over an amylose column
according to manufacturer’s instructions followed by elution in the above buffer amended

with 10 mM maltose.

Characterization of MBP-dioxygenase fusion protein.

The MBP-dioxygenase fusion protein was assayed for activity towards 2,4-D
using the 4—aminoantipyrene assay described previously (5). Activity towards sulfonate-
containing compounds was determined by quantifying the amount of sulfite released
using Ellman’s reagent (5,5’-dithiobis(2-nitrobenzoic acid)) (4). Standard assay
conditions were defined as 10 mM dimethylglutarate (pH 7.0), 1 mM a—KG, and 50 uM

Fe(H)-ascorbic acid salt (Sigma) for 5 min at 30°C in the presence of varied substrate

concentrations. Consumption of OL-[ 14C—1]-KG was determined using a previously
described method (5). One unit is defined as one umol of product formed per min using

the standard assay conditions.

Studies with the YLL05 7c deletion mutant

The S. cerevisiae YLL057c deletion mutant, BY4742-11545 and the
corresponding wild type strain, BY4742, were obtained from Research Genetics
(Huntsville, AL). Overnight cultures for inoculation of sulfur limited media were grown
in yeast peptone dextrose (YPD) broth. Mutant cultures were amended with 50 pig/ml

geneticin sulfate antibiotic. To assess the growth of S. cerevisiae BY4742—11545 and

89

 

 

BY4742 with alternative sulfur sources, strains were grown aerobically in defined
medium (3) with isethionate (250 uM), taurine (250 uM), sulfate (250 uM) or with no
sulfur source added. Since geneticin, is sold as a sulfate salt, the antibiotic was omitted

from experiments using low sulfur medium.

Results and Discussion

The product of YLL057c was found to be an F e(H)— and OL-KG-dependent
dioxygenase capable of utilizing a variety of sulfonates (Figure 12, Scheme 3) but was
inactive on 2,4-D. Maximum rates (Vmax), affinities (Km), and catalytic efficiencies
(kcat/Km) for various substrates are compared in Table 7. Whereas taurine was a
relatively poor substrate for the yeast sulfonate/oc—KG dioxygenase (with a moderate
reaction rate and high Km), isethionate (2—hydroxy-ethanesulfonate) and taurocholate
were tumed over more rapidly and bound with much higher affinity by the enzyme.
According to their catalytic efficiencies, the best substrates were the synthetic compounds
MOPS and N—phenyltaurine. Several of the better substrates for this enzyme had large
adducts on the amino group, indicating a relaxed substrate specificity for this portion of
the molecule. Of the substrates listed in Table 7, only taurine, MOPS (4) and taurocholate

(1) were also substrates of E. coli TauD. Notably, TauD activity toward taurine exhibits a

kcat of 133 min-1, which is the same order of magnitude as found here for the yeast

enzyme. While taurine utilization followed simple Michaelis-Menten kinetics

90

 

Q‘NNEI—OH N-phenyltaurine

I/\N/\/\E——-OH MOPS

 

 

 

”0 ~ 0“ lsethionic acid
I)
o
H l?
NH/VETOH Taurocholic acid
HO OH O

H2N\/\L| T _
g———_OH aunne

Figure 12. Structures of substrates for S. cerevisiae sulfonate/oc-KG dioxygenase.

COO-

R\/fi\ coo-
R 0 2+ + HSO - + +
\/\ + + 02 Fe H 3 C02

——>
C00-
C00-

Scheme 3. Reaction mechanism of S. cerevisiae sulfonate/a-KG dioxygenase.

91

 

(Figure 13A), the kinetic properties for several other substrates were complicated by
apparent inhibition of the enzyme at elevated substrate concentrations (Figure 13B and
C). In those cases, the data were fit to the standard equation for substrate inhibition

(Equation 1).

Equation 1 v = VmaXISl/([SJ+Km +[512/Kn

Inhibition constants (Ki) are listed in Table 7. TauD exhibited similar substrate inhibition
profiles for certain substrates other than taurine (1). High concentrations of substrate are
also inhibitory to y-butyrobetaine hydroxylase (K, is approximately 1 mM) (19), an
enzyme that catalyzes the conversion of y-butyrobetaine to camitine and to prolyl
hydroxylase (10). In y-butyrobetaine hydroxylase researchers demonstrated that the
uncoupled decarboxylation of oc-KG at high substrate concentrations did not explain the
apparent inhibition. The peptide substrate of prolyl hydroxylase is a competitive inhibitor
with respect to iron and oc-KG for that enzyme (10). Sulfonate/OL-KG dioxygenase
activity was not detected towards the following compounds: cysteate, aniline-2-sulfonic
acid, homotaurine, picrylsulfonic acid, sulfosalicylic acid, 3-[(3—cholamidopropyl)
dimethylammonio] propanesulfonic acid (CHAPS), 4-(2-hydroxyethyl)piperazine-1-
propanesulfonic acid (EPPS), piperazine-N,N'—bis(2-ethanesulfonic acid) (PIPES), and 3-

(cyclohexylamino)-1-propanesulfonic acid (CAPS).

92

 

Table 7. Sulfonate specificity and kinetic parameters for sulfonate/a-KG

dioxygenase.

 

Substratea

3-N-Morpholino-
propanesulfonic acid
(MOPS) [0-20 mM]

N-Phenyltaurine [0-3 mM]

2-Hydroxyethanesulfonate
(Isethionate) [0-20 mM]

3-Morpholino-2-hydroxy-
propanesulfonic acid
(MOPSO) [0-20 mM]

Taurocholate [0-10 mM]

2-(4-Morpholino)-
ethanesulfonic acid
(MES) [0-20 mM]

2-(Cyclohexylamino)
ethanesulfonic acid
(CITES) [0-20 mM]

N—(Tris[hydroxymethy1]
methyl)-2-amino-
ethanesulfonic acid (TES)
[0-15 mM]
N-[2-Hydroxyethyl]
piperazine-N'-[2-hydroxy-
propanesulfonic acid]
(HEPPSO) [0-20 mM]

N-(Tris(hydroxymethyl)—
methyl)-3-aminopropane-
sulfonic acid

(TAPS) [0-20 mM]

4-(2-Hydroxyethyl)-1-
piperazineethanesulfonic
acid (HEPES) [0-20 mM]

2-Aminoethanesulfonic
acid (Taurine) [0-20 mM]

Vmax

(umol/min/mg)

1.80 i 0.06

0.81 i 0.03

1.26 i- 0.05

1.51 i 0.03

0.99 i 0.05

1.48 i 0.05

0.56 i- 0.02

0.75 i 0.01

1.01 i 0.08

1.61i0.06

0.82 i 0.02

0.46 i 0.04

Km
(mM)

0.24 i- 0.02

0.12:0.0’1

0.27 i 0.05

0.33 i- 0.02

0.42 i 0.05

1.94i0.14

2.57 i 0.34

3.60:0.12

5.04 i 1.07

10.38 i- .80

16.62 i 0.80

10.37 i 1.82

kcat/Km
(min'l/mM)

670.9
606.4

418.4

411.2

210.7

68.4

19.6

18.7

18.0

13.9

4.4

4.0

Ki
(111M)

34.3 i 5.8b

1.5:r0.1b

20.3 i 2.5b

28.4 i 2.4b

8.2:r1.1b

45.2 i 6.6b

 

a The concentrations of substrate examined are shown in brackets
b Data for these substrates were fit to Equation 1
c No inhibition was detected. Kinetic parameters were fit to the Michaelis-Menten

equation.

93

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Additional properties of S. cerevisiae sulfonate dioxygenase
Several additional properties of the MBP-dioxygenase fusion protein were

characterized. The Km for a-KG was determined to be 30 i 16 uM. The approximate KD

for F e(H) was found to be 44 i 5 uM which is ten-fold higher than the value observed for

other OL-KG-dependent dioxygenases (see Chapter 1). The increase in apparent KD for
iron may reflect non-specific metal binding by lVIBP. Temperature stability experiments,
conducted by incubating the protein at varying temperatures for 1 hr prior to measuring
activity at 30 °C, showed that the enzyme lost 77% of its activity after incubation at 37°C
and was completely inactivated at higher temperatures. The fusion protein was shown to
be cleaved by Factor Xa (New England Biolabs) to yield active 47 kDa enzyme. There
was no significant difference in kcat towards MOPS for the fusion protein and the free

enzyme.

Phenotype of S. cerevisiae YLL05 7c deletion mutant

Studies with a S. cerevisiae YLL057c deletion mutant, BY4742-11545 (Research
Genetics, Huntsville, AL), demonstrated the importance of sulfonate/a-KG dioxygenase
for growth on alternative sulfur sources. Comparison of the growth profiles for these two
strains clearly showed that deletion of the sulfonate/oc-KG dioxygenase gene decreased
the extent to which S. cerevisiae could use taurine and isethionate as the sole source of
sulfur (Figure 14). The rate at which BY4742 grew in medium with isethionate was also
affected (data not shown). The growth rate and final yield was identical for the two

strains in medium containing sulfate.

95

 

0.6

 

0.5

0.4 3

0.3 ~

0.2 .

Absorbance (600 nm)

0.1 ~

 

 

 

 

 

 

_I:u_l:_

Sulfate lsethionate Taurine No Sulfur added

 

 

 

 

Sulfur Source

Figure 14. Role of YLL057c in the utilization of different sulfur sources. Growth of
S. cerevisiae BY4742 (solid bars) and the YLL057c deletion strain BY4742-11545 (open
bars) in medium containing either sulfate, isethionate, taurine, or no added sulfur source
was measured as absorbance at 600 nm of stationary phase cultures. Error bars represent
the standard deviation between replicate cultures.

96

Role of sulfonate/a-K G dioxygenase in S. cerevisiae

The yeast sulfonate/a-KG dioxygenase may allow cells to exploit one or more of
the many naturally occurring sulfonates. Some studies suggest that sulfonate sulfur is the
major form of organic sulfur in soils (2). Specific examples of environmental sources of
sulfonates include the sulfonolipids of gliding bacteria, the capsules of some Gram
positive bacteria (1 l), and the taurine produced in all vertebrates and invertebrates (6). In
addition, common oxidation products of cysteine and other S-containing amino acids
include sulfoacetaldehyde (9, 14), sulfopyruvate, sulfolactate, cysteate (20), and taurine
(20). In mice, the gut microflora have been shown to convert taurine and taurocholate to
isethionate (20). Perhaps significantly, a taurocholate transporter (YBTl) (12) resides on
the same segment of chromosome XH as YLL057c. The bile acid transporter has been
localized to the vacuolar membrane and efficiently imports taurocholate into the vacuole.
Sulfonate/oc-KG dioxygenase may play an additional role in the degradation of taurine-
conjugated compounds in viva.

Candida albicans, another fungus, contains a hypothetical protein sequence with
more than 50% amino acid identity to the sulfonate/or-KG dioxygenase suggesting that
other fungi have similar pathways to take advantage of these alternative sources of sulfur
(15). The fact that S. cerevisiae BY4742-11545 could grow to some degree on isethionate
suggests that an additional pathway exists for sulfonate utilization. Examples of enzymes
capable of the liberation of sulfite from sulfonates include a flavin mononucleotide-
dependent monooxygenase from Pseudomonas aeruginosa (7) and a sulfo-lyase from an
Acinetobacter isolate (8). Redundancy of sulfonate catabolic pathways accentuates the

importance of these compounds as sources of cell sulfur.

97

10.

11.

12.

Literature cited
Auchtung, T. A. 1999. Unpublished observations.

Autry, A. R., and J. W. Fitzgerald. 1990. Sulfonate S: A major form of forest
soil organic sulfur. Biol. Fertil. Soils 10:50-56.

Cherest, H., and Y. Surdin-Kerjan. 1992. Genetic analysis of a new mutation
conferring cysteine auxotrophy in Saccharomyces cerevisiae: updating of the
sulfur metabolism pathway. Genetics 130:51-58.

Eichhorn, E., J. R. van der Ploeg, M. A. Kertesz, and T. Leisinger. 1997.

Characterization of OL—ketoglutarate-dependent taurine dioxygenase from
Escherichia coli. J. Biol. Chem. 272:23031-23036.

Fukumori, F., and R. P. Hausinger. 1993. Purification and characterization of

2,4-dichlorophenoxyacetate/oc-ketog1utarate dioxygenase. J. Biol. Chem.
268:24311-24317.

Huxtable, R. J. 1992. Physiological actions of taurine. Physiol. Rev. 72:101-143.

Kertesz, M., K. Sehmidt—Larbig, and T. Wiiest. 1999. A novel reduced flavin
mononucleotide-dependent methanesulfonate sulfonatase encoded by the sulfur
regulated msu operon of Pseudomonas aeruginosa. J. Bacteriol. 181:1464-1473.

King, J. E., R. Jaouhari, and J. P. Quinn. 1997. The role of sulfoacetalaldehyde
sulfo-lyase in the mineralization of isethionate by an environmenal Acinetobacter
isolate. Microbiology 143:2339-2343.

Kondo, H., H. Anada, K. Ohsawa, and M. Ishimoto. 1971. Formation of
sulfoacetaldehyde from taurine in bacterial extracts. J. Biochem. 69:621-623.

Myllyla, R., L. Tuderman, and K. I. Kivirikko. 1977. Mechanism of the prolyl
hydroxylase reaction: 2. Kinetic analysis of the reaction sequence. Eur. J.
Biochem. 80:349-357.

Ohtomo, T., K. Yoshida, and C. L. San Clemente. 1981. Effect of bile acid
derivatives on taurine biosynthesis and extracellular slime production in
encapsulated Staphylococcus aureus S-7. Infec. Irnmun. 31:798-807.

Ortiz, D. F., M. V. St. Pierre, A. Abdulmessih, and I. M. Arias. 1997. A yeast

ATP-binding cassette-type protein mediating ATP-dependent bile acid transport.
J. Biol. Chem. 272:15358-15365.

98

13.

14.

15.

16.

17.

18.

19.

20.

Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: a
laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring
Harbor, NY.

Shimamoto, G., and R. S. Berk. 1979. Catabolism of taurine in Pseudomonas
aeruginosa. Biochim. Biophys. Acta. 569:287-292.

Unfinished Genome Database Website. 11 October 1999.
http://www.ncbi.n1m.nih.gov/BLAST/ unfinishedgenomehtml.

Uria-Nickelsen, M. R., E. R. Leadbetter, and W. Godchaux IH. 1994.
Comparative aspects of utilization of sulfonate and other sulfur sources by
Escherichia coli K12. Arch. Microbiol. 161:434-438.

Uria-Nickelsen, M. R., E. R. Leadbetter, and W. Godchaux III. 1993.
Sulfonate-sulfur assimilation by yeasts resembles that of bacteria. FEMS
Microbiol. Letts 114:73-78.

Uria-Nickelsen, M. R., E. R. Leadbetter, and W. Godchaux III. 1993.
Sulphonate utilization by enteric bacteria. J. Gen. Microbiol. 139:203-208.

Wehbie, R. S., N. S. Punekar, and H. A. Lardy. 1988. Rat liver y-butyrobetaine
hydroxylase catalyzed reaction: influence of potassium, substrates and substrate
analogues on hydroxylation and decarboxylation. Biochemistry 27 :2222-2228.

Weinstein, C. L., and O. W. Griffith. 1988. Cysteinesulfonate and 13-
sulfopyruvate metabolism. J. Biol. Chem. 263:3735-3743.

99

CHAPTER 4

DISTRIBUTION OF tfdA IN THE ENVIRONIVIENT

Many of these results have been published in the article: Hogan, D. A., D. H.
Buckley, C. H. Nakatsu, T. M. Schmidt, and RP. Hausinger (1997). “Distribution of the
tfdA gene in soil bacteria that do not degrade 2,4-dichlorophenoxyacetic acid (2,4-D).”
Microb. Ecol. 34:90-96.

100

Introduction

The tfa’A gene (30) encodes 2,4-D/a-KG dioxygenase (deA) (9), an enzyme that
catalyzes removal of the acetate side chain of 2,4-D yielding 2,4-dichlorophenol. Thus
far, tfdA is uniquely associated with 2,4-D catabolism, and is most often found with the
other tfd genes necessary to complete the degradation of 2,4-D. While tfdA is unlike any
previously characterized genes involved in aromatic compound decomposition, the other
tfd genes are similar to components of a number of different catabolic pathways. For
instance, genes similar in sequence to lde encode hydroxylases that act on various
substituted or unsubstituted phenols in a variety of microorganisms (15, 39). Likewise,
the tdeDEF operon is homologous to other catechol degradation genes involved in many
different aromatic degradative pathways (3 9). Because the downstream tfd genes are
closely related to common bacterial genes, it is likely that homologs to tfdA can also be
identified.

Several pieces of evidence suggest that the assembly of the tfd genes to form 2,4-
D degradation plasmids has occurred numerous times. First, the 2,4-D degradative genes
are found in different arrangements on a variety of different plasmid backbones (35, 37).
Second, plasmids within the same incompatibility group can have different alleles of the
genes that encode the first three enzymes of the 2,4-D degradative pathway (10, 36).
Third, on some plasmids, tfdA is physically distant from the zdeDEFB gene cluster (7).
For there to be a “source” of tfd genes in the environment prior to their incorporation in
2,4-D degradative pathways, these genes must either be maintained without selection or

serve an alternative purpose. Consistent with a widespread availability of tfdA homologs,

101

evidence for deA or deA-like activity has been found in diverse phylogenetic groups
including the y-(21) and B-Proteobacteria (2, 6, 10, 32), a member of the Flexibacter-
Cytophaga— Bacteroides phylum (3), and Arthrobacter sp. in the High G+C Subdivision
of the Gram-positive phylum (34).

To assess the natural distribution of tfdA, I used PCR primers internal to the tfdA
gene to screen a collection of soil microorganisms, isolated on a non-selective medium,
for the presence of tfdA or tfdA—like genes. In this chapter, I provide evidence that tfdA-
like sequences are present in phylogenetically diverse soil microbes that do not degrade
2,4-D. These data suggest that tfdA is maintained in natural bacterial populations,

possibly for a purpose other than 2,4-D degradation.

Materials and Methods
Bacterial strains

Bacterial strains were isolated from soils obtained from a Long Term Ecological
Research (LTER) plot (24) at the Kellogg Biological Research Station in Hickory
Comers, IVH, USA which had been managed as a single cropping system to produce grain.
Soil extracts were serially diluted in 50 mM phosphate buffer (pH 7), then plated on R2A
medium (Difco, Detroit, MI) and incubated at 28°C for 10 days. Plates with 30 to 150
colonies were divided into quadrants. All colonies from one quadrant were purified on
R2A agar and incubated at 28°C for three to four days. Additional colonies were chosen
on the basis of colony morphology to include several groups of similar isolates as well as

a representation of distinctive colony morphologies. The origins of Alcaligenes eutrophus

102

JMP134 (pJP4) (6), JMP228 (30), and Burkhola'eria sp. RASC (32) have been described

previously.

I6S rDNA partial sequence analyses

Total DNA was obtained from these isolates by detergent lysis (1) and 168 rDNA
was amplified by PCR using the eubacterial primers complementary to Escherichia coli
nucleotide positions 8-27 and 1510-1492 (8). Amplified rDNA was purified with Ultra-
free MC (30,000 MWCO) centrifugal filter units (Millipore, Bedford, MA), then
sequenced with primers complementary to E. coli positions 536-519 or 8-27 (27). The
rDNA sequence was determined by automated fluorescent sequencing performed by the
MSU-DOE-PRL Plant Biochemistry Facility using an ABI Catalyst 800 for Taq cycle
sequencing and the ABI 373A Sequencer for the analysis of products. Partial 16S rDNA
sequences were aligned manually against a collection of 168 rRNA sequences (18) on the
basis of conserved regions of sequence and secondary structure of 16S rRNA. Regions of
ambiguous alignment were omitted from the subsequent analysis. Phylogenetic trees
were generated using the Neighbor Joining method and the topology was confirmed by

Parsimony and Maximum Likelihood analyses (3 3).

PCR Amplification of the tfdA gene
Total DNA samples were isolated from each of the LTER strains (1). Primers
were derived from conserved regions of sequence in the tfdA genes of pJ P4 and

Burkholderia strain RASC (79% identical to tfdA—pJP4). The sequence of the forward

103

primer, TVU, was 5’-ACG GAG TTC TG(C/T) GA(C/T) ATG-3’; the sequence of the
reverse primer, TVL, was 5’-AAC GCA GCG (G/A)TT (G/A)TC CCA-3’(3 8). The PCR
mix contained 5 ul 10X PCR buffer (Gibco, Gaithersberg, MD), 1.5 mM MgC12, 1 uM
primer TVU, 1 uM primer TVL, 1 mM dNTPs (Boehringer Mannheim, Indianapolis, W),
1.5 units of Taq polymerase (Gibco), 10-100 ng template DNA, and distilled water to
bring the final volume to 50 al. The PCR amplification protocol consisted of thirty five
cycles with a one min denaturation at 94°C, one min amiealing at 55°C and one min
extension at 72°C. The predicted product size was 360 bp. Amplified products were
separated on a 1% agarose gel and compared to PCR products generated from JMP134
and RASC genomic DNA as template. Generation of PCR products was confirmed by at
least one additional PCR amplification reaction. Products were purified with Promega
PCR mini—prep columns (Promega, Madison, WI). Selected PCR products were
sequenced with either primer TVL or TVU by the method described for 168 rDNA

above.

Dot Blot of PCR products

PCR products (~50 ng) were applied to a Hybond N nylon membrane (Amersharn,
Arlington Heights, H.) in 1 ul aliquots and UV cross-linked to the membrane with a
Stratalinker (Stratagene, La J olla, CA). To create probes, PCR products obtained using
DNA from JMP134 and RASC were labeled with digoxigenin using the DIG High Prime
labeling mix (Boehringer Mannheim) according to the manufacturer’s instructions.
Hybridization, detection, and probe removal were performed according to protocols in the

Genius Detection System manual (Boehringer Mannheim). To assure high stringency, the

104

hybridization solution contained 50% forrnamide and hybridizations and stringency
washes were at 68°C. Hybridization signals were detected by the colorimetric detection

system (Boehringer Mannheim).

Analysis 0f2,4-D degradation and deA activity

Isolates were streaked on R2A agar and incubated at 30°C for three days to allow
for growth of all cultures. Isolated colonies were transferred to tubes containing 3 ml of
A+N medium (41) amended with 0.1% yeast extract and 1 mM 2,4-D. Cultures were
incubated for two weeks at 30°C on a shaker. After incubation, cells were removed by
centrifugation and the supernatant was filtered (0.45 urn) into an autosampler vial.
Duplicate samples of each culture were either analyzed immediately or frozen for later
analysis. 2,4-D and 2,4-dichlorophenol concentrations were determined by FH’LC with
70/30 methanol-0.1% phosphoric acid as eluent, with a Lichosorb RP-18 column (EM
Separations, Gibbstown, NJ) and monitored with a UV detector at 230 nm. For induction
experiments, cells were grown on MMO medium (29) with 5 mM succinate, harvested
and washed in phosphate-buffered saline, then resuspended in medium with
phenoxyacetic acid, 3-chlorobenzoate, 3-chlorocatechol, catechol, succinate (all 1 mM
final concentration), or with no carbon source. deA activity was measured by detection

of 2,4-dichlorophenol with 4-aminoantipyrene (9).

Incorporation of radiolabel from 14C-2,4-D

Single colonies from cultures grown on R2A plates were transferred to sterile
nylon membranes overlaid on R2A agar. After overnight incubation at 30°C, all cultures

had visible growth. Each filter was replicated (25), then both master and replica filters

105

were transferred to minimal agar containing either l4C-ring-labeled or chain—labeled 2,4-
D and incubated for three days at room temperature. The filters were exposed to X-ray

film (Kodak, Rochester, NY) for 18 hours to detect incorporation of radiolabeled carbon.

Results

Composition of the LTER collection

The analysis of 16S rDNA partial sequence from strains in the LTER collection
showed that these isolates represent diverse phylogenetic divisions (Figure 15,Table 8).
The taxonomic divisions used to describe the isolates are defined in the Ribosomal
Database Project (18). Among the seventy-six isolates analyzed, twenty seven were found
to cluster within the Bacillus-Lactobacillus-Streptococcus subdivision and thirteen strains
fall within the High G+C Subdivision of the Gram-positive phylum. The remaining
isolates included five a-Proteobacteria, two B-Proteobacteria, eleven y-Proteobacteria,
and eighteen members of the Cytophaga group. In a polyphasic taxonomy study, more
than 95% of these isolates were found to be distinct taxa based on BIOLOG analyses
(Biolog, Hayward, CA), fatty acid methyl ester (FAME) profiles (13), and genomic
fingerprints from rep-PCR analyses (40). The polyphasic taxonomy study of the LTER
isolates, including refined phylogenetic analyses based on complete 16S rDNA sequence
is underway and will be presented elsewhere. Because the LTER collection includes both
closely related isolates as well as organisms from several different phyla, this collection
provides a good opportunity to test for the dispersion of tfdA between distantly related

bacteria as well as for the presence of tfdA in similar strains within a bacterial species.

106

Identification of [@211 in LT ER collection

PCR amplification with internal tfdA primers generated products using genomic
DNA from twenty eight of the seventy six isolates. The products were similar in size to
the tfdA product amplified from M134 and RASC (360 bp), and hybridized at high
stringency to labeled products from PCR amplification of tfdA from JMP134 (Figure 16)
and Burkholderia RASC. Amplification product from each isolate was assigned to either
the tfdApm or tfdARAsc hybrization group. When primers TVU and TVL were applied
directly to the membrane, hybridization to the probe was not detected.

Amplified PCR products from four isolates were sequenced to assess the
similarity of the tfdA-like gene in organisms from different phylogenetic divisions to each
other and to previously characterized tfdA genes. Three of the isolates, LTER 1 and LTER
11 from the Bacillus-Lactobacillus-Streptococcus subdivision as well as LTER 8 from the
Rhizobium-Agrobacterium group were identical to the tfdA partial sequence (Genbank
U22499) from Halomonas sp. I-l 8, a moderate halophile (21) and to the tfdA sequence
identified on plasmid pEMT8 (36) (Dr. Terrence Marsh, unpublished data). Although
pEMT8 does not confer the ability to degrade 2,4-D to its host, it can complement R.
eutropha(pBH501aE) (36), a tfdA" derivative of pJ P4, to restore the 2,4-D degradation
phenotype (37) suggesting that it does encode a functional deA. The sequence of the
PCR product from LTER 40, a member the B-Proteobacteria division, was identical to the

sequence of tfdA found on pJP4.

107

Figure 15. 16S rDNA phylogeny of the LTER isolates

Phylogeny is based on partial sequences corresponding to E. coli rDNA sequence

between positions 27 and 519. Names of phylogenetic divisions are derived from the
RDP 5.0 release (18). The scale bar indicates the number of nucleotide changes per base
position analyzed. Isolates that produce a PCR amplification product with primers
internal to the tfdA gene are enclosed in boxes. The abbreviation Rub. indicates
Rubrivivax, th. indicate Rhizobium, and Ag. signifies Agrobacterium. LTER isolates in
dark boxes yielded a PCR product belonging to tfdApm hybridization group; grey boxes
indicate tfdA-like genes more similar to ddARAsc-

108

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2,4-D degradation by LTER isolates

HPLC analyses showed that 2,4-D was not degraded to a significant extent by any
of the LTER isolates after a two week incubation. In contrast, JMP134 (pJP4) or
Burkholderia sp. RASC completely consumed the 2,4-D during this time period. Uptake
experiments provided additional evidence that none of the LTER isolates utilized 2,4-D
for cellular carbon. Neither ring- nor chain-labeled 14C-2,4-D was incorporated into the
LTER cultures after a three day incubation period while JMP134(pJP4) produced a strong

positive signal after incubation on medium with either radiolabeled substrate.

Experiments to determine if the [fl-like gene encodes a functional deA

To determine if the tfdA-like genes encode functional enzymes, selected isolates
from different phylogenetic groups were tested for deA activity and the presence of a
deA—like protein in cell extracts. Ten isolates from different phylogenetic groups were
selected for fiirther study including: LTER 28 (B-Proteobacterium); LTER 38 and LTER
82 (or-Proteobacteria); LTER 40 (y-Proteobacterium); LTER 20 (Cytophaga Group);
LTER 1, LTER 11 and LTER 24 (Bacillus group); and LTER 3 and LTER 5
(Arthrobacter group). While JMP134 and RASC had high levels of deA activity after
induction with 2,4-D, none of the cell lysates from the above LTER strains exhibited any
activity after induction with 2,4-D, phenoxyacetic acid, 3-chlorobenzoate (3-CBA),
catechol, 3-chlorocatechol, succinate or with no added carbon source. JMP134 was the
only strain that degraded 3-CBA, as determined by HPLC analysis. Western blot analysis

with anti-deA(pJP4) antibodies did not indicate the presence of a cross-reacting deA in

110

1 2 3 4 8 9 10 11 I2 14
4Q o :T '3 # Q G . C
18 20 22 24 25 27 28 31 38 40
O . c O
48 65 67 68 69 75 77 82 90 NC
«... . {3 .
P R 69~

Figure 16. Hybridization of amplified products from LTER isolates to a
digoxigenin-labeled fragment internal to the tfdA gene.

The numbers correspond to the LTER isolates that yielded the template DNA for the PCR
reaction; 69* is an amplified product that is 40 bp longer than the predicted product size;
as a negative control, NC, PCR primers were applied directly to the membrane; P is the
amplified product from JMP134 (pJ P4); R, amplified product from Burkholderia strain
RASC. Results from hybrization to RASC probe not shown.

111

Table 8. Distribution of isolates with tfdA among the different phylogenetic groups
within the LTER collection.

 

 

 

Phylogenetic subdivison Total no. No. of isolates % of isolates with
of isolates with tfd/la {fdA
pfll RASC

a—Proteobacteria 5 2 2 60
B-Proteobacteria 2 O 2 1 OO
y-Proteobacteria 1 1 1 2 27

Cytophaga 18 0 1 6

High G+C Gram-positives 13 1 5 46
Bacillus-Lactobacillus- 27 5 7 40

Streptococcus

Total 76 9 19 37

 

a Amplified PCR products were categorized into hybridization groups based on the
strength of the hybridization signal to the pJP4 and RASC probe.

112

lysates from any of the LTER isolates. In a separate experiment, cell extracts of
Halomonas sp. LI 8 grown on 2,4-D possessed a band of approximately 32 kDa (similar
in size to pJP4 and the predicted MW of pEMT8 deA) that cross reacted with the anti-
deA (pJP4) antibody, however, RASC, grown with 2,4-D as the sole source of carbon,
did not.

Mating experiments similar to those published by Top et al. (3 6) were also
performed to determine if any of the LTER isolates could transfer a functional tfdA to a
recipient strain. No 2,4-D+ transconjugants were generated after incubation of the LTER
isolates with R. eutropha JMP228 carrying pBHSOlaE (a tfdA‘ pJP4-derived plasmid) or

plasmid-free R. eutropha JMP228 (N alR) (36).

Discussion

A significant percentage (37%) of organisms in this collection of soil isolates
contains tfdA or tfdA-like genes. Two aspects of these data are particularly interesting.
First, the tfdA-like genes are found in very phylogenetically diverse taxa (Figure 15, Table
8) including both Gram positive and Gram negative bacteria. Second, isolates within the
same phylogenetic division contain tfdA-like genes from different hybridization groups
(Table 8). Previous research provides evidence for tfdA in only a small number of Gram-
positive 2,4-D-degraders, such as Arthrobacter sp. (34), compared to a vast collection of
tfdA-containing Gram-negative bacteria (2, 6, 21 , 32, 35). This is the first report of a tfdA-
like gene in Bacillus or C ytophaga.

Despite the fact that 2,4-D is readily degraded by many different microbial

communities (26), the isolation of axenic cultures that degrade 2,4-D has proven to be

113

more difficult (11, 17). These data suggest that, in many instances, 2,4-D is not degraded
by individual organisms, but by the concerted activities of several different bacteria. The
presence of tfdA in isolates that do not degrade 2,4-D may indicate the source of the 2,4-
D-degradation potential of microbial communities. Perhaps prolonged exposure to 2,4-D
is necessary to select for organisms or plasmids that contain both tfdA and the tdeCDEF
genes. If tfdA is present in bacteria lacking the additional genes needed for 2,4—D
mineralization, deA activity may be difficult to detect in pure culture due to the
accumulation of toxic products, such as 2,4-dichlorophenol, or lack of the proper inducer
molecule (19).

The high identity of the partial tfdA—like sequences, amplified from the DNA of
phylogenetically diverse bacteria, to previously characterized tfdAs suggests extensive
horizontal transfer of these genes. The tfdA gene is most often found on self-
transmissible, broad host-range plasmids that confer the ability to degrade 2,4-D (2, 6,
36). Several studies (4-6) have shown that plasmid pJP4, which encodes all of the
enzymes necessary for 2,4-D degradation, can be maintained in a number of different
bacteria. It is worth noting, however, that not all organisms containing pJP4 express the
2,4-D-degrading phenotype (6).

Genetic characterization of plasmid-encoded 2,4-D degradation pathways indicate
that tfdA most often resides on transposable elements. Plasmids that confer the 2,4-D+
phenotype are often unstable. Multiple researchers have reported that growth on non-
selective media induces the loss of a 20- to 40-kb fragment from a plasmid concurrent
with the inability to degrade 2,4-D (2, 7, 20, 42). A plasmid isolated from Michigan soils,

pEMT8, (3 7) contains tfdA but does not have the other genes required for 2,4-D

114

degradation. The pEMT8 tfdA, which is identical to the partial sequences amplified from
three of the LTER isolates, is adjacent to a transposase of the ISI 1 1 1/ISl328/IS15 33
family of elements (Dr. Terrence Marsh, unpublished data). Similar transposable
elements have been found in diverse organisms including strains of Streptomyces (16),
Mycobacterium (l6), Pseudomonas (Genbank # AF052750) and Rhizobium (Genbank#
P55615). With this, it is reasonable to hypothesize that the tfdA—like genes in the LTER
isolates are associated with a similar transposable element. Consistent with this
hypothesis is the observation that the tfdA-like gene can be lost, as evidenced by the
inability to amplify the 360 bp product (data not shown), upon multiple transfers of the
isolates on rich medium.

The wide spread presence of tfdA, even on a transposon, in organisms that do not
degrade 2,4-D may indicate that these genes serve an important metabolic purpose apart
from herbicide degradation. There are several examples of genes that reside primarily on
mobile elements, including those encoding virulence factors (14), plant-microbe
symbiosis functions (31), and antibiotic resistance (23). The strategy of maintaining
important genes in only a fraction of the population by use of mobile genetic elements is
poorly understood, but may prove to be important to microbial ecology. Naturally-
occurring compounds with structural similarities to 2,4-D that are potential substrates of
native deA include the numerous aryl-ether compounds that are released during fungal
degradation of lignin (22) or the different halogenated aromatic metabolites produced by
fungi and bacteria (12, 28). Alternatively, it is possible that the natural substrate for deA

shows little resemblance to 2,4-D and the ability to degrade this compound is fortuitous.

115

Acknowledgements.

I am grateful to Helen Corlew-Newman and Frank Lowes for providing the LTER
strains and the past and present members of the Research on Microbial Evolution
(ROME) Laboratory for helpful suggestions and discussions. Particular thanks go to
Terry Marsh for providing the unpublished pEMT8 tfdA and transposase sequences and to

Olga Maltseva for providing DNA and cell material from Halomonas strain I—18.

116

10.

Literature cited

Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A.
Smith, and K. Struhl. 1987. Current protocols in molecular biology. John Wiley
and Sons, New York.

Bhat, M. A., M. Tsuda, K. Horiike, M. Nozaki, C. S. Vaidyanathan, and T.
Nakazawa. 1994. Identification and characterization of a new plasmid carrying
genes for degradation of 2,4-dichlorophenoxyacetate from Pseudomonas cepacia
CSV90. Appl. Environ. Microbiol. 60:307-312.

Chaudry, G. R., and G. H. Huang. 1988. Isolation and characterization of a new
plasmid from a F lavobacterium sp. which carries the genes for the degradation of
2,4-dichlorophenoxyacetate. J. Bacteriol. 170:3897-3902.

Daane, L. L., J. A. E. Molina, E. C. Berry, and M. J. Sadowsky. 1996.
Influence of earthworm activity on gene transfer from Pseudomonas fluorescens
to indigenous soil bacteria. Appl. Environ. Microbiol. 62:515-521.

DiGiovanni, G. D., J. W. N eilson, I. L. Pepper, and N. A. Sinclair. 1996. Gene
transfer of Alcaligenes eutrophus JMP134 plasmid pJP4 to indigenous soil
recipients. Appl. Environ. Microbiol. 62:2521-2526.

Don, R. H., and J. M. Pemberton. 1981. Properties of six pesticide degradation
plasmids isolated from Alcaligenes paradoxus and Alcaligenes eutrophus. J.
Bacteriol. 145:681-686.

Don, R. H., A. J. Weightman, H. J. Knackmuss, and K. N. Timmis. 1985.
Transposon mutagenesis and cloning analysis of the pathways for degradation of
2,4-dichlorophenoxyacetic acid and 3-chorobenzoate in Alcaligenes eutrophus
JMP134 (pJP4). J. Bacteriol. 161:85-90.

Eden, P. A., T. M. Schmidt, R. P. Blakemore, and N. R. Pace. 1991.
Phylogenetic analysis of Aquaspirillum magnetotacticum using polymerase chain
reaction-amplified 16S rRNA-specific DNA. Int. J. Syst. Bacteriol. 42:166-170.

Fukumori, F., and R. P. Hausinger. 1993. Purification and characterization of

2,4-dichlorophenoxyacetate/OL-ketoglutarate dioxygenase. J. Biol. Chem.
268:24311-24317.

Fulthorpe, R. R., C. McGowan, O. V. Maltseva, W. E. Holben, and J. M.

Tiedje. 1995. 2,4-Dichlorophenoxyacetic acid-degrading bacteria contain mosaics
of catabolic genes. Appl. Environ. Microbiol. 61:3274-3281.

117

.11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

Fulthorpe, R. R., A. N. Rhodes, and J. M. Tiedje. 1996. Pristine soils
mineralize 3-chlorobenzoate and 2,4-dichlorophenoxyacetate via different
microbial populations. Appl. Environ. Microbiol. 62:1 159-1 166.

Gribble, G. W. 1992. Naturally occurring organohalogen compounds-a survey. J.
Nat. Prod. 55:1353-1395.

Haack, S. K., H. Garchow, D. A. Odelson, L. J. Forney, and M. J. Klug. 1994.
Accuracy, reproducibility, and interpretation of fatty acid methyl ester profiles of
model bacterial communities. Appl. Environ. Microbiol. 60:2483-2493.

Hacker, J., G. Blum-Oehler, I. Muhldorfer, and H. Tschape. 1997.
Pathogenicity islands of virulent bacteria: structure, function and impact on
microbial evolution. M01. Microbiol. 23: 1089-1097.

Hiiggblom, M. M. 1992. Microbial breakdown of halogenated aromatic
pesticides and related compounds. FEMS Microbiol. Rev. 103:29-72.

Hoover, T. A., M. H. Vodkin, and J. C. Williams. 1992. A Coxiella burnetti
repeated DNA element resembling a bacterial insertion sequence. J. Bacteriol.
174:5540-5548.

Ka, J. 0., W. E. Holben, and J. M. Tiedje. 1994. Use of gene probes to aid in
recovery and identification of functionally dominant 2,4-dichlorophenoxyacetic
acid-degrading populations in soil. Appl. Environ. Microbiol. 60:1116-1120.

Larsen, N., G. J. Olsen, B. L. Maidak, M. J. McCaughey, R. Overbeek, T. J.
Macke, T. L. Marsh, and C. R. Woese. 1993. The ribosomal database project.
Nucl. Acids Res. 21:3021-3023.

Leveau, J. H., F. Konig, H. F uchslin, C. Werlen, and J. R. Van Der Meer.
1999. Dynamics of multigene expression during catabolic adaptation of Ralstonia
eutropha JMP134 (pJP4) to the herbicide 2, 4- dichlorophenoxyacetate. Mol.
Microbiol. 33:396—406.

Mae, A. A., R. O. Martis, N. R. Ausmees, V. M. K6iv, and A. L. Heinaru.
1993. Characterization of a new 2,4-dichlorophenoxyacetic acid degrading
plasmid pEST4011: physical map and localization of catabolic genes. J. Gen.
Microbiol. 139:3165-3170.

Maltseva, O. V., C. McGowan, R. Fulthorpe, and P. J. Oriel. 1996.

Degradation of 2,4-dichlorophenoxyacetic acid by haloalkaliphilic bacteria.
Microbiol. 142:1115-1122.

118

22.

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

Ribbons, D. W. 1987. Presented at the Technology in the 19903: Utilization of
lignocellulosic wastes, London, UK.

Roberts, M. C. 1996. Tetracycline resistance determinants: mechanisms of
action, regulation of expression, genetic mobility, and distribution. FEMS
Microbiol. Rev. 19:1-24.

Robertson, G. P., and D. W. Freckman. 1995. The spatial distribution of
nematode trophic groups across a cultivated ecosystem. Ecology 76: 1425-1432.

Sambrook, J., E. F. F ritsch, and T. Maniatis. 1989. Molecular cloning: a
laboratory manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring
Harbor, NY.

Sandmann, E. R. I. C., M. A. Loos, and L. P. van Dyk. 1988. Microbial
degradation of 2,4-dichlorophenoxyacetic acid in soil. Rev. Environ. Contamin.
Tox. 101:1-53.

Schmidt, T. M., E. F. DeLong, and N. R. Pace. 1991. Analysis of a marine
picoplankton community by 16S rRNA gene cloning and sequencing. J. Bacteriol.
173:4371-4378.

Siuda, J. F., and J. F. deBernardis. 1973. Naturally occuring halogenated
compounds. Lloydia 36: 107-143.

Stanier, R. Y., N. J. Palleroni, and M. Doudoroff. 1966. The aerobic
pseudomonads: a taxonomic study. J. Gen. Microbiol. 43: 159-271.

Streber, W. R., K. N. Timmis, and M. H. Zenk. 1987. Analysis, cloning, and
high-level expression of 2,4-dichlorophenoxyacetate monooxygenase gene tfdA of
Alcaligenes eutrophus JMP134. J. Bacteriol. 169:2950-2955.

Sullivan, J. T., and C. W. Ronson. 1998. Evolution of rhizobia by acquisition of
a 500-kb symbiosis island that integrates into a phe-tRNA gene [published
erratum appears in Proc Natl Acad Sci U S A 1998 Jul 21 ;95(15):9059]. Proc.
Natl. Acad. Sci. U S A 95:5145-9.

Suwa, Y., A. D. Wright, F. Fukumori, K. A. N ummy, R. P. Hausinger, W. E.
Holben, and L. J. Forney. 1996. Characterization of a chromosomally encoded
2,4-dichlorophenoxyacetic acid/a-ketoglutarate dioxygenase from Burkholderia
sp. strain RASC. Appl. Environ. Microbiol. 62:2464-2469.

Swofford, D. L., and G. J. Olsen. 1990. Phylogeny reconstruction. In D. Hillis
and C. Moritz (ed.), Molecular Systematics. Sinaur Associates, Sunderland, MA.

119

34.

35.

36.

37.

38.

39.

40.

41.

42.

Tiedje, J. M., and M. Alexander. 1969. Enzymatic cleavage of the ether bond of
2,4-dichlorophenoxyacetate. J. Agr. Food Chem. 17:1080-1084.

Tonso, N. L., V. G. Matheson, and W. E. Holben. 1995. Polyphasic
characterization of a suite of bacterial isolates capable of degrading 2,4-D.
Microb. Ecol. 3:3-24.

Top, E. M., W. E. Holben, and L. J. Forney. 1995. Characterization of diverse
2,4-dichlorophenoxyacetic acid-degradative plasmids isolated from soil by
complementation. Appl. Environ. Microbiol. 6 1: 1691-1698.

Top, E. M., O. V. Maltseva, and L. J. Forney. 1996. Capture of a catabolic
plasmid that encodes only 2,4-D/0L-ketoglutarate dioxygenase (deA) by genetic
complementation. Appl. Environ. Microbiol. 62:2470-2476.

Vallaeys, T ., R. R. Fulthorpe, A. M. Wright, and G. Soulas. 1996. The
metabolic pathway of 2,4-dichlorophenoxyacetic acid degradation involves
different families of tfdA and tde genes according to PCR-RFLP analysis. FEMS
Microbiol. Lett. 20: 163-172.

van der Meer, J. R., W. M. de Vos, S. Harayama, and A. J. B. Zehnder. 1992.
Molecular mechanisms of genetic adaption to xenobiotic compounds. Microbiol.
Rev. 56:677-694.

Versalovic, J., M. Schneider, F. J. de Bruijn, and J. R. Lupski. 1994. Genomic
fingerprinting of bacteria using repetitive sequence-based polymerase chain
reaction. Methods Mol. Cell Biol. 5:25-40.

Wyndham, R. C. 1986. Evolved aniline catabolism in Acinetobacter
calcoaceticus during continuous culture of river water. Appl. Environ. Microbiol.
51:781-789.

Xia, X.-S., S. Aathithan, K. Oswiecimska, A. R. W. Smith, and I. J. Bruce.
1998. A novel plasmid pIJBl possessing a putative 2,4—dichlor0phenoxyacetate
degradative transposon TN5530 in Burkholderia cepacia strain 2a. Plasmid

39:154-159.

120

 

CHAPTER 5

CONCLUSIONS AND FUTURE RESEARCH

121

In this dissertation, I presented my research on deA as both a model enzyme for
the a-KG—dependent dioxygenase superfamily and as an enzyme with a unique catalytic
activity in nature. The concluding chapter reviews the model for activity in or-KG-
dependent dioxygenases and highlights three major questions that need to be addressed to
understand the catalytic cycle of these enzymes. Although it is hardly necessary to justify
the study of enzymes so critical to different areas of biology, some potential applications
of the research on different a—KG-dependent dioxygenases in medicine, industry and
agriculture are discussed. Lastly, I describe the ongoing efforts in our laboratory to find
the role of deA or a similar enzyme in catabolic pathways other than those for the

decomposition of 2,4-D.

Understanding the mechanism of oc-KG-dependent dioxygenases

Although the a-KG-dependent dioxygenases are not unified by similarities in
primary structure, data from spectroscopic and biochemical experiments suggest that
these enzymes share many common elements. Thus, the three groups of enzymes, defined
by their amino acid sequences (Chapter 1), are likely derived from a common enzyme
ancestor that underwent rearrangements, insertions or deletions during the evolution of
novel catabolic activities. The similarities in tertiary structure between seemingly
unrelated members of the superfamily is revealed by recent X-ray crystallographic
characterizations (Prof. C. Schofield, unpublished data). Information taken from the
examination of multiple a—KG-dependent dioxygenases has been incorporated into a

general mechanistic model for enzymes in this superfamily.

122

The overall reaction catalyzed by oc—KG-dependent dioxygenases can be distilled
down to three major steps. In this discussion, the proposed OL-KG-dependent dioxygenase
mechanism is compared to that for IPNS, a related enzyme that does not use an or-keto
acid as a co—substrate, to illustrate the common elements in the mechanisms of these
disparate enzymes. The three stages of the reactions (A-C) are described below and
depicted in Figure 17.

(A) Substrate binding to the active site alters the coordination environment of the
iron center to enable oxygen binding and formation of the first reactive oxygen species.
The coordination geometry and electronic nature of ligand donors act in concert to
control the reaction between iron and oxygen (19). In enzymes that use oc-KG as a co-
substrate, the oc-keto acid chelates the metal through the C-1 carboxylate and the C-2 keto
group (11, 25, 28, 36). The replacement of neutral solvent ligands with anionic species is
proposed to cause a shift in the iron’s redox potential that may be necessary for
subsequent steps in the reaction. In deA (4, 42), clavaminate synthase (25), and DAOCS
(36), the unique substrate is not thought to interact directly with the metal, but rather
induces a change in the metallocenter geometry from six— to five-coordinate. Together,
the altered geometry and changes in redox potential promote 02 binding to the iron and
generation of an Fe(HI)-superoxide intermediate. Although IPNS does not bind Ot-KG, a
similar shift to five-coordinate geometry is brought about by direct ligation of a substrate
thiol to the iron center (26, 27), thus enabling the binding of oxygen and formation of the
proposed F e(III)-superoxide species.

(B) The first oxidation reaction generates an F e(I V)-oxo intermediate. Iron-

containing model compounds with ligands resembling the 2 His-l-carboxylate facial triad

123

and a bound oc-keto acid moiety have been used to study the initial steps in catalysis (10).
Upon exposure to diatomic oxygen, the synthesized compound undergoes oxidative
decarboxylation concomitant with the hydroxylation of an adjacent aromatic ring.
Comparison of the rate for dioxygenase-like activity for this compound to the decreased
rates with other model compounds suggests that the decarboxylation of a-KG provides a
large driving force for cleavage of the oxygen-oxygen bond (10). This portion of the
reaction catalyzed by oc-KG-dependent enzymes is analogous to the first two-electron
oxidation reaction in IPNS where a new nitrogen-carbon bond is created upon ring
closure and two electrons are transferred from the substrate to oxygen to form a molecule
of water.

(C) The reactive F e(I V)-oxygen species participates in a hydroxylation reaction
or alternative oxidation (desaturation, ring expansion, cyclization) regenerating the
F e(II) center. In hydroxylating enzymes, a specific hydrogen atom is abstracted from the
substrate to form an F e(111)-OH species. The iron-oxygen bond breaks, yielding a
hydroxyl radical which recombines with the substrate to give the hydroxylated product
(32). In IPNS, a series of bonds are cleaved homolytically to generate a ring via a new
carbon-sulfur bond. Again two electrons are transferred from substrate to reduce the
second atom of oxygen to water, thus restoring the iron to its Fe(H) oxidation state.

Additional research is needed in three particular areas to provide experimental
evidence and additional information for several of elements of the above model. First, the

ligands to the metal during different stages of catalysis must be identified and other

124

 

Figure 17. Comparison of the proposed reaction mechanisms for an OL-KG-
dependent hydroxylase and IPN S.

Both resting enzymes possess an Fe(II) that is bound by a facial triad of residues in an H-
X-D-Xn-H motif. (A) In the oc-KG-dependent hydroxylases (left branch), two water
molecules are displaced upon binding of anionic oc-KG to the metal and a third is lost
upon binding of a second substrate (S-H) nearby. The activated metal reacts with 02 to
form F e(III)—superoxide. A similar intermediate is generated in IPNS by stepwise binding
of the substrate and oxygen (right branch). (B) An Fe(IV)-oxo species is generated by a
reaction driven by oc-KG decarboxylation (left) or by oxidative cyclization of the lactam
ring (right). (C) The highly reactive Fe(IV)-oxo species catalyzes fLuther oxidative
chemistry and the metal returns to its resting Fe(H) state as products dissociate.

125

 

 

0H2
O
H.
M H20/ln,_ ('eF +2 ‘\\ ‘81
-00C 000- \Asp
8"" His:
-000 O2 02
/o.
O/
—OO/lll.le -i>‘3\\\His113 A
O— ‘9 ASP115
HIS
-OOC
O--0\
3\\\\H|5113
O o-e\/| ASP115
His
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L 0’
®Fe [+4\\\Hi3113
-OOC
/\/coo-'F l e\Asp115
His
i //O
S- C
//
HO 9 .
Q.,3.\\H'$113
-ooc , e
/\/coo-’ | ‘Aspns
His
0H2
- His
8 OH HZO/llli..Fe+2‘\\\ 1
Hp" \Asp
H152

N H-AA
”Ill/HS

O NH-AA
N\/H\"’//H
H0 is
-OOC \\\ \O \ei +3 \\\H15214
H3O CH3 (I\
H20 Hisz7 A59216

H

NH-AA

N ”III/H

. HKJS

-ooc\‘\\ VON} I)
H3O

H

e+4-I\\HISZ14

H20 _ ASPzie
Hl5270

NH-AA

N "’//H
H
5‘ 3
-00C
H3O CH3

Figure 17 . Comparison of the proposed mechanisms for an a-KG—dependent

hydroxylase and IPNS

126

essential residues defined. Multiple studies of several enzymes have thoroughly
characterized the iron ligands in the resting enzyme (2 His, 1 Asp, 3 waters). The identity
and geometry of the ligands to the metal during the course of the reaction, however, have
not been characterized. The order of dissociation of succinate and C02 (or bicarbonate)
suggests that these products remain in the active site until after the second oxidation
reaction occurs, however, no role for either product has been identified. The bicarbonate
dependence of ACC oxidase, an enzyme related to IPNS that does not use OL-KG as a co—
substrate, may indicate the involvement of bicarbonate in the catalytic cycle of other
enzymes. In addition, mutagenesis studies suggest that other amino acids present at the
active site participate in the reaction; these residues need to be identified and
characterized.

The second set of critical experiments needed to support the proposed model is
detection of the oxygen intermediates to confirm their identity. Researchers in our lab and
others have proposed experiments that may detect Fe(HI)-containing intermediates (e. g.
with bound superoxide or peroxide species) that may be generated during catalysis.

F e(HI) is paramagnetic and therefore can be detected by EPR spectroscopy of freeze—
quenched samples. Alternatively, stopped-flow UV-visible spectroscopy can also be used
to detect reactive intermediates by their characteristic chromophores. Although detection
of an Fe(IV)—oxo species may be difficult due to its high reactivity, model compound
studies or the use of non-reactive substrate analogs might trap the reactive oxygen
intermediate and aid in its identification.

The third area of study that is important for the creation of a robust model

involves the interactions between the unique substrate and the F e(lV)-oxo species. Apart

127

from IPNS, which has been structurally characterized with different substrate analogs,
little is known about the positioning of the substrate or its interaction with the metal in
other enzymes. a-KG-dependent dioxygenases demonstrate an amazing flexibility in the
number and types of substrates that can be oxidized in a single catalytic center. For
example, thymine 7-hydroxylase, which catalyzes the hydroxylation of methyl group of
thymine, can also epoxidize olefins, oxidize thiol ethers to sulfones and sulfoxides, and
demethylate the l-methyl group of 1-methyl thymine (3 5). Similarly, clavaminate
synthase catalyzes a hydroxylation, a desaturation and a ring expansion on three different,
though related substrates (Figure 2) (20). In this enzyme, the regio- and stereospecificity
of the three reactions suggest significant differences in the binding modes for the
structurally similar substrates (20). Alternatively, it has been proposed that clavaminate
synthase may modulate its reactivity by altering the position of the F e(IV ) oxo-
intermediate in response to different compounds. These experiments clearly show that the
substrates themselves play a significant role in influencing the aspects of the reaction;
however, the rules governing these interactions are not known. Information about the
importance of different substrate characteristics can be obtained by studying the products
of alternative substrates, spectroscopic characterization of the enzyme in the presence of
different compounds, and ultimately by crystal structure studies of enzyme-substrate
complexes.

The site-directed mutagenesis studies described in Chapter 2 lay the foundation
for examination of some of the questions outlined above. Knowledge of the metallocenter
ligands are not only important in the design of model compounds, but are necessary for

the interpretation of spectroscopic data and ultimately for our understanding of the

128

catalytic chemistry. For example, with both metal-bound histidines identified, we can
now learn more about the rearrangements that occur upon substrate binding and how the
substrate is positioned with respect to the metal. Furthermore, the ligands also play a
crucial role in controlling the reactivity of the metal. For example, enzymes with heme-
bound iron, such as cytochrome P450s, stabilize their iron(IV)—oxo intermediate with the
porphyrin ring. In the absence of such a cofactor, oc-KG—dependent dioxygenases may
rely on histidines to delocalize the charge on the metal. This potential role for histidines
may explain why glutamine or carboxylates are poor substitutes, and may suggest that
other amino acid substitutions (His to Tyr or Cys) may be capable of sustaining activity.
Continuing efforts to spectroscopically characterize deA (and the related enzyme TauD)
with different amino acid substitutions or different substrates and substrate analogs will
address some of these important questions. In addition, for the past four years, I have
supplied our collaborator Peter Roach at Oxford University with purified deA for use in
crystallization studies. Although crystals were obtained, crystallization conditions are still
being optimized. Structural characterization has not yet been successful. If a crystal
structure of deA or a closely related enzyme becomes available, more specific

experiments can be designed to probe the catalytic mechanism.

Biological significance and potential applications of oc-KG-dependent dioxygenases

Several or-KG-dependent dioxygenases are already being studied for their
potential applications to the fields of medicine, industry, and agriculture. Prolyl and lysyl
hydroxylases have long been studied for their role in collagen formation. Hydroxylated
proline and lysine residues, which comprise between 0.5—5% of the amino acids in

collagen or collagen-like proteins, are necessary to stabilize its triple helical structure

129

under physiological conditions (18). People with defects in lysyl hydroxylase suffer from
Ehlers-Danlos syndrome, which is characterized by hypermobility of the joints,
hyperextensibility of the skin, and other traits indicative of a connective tissue disorder
(18). The excessive accumulation of collagen, termed fibrosis, can also impair function in
a variety of tissues including the liver, lungs, and kidneys. Although several different
inhibitors and inactivators of collagen hydroxylases prevent collagen formation in
cultured human skin fibroblasts, none are currently in clinical use (18). In industry, the
enzymes involved in the synthesis of B-lactam antibiotics are being scrutinized for
opportunities to improve the large-scale synthesis of these complex molecules. Due to
their intricate structure which defies chemical synthesis, all penicillin and cephalosporin
antibiotics are still produced as natural fermentation products (31). Research on IPNS,
DAOCS and related enzymes has focused on improving the specificity, rate and range of
precursor substrates in hopes of optimizing steps in the synthesis and purification of these
chemicals in addition to creating additional derivatives with different biological
properties. Finally, in agriculture, deA has already been used to create transgenic plants
that are resistant to the broad—leaf herbicide 2,4—D and related phenoxyacetic acids (1). It
has also been proposed that deA could be used in the preparation and processing of
phenoxy herbicides (29). Since only one isomer of chiral 2,4-D derivatives, such as
mecoprop, is biologically active, deA-containing microorganisms or immobilized
enzyme could be used to remove the inactive isomer from a chemically-synthesized

racemic mixture, thus reducing the load of xenobiotics to soils.

130

Role of deA in the environment

Enzymes involved in the degradation of xenobiotic compounds are generally
derived from proteins that act on chemically-similar, natural compounds. For example,
benzoate and chlorobenzoate dioxygenases are related in both sequence and mechanism
but have differing reactivities towards chlorinated and unsubstituted substrates (24).
Similar examples include phenol hydroxylases (9), catechol dioxygenases (8), and
benzene dioxygenases (2). deA acts on a variety of natural and synthetic substrates
(Figure 18), but none with kinetic parameters approaching those for 2.4-D (R.
Padmakumar, J. Hotopp, R. P. Hausinger, unpublished data). One abundant source of
natural compounds that resemble known substrates of deA is lignin (9). This compound,
the second most abundant polymer on earth, is an assemblage of aromatic units joined
primarily through ether linkages. The aerobic degradation of lignin begins with
“combustion” of the polymer to its constituent subunits by the action of fungal
peroxidases to release a variety of monomeric and dimeric compounds. The types and
relative percentages of the different intermonomer linkages found in native lignin are
shown in Figure 19, and indicate the range of monomeric and dimeric compounds that
are liberated during lignin decomposition. Three types of compounds in particular, B-aryl
ethers, derivatives of cinnamic acids (including the corresponding alcohol (coumaryl
alcohol) and cinnamic acid esters through the side-chain carboxylate), and phenoxyacetic
acids bear particular resemblance to known substrates of deA (17).

One approach to determining if deA-like enzymes are involved in alternative

degradative pathways, such as lignin degradation, is to examine the roles of proteins

 

phenoxyacetic acids naphthoxyacetic acid thiophenoxyacetic acid
0
I \ \ 0' W
/ / O O-
X
cinnamic acids benzofurancarboxylic acid

(couman'llic acid)

Figure 18. Structures of various substrates of deA

OCH3
fHZOH
H
fHSOH fl H2 /O\CH
H CH0 H OOH OOH T |
F r—r
2 0 2
“r“ O 0 w
fH H3CO OCH,
0— H
OH OH
OCHg H3CO
OH OH
B-aiyl ethers (51%) phenylcoumarane (12%) biphenyls (l 1%) pinoresinol (2%)

Figure 19. Representative structures of the major intermonomeric linkages
in lignin. The relative percentages of these linkages are shown in parentheses (17, 22).

132

 

encoded by nearby genes. Efforts are currently underway in our laboratory to clone a
tfdA-like gene, described in Chapter 4, from isolates that do not appear to degrade 2,4-D.
The extragenic sequence of these elements may provide clues about the in vivo role of
this gene. Alternatively, the surrounding regions of sequences encoding proteins with
high identities to deA may indicate their potential functions in nature. In Bordetella
pertussis, the causative agent of whooping cough, there is a hypothetical protein with
42% identity to deA over the entire amino acid sequence (Figure 20). Interestingly, the
adjacent open reading frame, which is divergent from the tfdA-like gene, is 37% identical
in amino acid sequence to a glutathione-S-transferase (GST) from Sphingomonas
(formerly Pseudomonas) paucimobilis SYK-6 strain cleaves the B-aryl ether linkage of
dimeric lignin components (Figure 21) (22, 23). Other open reading frames adjacent to
the putative dioxygenase gene in B. pertussis are also homologous to enzymes involved
in aromatic degradation (Figure 20), including a LysR-type transcriptional regulator (37)
and a sequence homologous to clcE, which encodes maleyl acetate reductase (16).
Although B. pertussis has not been found in the environment, Bordetella spp. have been
isolated from soils (7, 33) and paper effluents (6), and at least one of these strains
degrades 2,4-D (7).

Apart from lignin degradation, GSTs are associated with a number of other
pathways involved in degradation of aromatics. Genes encoding GSTs are found in
degradative pathways for biphenyl (Bth) in Burkholderia sp. LB400 (l2), 2,4,5—
trichlorophenoxyacetic acid (the product of orf3) in Burkholderia cepacia AC1 100 (5),
and polycyclic aromatic hydrocarbons (Xle) in Cycloclasticus oligotrophicus (41 ).

Although gene disruption studies demonstrate that these enzymes are not required for

133

 

_ ORFl-putative OL-KG-dependent dioxygenase (287 aa) with 42% identity to deA (M16730)
ORF2-putative glutathione S-tranferase (203 aa) with 37% identity B-etherase, LigE
ORF3-putative LysR-type transcriptional regulator with 29% identity to N-terminus of SalR(3l6 aa).
ORF4-putative maleyl acetate reductase with 37% identity to ClcE (329 aa) (Q47100)

Figure 20. Open reading frames adjacent to a putative oc-KG-dependent
dioxygenase in Bordetella pertussis. Genbank accession numbers are shown in
parentheses.

activity, the multiple occurrences of GST—encoding genes in aromatic degradation
operons suggest that they do serve a purpose. Based on the activities observed for other
GSTs, proposed functions for the above enzymes include double bond isomerization,
dehalogenation, and epoxide cleavage (40), but none of these activities have been
demonstrated.

Despite the prevalence of lignin and its degradation products in soils, bacterial
decomposition of these compounds has not been well studied due to the difficulties in
working with such a large, heterogeneous substrate. The scheme for the degradation of
lignin-derived aromatics in S. paucimobilis SYK—6 demonstrates the number and
diversity of pathways and intermediates involved in this process (Figure 21) (22), but
certainly does not represent the only means for lignin mineralization (17). Of the few
previously characterized lignin-degrading isolates, most have generally belonged to the
Actinomycete group (3, 21, 34, 39). Although not enriched in cultures with polymeric
lignin as a substrate, a vast number of isolates including oc-, B-, and y—Proteobacteria and

numerous Gram positive bacteria degrade a variety of aromatic compounds, including

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135

biphenyls and aromatic acids (38, 39). Thus, it is not unlikely that more of these
organisms possess the ability to degrade at least a fraction of the available lignin
degradation products. As an aside, different S. paucimobilis strains are commonly
isolated from 2,4-D enrichment cultures (13-15), have deA activity (30), but lack a tfdA
gene with enough sequence identity to be picked up by gene hybridization methods (13).
When I began this research, there were no known enzymes with obvious sequence
similarity to deA. Thus, little could be said about important or conserved residues or
active site structure. Five years later, deA is related in sequence to a growing number of
characterized and putative dioxygenases (Chapter 1). Sequence alignments of deA to
two related sulfonate/OL-KG dioxygenases indicated a number of conserved residues that
proved to be important for activity (Chapter 2). Comparison of these findings to studies
of other enzymes in the superfamily shows both similarities in metallocenter ligands and
differences in overall protein structure among the a-KG-dependent dioxygenases. In
Chapter 4, I reported the intriguing presence of a tfdA-like gene in diverse soil isolates
that do not degrade 2,4-D suggesting that this gene is maintained for a function other than
in herbicide degradation. Although no alternative substrate for deA in soil
microorganisms has been identified, there is circumstantial evidence for its involvement
in the degradation of plant-synthesized ether- and cinnamic acid-containing products
derived possibly from lignin. Further biochemical characterization of pJP4-derived deA
and dioxygenases from other environmental bacteria are ongoing in the continued search

for other oc-KG-dependent dioxygenases with activity similar to that of deA.

136

Literature cited

Bayley, C., N. Trolinder, C. Ray, M. Morgan, J. E. Quisenberry, and D. W.
Ow. 1992. Engineering 2,4-D resistance into cotton. Theor. Appl. Gen. 83:645-
649.

Beil, S., K. N. Timmis, and D. H. Pieper. 1999. Genetic and biochemical
analyses of the tee operon suggest a route for evolution of chlorobenzene
degradation genes. J. Bacteriol. 181:341-346.

Cespedes, R., L. Salas, I. Calderon, B. Gonzales, and R. Vienna. 1992.
Microbial and biochemical-characterization of a bacterial consortium isolated

from decaying wood by grth on a B-O-4 lignin-related dimeric compound.
Arch. Microbiol. 158: 162-170.

Cosper, N. J., C. M. V. Stalhandske, R. E. Saari, R. P. Hausinger, and R. A.
Scott. 1999. X-Ray absorption analysis of F e(II) and Cu(H) forms of an

herbicide-degrading a—ketoglutarate dioxygenase. J. Biol. Inorg. Chem. 4: 122-
129.

 

Daubaras, D. L., C. D. Hershberger, K. Kitano, and A. M. Chakrabarty.
1995. Sequence-analysis of a gene-cluster involved in metabolism of 2,4,5-
trichlorophenoxyacetic acid by Burkholderia cepacia AC1100. Appl. Environ.
Microbiol. 61:1279-1289.

Foss, S., U. Heyen, and J. Harder. 1998. Alcaligenes defragrans sp. nov.,
description of four strains isolated on alkenoic monoterpenes ((+)-menthene,
alpha-pinene, 2-carene, and alpha-phellandrene) and nitrate. Syst. Appl.
Microbiol. 21 :23 7-244.

Greer, L. E., J. A. Robinson, and D. R. Shelton. 1992. Kinetic comparison of
seven strains of 2,4-dichlorophenoxyacetic acid-degrading bacteria. Appl.
Environ. Microbiol. 58: 1027-1030.

Haggblom, M. M. 1992. Microbial breakdown of halogenated aromatic
pesticides and related compounds. FEMS Microbiol. Rev. 103:29-72.

Hausinger, R. P., F. Fukumori, D. A. Hogan, T. M. Sassanella, Y. Kamagata,
H. Takami, and R. E. Wallace. 1996. Biochemistry of 2,4-D degradation:
evolutionary implications, p. 35-51. In K. Horikoshi, M. F ukuda, and T. Kudo
(ed.), Microbial Diversity and Genetics of Biodegradation. Japan Scientific Press,

Tokyo.

137

 

20.

Hegg, E. L., R. Y. N. Ho, and L. Que Jr. 1999. Oxygen activation and arene
hydroxylation by functional mimics of oc-keto acid-dependent iron(II)
dioxygenases. J. Am. Chem. Soc. 121:1972-1973.

Hegg, E. L., A. K. Whiting, J. McCracken, R. P. Hausinger, and J. Que, L.
1999. The herbicide-degrading oc-keto acid enzyme deA: metal coordination
environment and mechanistic insights. Biochemistry In press.

Hofer, B., S. Backhaus, and K. N. Timmis. 1994. The biphenyl polychlorinated
biphenyl-degradation locus (Bph) of Pseudomonas Sp. LB400 encodes four
additional metabolic enzymes. Gene 144:9-16.

Ka, J. 0., W. E. Holben, and J. M. Tiedje. 1994. Genetic and phenotypic
diversity of 2,4-dichlorophenoxyacetic acid (2,4-D)-degrading bacteria isolated
from 2,4-D-treated field soils. Appl. Environ. Microbiol. 60:1 106—1 1 l5.

Ka, J. 0., W. E. Holben, and J. M. Tiedje. 1994. Use of gene probes to aid in
recovery and identification of functionally dominant 2,4-dichlor0phenoxyacetic
acid-degrading populations in soil. Appl. Environ. Microbiol. 60:1116-1120.

Ka, J. 0., and J. M. Tiedje. 1994. Integration and excision of a 2,4-
dichlorophenoxyacetic acid degradative plasmid in Alcaligenes paradoxus and
evidence of its natural intergeneric transfer. J. Bacteriol. 176:5284—5289.

Kasberg, T., D. L. Daubaras, A. M. Chakrabarty, D. Kinzelt, and W.
Reineke. 1995. Evidence that Operon-ch, operon-de, and operon-Clc encode
maleylacetate reductase, the fourth enzyme of the modified ortho pathway. J.
Bacteriol. 177:3885-3889.

Kirk, T. K., and R. L. Farrell. 1987. Enzymatic combustion - the microbial-
degradation of lignin. Ann. Rev. Microbiol. 41:465-505.

Kivirikko, K. I., and T. Pihlajaniemi. 1998. Collagen hydroxylases and the
protein disulfide isomerase subunit of prolyl 4-hydroxy1ases. Adv. Enzymol.
Relat. Areas Mol. Biol. 72:328-398.

Lippard, S. J., and J. M. Berg. 1994. Principles of bioinorganic chemistry.
University Science Books, Mill Valley, CA.

Lloyd, M. D., K. D. Merritt, V. Lee, T. J. Sewell, B. Wha-Son, J. E. Baldwin,
C. J. Schofield, S. W. Elson, K. H. Baggaley, and N. H. Nicholson. 1999.
Product-substrate engineering by bacteria: Studies on clavaminate synthase, a
trifunctional dioxygenase. Tetrahedron 55: 10201-10220.

138

'21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

Lodha, S. J., R. A. Korus, and D. L. Crawford. 1991. Synthesis and properties
of lignin peroxidase from Streptomyces viridosporus T7a. Appl. Biochem.
Biotechnol. 28-9:41 1-420.

Masai, E., Y. Katayama, S. Kawai, S. Nishikawa, M. Yamasaki, and N.
Morohoshi. 1991. Cloning and sequencing of the gene for a Pseudomonas
paucimobilis enzyme that cleaves B-Aryl ether. J. Bacteriol. 173:7950-7955.

Masai, E., Y. Katayama, S. Kubota, S. Kawai, M. Yamasaki, and N.
Morohoshi. 1993. A bacterial enzyme degrading the model lignin compound [3-

etherase is a member of the glutathione-S-transferase superfamily. FEBS Lett.
323: 135-140.

N akatsu, C. H., N. A. Straus, and R. C. Wyndham. 1995 . The nucleotide-
sequence of the Tn5271 3-chlorobenzoate 3,4—dioxygenase genes (cbaAB) unites
the class Ia oxygenases in a single lineage. Microbiol. UK 141:485-495.

Pavel, E. G., J. Zhou, R. W. Busby, M. Gunsior, C. A. Townsend, and E. 1.
Solomon. 1998. Circular dichroism and magnetic circular dichroism
spectroscopic studies of the non-heme ferrous active site in clavaminate synthase

and its interaction with a-ketoglutarate cosubstrate. J. Am. Chem. Soc. 120:743-
754.

Roach, P. L., I. J. Clifton, V. Fiiliip, K. Harlos, G. J. Barton, J. Hajdu, I.
Andersson, C. J. Schofield, and J. E. Baldwin. 1995. Crystal structure of

isopenicillin N synthase is the first from a new structural family of enzymes.
Nature 375:700-704.

Roach, P. L., I. J. Clifton, C. M. H. Hensgens, N. Shibata, C. J. Schofield, J.
Hajdu, and J. E. Baldwin. 1997. Structure of isopenicillin N synthase complexed
with substrate and the mechanism of penicillin formation. Nature 387:827-830.

Ryle, M. J., R. Padmakumar, and R. P. Hausinger. 1999. Stopped-flow kinetic
analysis of Escherchia coli taurine/a—ketoglutarate dioxygenase: interactions with
oc-ketoglutarate, taurine, and oxygen. Biochem. In press.

Saari, R. E., D. A. Hogan, and R. P. Hausinger. 1999. Stereospecific
degradation of the phenoxypropionate herbicide dichlorprop. J. Molec. Catalysis
B: Enzymatic 266:421-428.

Sassanella, T. M., F. Fukumori, M. Bagdasarian, and R. P. Hausinger. 1997.
Use of 4-nitrophenoxyacetic acid for the detection and quantitation of 2,4-
dichlorophenoxyacetic acid (2,4-D)/0L-ketoglutarate dioxygenase activity in 2,4-D
degrading microorganisms. Appl. Environ. Microbiol. 63: 1 1 89-1 191 .

139

31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

Schofield, C. J., J. E. Baldwin, M. F. Byford, 1. Clifton, J. Hajdu, C.
Hensgens, and P. Roach. 1997. Proteins of the penicillin biosynthesis pathway.
Ctur Opin Struct Biol 7 :857—864.

Shilov, A. E., and A. A. Shteinman. 1999. Oxygen atom transfer into C—H bond
in biological and model chemical systems. Mechanistic aspects. Accounts of
Chemical Research 32:763-771.

Sturz, A. V., B. R. Christie, B. G. Matheson, and J. Nowak. 1997. Biodiversity
of endophytic bacteria which colonize red clover nodules, roots, stems and foliage
and their influence on host growth. Biol. Fert. Soil 25:13-19.

Thomas, L., and D. L. Crawford. 1998. Cloning of clustered Streptomyces
viridosporus T7A lignocellulose catabolism genes encoding peroxidase and
endoglucanase and their extracellular expression in Pichia pastoris. Can. J.
Microbiol. 44:364-372.

Thornburg, L. D., M. T. Lai, J. S. Wishnok, and J. Stubbe. 1993. A non—heme
protein with heme tendencies: an investigation of the substrate specificity of
thymine hydroxylase. Biochemistry 32:14023-14033.

Valegz‘ird, K., A. C. T. van Scheltinga, M. D. Lloyd, T. Hara, S. Ramaswamy,
A. Perrakis, A. Thompson, H.—J. Lee, J. E. Baldwin, C. J. Schofield, J. Hajdu,
and I. Andersson. 1998. Structure of a cephalosporin synthase. Nature 394:805-
809.

van der Meer, J. R., A. C. J. Frijters, J. J. Leveau, R. I. L. Eggen, A. J. B.
Zhender, and W. M. De Vos. 1991. Characterization of the Pseudomonas sp.
strain P51 tch, a LysR-type transcriptional activator of the tchDEF
chlorocatechol oxidative operon, and analysis of the regulatory region. J.
Bacteriol. 173:3700-3708.

Vicufia. 1988. Bacterial degradation of lignin. Enzyme Microb. Techno]. 10:646—
655.

Vicuna, R., B. Gonzalez, D. Seelenfreund, C. Ruttimann, and L. Salas. 1993.
Ability of natural bacterial isolates to metabolize high and low—molecular—weight
lignin-derived molecules. J. Biotechnol. 30:9-13.

Vuilleumier, S. 1997. Bacterial glutathione S-transferases: What are they good
for? J. Bacteriol. 179:1431—1441.

Wang, Y., P. C. K. Lau, and D. K. Button. 1996. A marine oligobacterium

harboring genes known to be part of aromatic hydrocarbon degradation pathways
of soil pseudomonads. Appl. Environ. Microbiol. 62:2169—2173.

140

 

42.

Whiting, A. K., L. Que Jr., R. E. Saari, R. P. Hausinger, M. A. Fredrick, and
J. McCracken. 1997. Metal coordination environment of a Cu(H)—substituted Ot-
keto acid-dependent dioxygenase that degrades the herbicide 2,4-D. J. Amer.
Chem. Soc. 119:3413-3414.

141

 

APPENDIX A

AMINO ACID SEQUENCE ALIGNMENTS

Amino acid sequence alignments for Group 11 (Figure 22) and Group III (Figure
23) enzyme and different deAs (Figure 24) are shown. Alignments were performed
using either the PIMA (3) or CLUSTAL (5) algorithms, which are available through the
Baylor College of Medicine (BCM) Search Launcher (4). Figures were generated using

GeneDoc software (1, 2).

142

 

 

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Literature cited

Nicholas, K. B., N. H. B. Jr., and I. Deerfield, D.W. 1997. GeneDoc: analysis
and visualization of genetic variation. EMB News 4:14.

Nicholas, K. B., and J. Nicholas, H.B.GeneDoc: Analysis and Visualization of
Genetic Variationhttp://www.cris.com/~Ketchup/genedoc.shtml September, 10
1999

Smith, R. F., and T. F. Smith. 1992. Pattem-Induced Multi-sequence Alignment
(PIMA) algorithm employing secondary structure-dependent gap penalties for
comparitive protein modelling. Prot. Engineering 5:35-41.

Smith, R. F ., B. A. Wiese, M. K. Wojzynski, D. B. Davison, and K. C. Worley.
1996. BCM Search Launcher-—an integrated interface to molecular biology data
base search and analysis services available on the World Wide Web. Genome
Res. 6:454-462.

Thompson, J. D., D. G. Higgins, and T. J. Gibson. 1994. CLUSTAL W:
improving the sensitivity of progressive multiple sequence alignment through
sequence weighting, position-specific gap penalties and weight matrix choice.
Nucl. Acids Res. 22:4673—4680.

151

APPENDIX B

STEREOSPECIFIC DEGRADATION OF THE PHENOXYPROPIONATE
HERBICIDE MECOPROP

The following article, Saari, R. E., D. A. Hogan, and R. P. Hausinger (1999)
f‘Stereospecific degradation of the phenoxypropionate herbicide dichlorprop.” J.
Mol.Catal. B: Enzymatic (6): 421-428 reprinted with permission from Elsevier Science,
is the result of a collaboration between a former graduate student in the Hausigner lab,
Ruth E. Saari, and myself (D.A.H.). In this report, the stereospecific degradation of
dichlroprop, a chiral derivative of 2,4-D, is examined in cell extracts of Ralstonia
eutropha JMP134, Burkholderia cepecz'a strain RASC and a mecoprop-degrading strain
Alcaligenes denitrificans. These studies indicate that oc-KG-dependent dioxygenases with
altered substrate specificities exist in the environment and suggest that deA-like
enzymes can act on substituted phenoxyacetic acids that may more closely resemble the

structure of aromatics liberated during the course of lignin degradation.

152

 

 

 

 

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JOURNAL OF
MOLECULAR
CATALYSIS
B: ENZYMATIC

EISEVIE

Journal of Molecular Catalysis B: Enzymatic 6 (I999) 42 I —428

 

Stereospecific degradation of the phenoxypropionate herbicide
dichlorprop '

Ruth B. Saari a'b, Deborah A. Hogan 3‘, Robert P. Hausinger “b“

" Center for Microbial Ecology. I60 Giitner Hall, Michigan State University, East Lansing, MI 48824, USA
b Department of Biochemistry, I 60 Gilmer Hall, Michigan State University, East Lansing, MI 48824, USA
C Department of Microbiology, 160 Giltner Hall, Michigan State University, East Lansing, MI 48824. USA

Received 31 August 1998; accepted 7 December 1998

 

Abstract

2,4-Dichlorophenoxyacetic acid (2,4-D)/o¢-ketog1utarate (oz-KG) dioxygenase, deA, from Ralstonia eutropha JMP134,
was purified from recombinant cells and shown by gas chromatographic and colorimetric methods to degrade only the S
enantiomer of dichlorprop, a phenoxypropionate herbicide. Similarly, cell extracts of Burkholderia cepacia RASC,
containing a biochemically and genetically related or-KG—dependent dioxygenase, also were shown to oxidize (5)-dichlor-
prop using chiral HPLC and colorimetric methods. In contrast, cell extracts of a mecoprop—degrading strain of Alcaligenes
denitri/icans were shown to catabolize (R)-dichlorprop. Although the A. denitrificans activity exhibited stereospecificity
opposite to that of the JMP134 and RASC strains, its cofactor requirements were found to be characteristic of an
a-KG-dependent dioxygenase. A PCR amplification product from the DNA of this strain was shown to encode an amino
acid sequence that was 95% and 86% identical to the conesponding region of deA in RASC and JMP134, respectively.
Thus, closely related herbicide-degrading gene products appear to be capable of exhibiting opposite stereochemical
degradative capabilities. © 1999 Elsevier Science B.V. All rights reserved.

Keywords: Stereospecificity; Herbicide; Dichlorprop; a-Ketoglutarate; deA

 

1. Introduction a chiral carbon atom, phenoxypropionates can

exhibit either of two enantiomeric forms be—

 

Ring—substituted phenoxyacetates and phen-
oxypropionates are widely used as herbicides to
control the growth of broadleaf weeds and
woody plants [1]. While phenoxyacetates such
as 2,4-dichlorophenoxyacetic acid (2,4-D) lack

' Corresponding author. Michigan State University, Center for
Microbial Ecology, Department of Microbiology, 160 Giltner
Hall, East Lansing, MI, USA. Tel.: + 1-517-353-9675; Fax: + 1-
517-353—8957; Email: hausinge@pilot.msu.cdu

cause of the methyl group attached to the C—2
carbon atom. Notably, only the R enantiomers
are active towards plants [2]. A current trend in
agriculture is increasing use of only the active
enantiomer of chiral herbicides in order to re-
duce environmental loads of organOpesticides
[2]. For example, about 75% of dichlorprop
(2-(2,4-dich1orophenoxy)pr0pionate) and meco-
prop (2-(2-methy1—4—chlorophenoxy)propionate)
are sold in their active enantiomeric forms in
the EurOpean Union, with the expectation that

1381-1 177/99/$ - see front matter © 1999 Elsevier Science B.V. All rights reserved.

PII: 81381-1 177(99)OOOO8-9

153

422 R. E. Saari et al./Joumal ofMolecular Catalysis B: Enzymatic 6 (1999) 421—428

within 5 years only the R isomer will be com-
mercially available (R. Akers, personal commu—
nication).

Phenoxyacid herbicides are rapidly degraded
in soil by bacteria and fungi [1]. One of the best
characterized herbicide-degrading organisms is
Ralstonia eutropha (formerly Alcaligenes eutro-
phus [3]) strain JMP134, capable of using 2,4-D
as its sole carbon and energy source. The first
step in 2,4-D catabolism is its conversion to
2,4-dichlorophenol (2,4-DCP), catalyzed by the
enzyme 2,4-D /a-ketog1utarate (or-KG) dioxy-
genase [4] or deA [5]. This ferrous ion-depen-
dent enzyme couples the oxidative decarboxyla-
tion of a-KG to hydroxylation of the substrate
at the C-2 carbon to yield a postulated hemiac-
etal intermediate which spontaneously decom—
poses to form 2,4-DCP and glyoxylate [4]
(Scheme 1). The chlorinated phenol is subse-
quently metabolized by a series of well-studied
enzymes that eventually generate an intermedi-
ate .of the tricarboxylic acid cycle [6]. It is
important to emphasize, however, that not all
2,4-D-degrading microorganisms use this path-
way. The biochemistry of phenoxypropionate
degradation is much less well understood than
that of 2,4-D. Here too, however, side chain
removal to release the corresponding phenol has
been shown to be an essential step in catabolism
of these compounds (e.g., Refs. [7—14]).

Studies to characterize the stereospecificity
of enzymes that degrade phenoxypropionate
herbicides can have important biotechnological
ramifications. For example, a synthetic process

a-KG succinate

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2.4-D

—

V

could be devised in which a particular herbicide
is initially synthesized as the racemic mixture,
the S enantiomer is degraded by a stereoselec-
tive enzyme, and the remaining R isomer is
purified away from the other reaction products
which are recycled. As an alternate illustration,
the genes encoding an enzyme specific for the
R isomer could be used to develop genetically
engineered crops that are resistant to the herbi-
cide of interest. Genetic engineering has already
been used for crop protection from (achiral)
phenoxyacetate herbicides: cotton has shown
enhanced tolerance towards 2,4-D when tfdA
was transformed directly into the plant [15] or
into root-associated microbes [16].

Here, we demonstrate that purified R. eu-
tropha deA specifically degrades the S enan-
tiomer of dichlorprop and characterize the kinet-
ics of this process. In addition, we establish that
Burkholderia cepacia strain RASC, a 2,4-D—de-
grader containing a deA with 80% sequence
identity to that from JMP134, has the same
stereopreference for the S enantiomer. By con-
trast, we show that cell extracts of a strain of
Alcaligenes denitrificans, intact cells of which
are lmown to utilize the R enantiomer of meco-
pr0p [14], specifically degrade the R enantiomer
of dichlorprop in an a-KG-dependent process.
Although the latter activity exhibits opposite
stereospecificity from that found in the 2,4-D-
degrading strains, we provide evidence that a
tfdA-like gene is present in A. denitrificans,
suggesting that all of these enzymes are closely
related to each other.

C1 0
__, O glyoxylate
spontaneous

.. it
© ..

 

2,4-DCP

Scheme 1. Reaction catalyzed by deA with 2.4-D.

154

RE. Saari et al. / Journal of Molecular Catalysis 8: Enzymatic 6 ( 1999) 421—428 423

2. Experimental
2. I . Chemicals

(+)-(R)-Dichlorprop was a kind gift from
BASF. Racemic and (+)-(R)—mecoprop were
generously donated by AH. Marks. Racemic
dichlorpr0p and 2,4—D were purchased from
Aldrich.

2.2. Bacterial strains

R. eutropha JMP134 [17] and B. cepacia
RASC [18] were obtained from the Research on
Microbial Evolution laboratory at Michigan
State University. A strain of A. denitrificans
isolated for its ability to degrade mecoprop [14]
was obtained from Hilary Lappin-Scott (Univer—
sity of Exeter).

2.3. Activity assay with colorimetric detection of
derivatized phenols

The standard assay mix contained 1 mM
a-KG, 50 uM each of Fe(II) and ascorbate, 1
mM phenoxyacid substrate in 10 mM imidazole
buffer (pH 6.75), and enzyme. In studies of cell
extracts, the concentrations of selected reagents
were varied in some experiments as noted in the
text. The iron and ascorbate were either sup-
plied together as iron(II) ascorbate salt or sepa~
rately as (NH4)2Fe(SO4)2 and ascorbic acid.
Reactions were carried out at 30°C and termi-
nated by addition of EDTA to a final concentra-
tion of 5 mM. The phenolic product was de-
tected at 510 nm following addition of 4-
aminoantipyrene, pH 10 borate buffer, and
potassium ferricyanide, as described [19]. Mea-
sured extinction coefficients for the 4-aminoan-
tipyrene adducts were 15,700 M"1 cm‘1 for
2,4-DCP and 11,300 M"1 cm‘l for 4-chloro-
2-methy1phenol.

2.4. Preparation of cell extracts and purified
enzyme

deA was purified from Escherichia coli
DH5a cells carrying the cloned R. eutropha

JMP134 tfdA gene on plasmid pUS311 by se-
quential chromatography on DEAE Sepharose
and MonoQ (Pharmacia) columns [4,20].

2,4-D and mecoprop degraders were grown
aerobically at 30°C in MO mineral salts
medium [21] with 50 ppm yeast extract and the
relevant herbicide at initial concentrations be—
tween 1 and 5 mM. Cells were harvested when
60—90% of the herbicide was consumed as mea-
sured by disappearance of the chromophore at
230 nm. Cell extracts were prepared by resus-
pending the bacteria in Tris buffer (pH 7.7) with
1 mM EDTA followed by disruption in a French
pressure cell and ultracentrifugation (100,000 X
g, 30 min) or centrifugation in a microcentri-
fuge (16,000 X g, 5 min). Protein concentra-
tions were determined by using the BioRad
protein assay with bovine serum albumin as the
standard.

2.5. Formation of diastereomers and gas chro—
matography (GC)

The levels of (R)— and (5)-dichlorprop in
samples were assessed by using a modification
of a published GC method [22]. Aqueous reac-
tion mixtures (1 ml) were acidified with 25 it]
of HCI (cone) and extracted with 1 ml methy-
lene chloride. The organic phase (0.5 ml) was
dried under a gentle stream of air, 1 drop of
thionyl chloride was added, and the sample was
incubated 10 min over a steam bath. Following
removal of the thionyl chloride under vacuum,
the sample was dissolved in 0.1 ml chloroform
with 1 drop‘of (R)—phenylethy1amine added,
allowed to react 25 min at room temperature to
form diastereomers (Scheme 2), dried, and
transferred to 0.5 m1 of ethyl acetate. An aliquot
(1 ul) of the sample was injected onto a DB-5
column (polymethylsiloxane with 5% phenyl
groups; I &W Scientific), with a temperature
program set to increase linearly from 200 to
270°C in 7 min, and the analytes were moni-
tored by electron capture detection (rather than
the less sensitive flame ionization detection
originally used [22]). Peaks were identified by

155

424 R. E. Saari et al. /Journal of Molecular Catalysis 1?." Enzymatic 6 ([999) 421—428

 

(8)-dichlorprop (R)-phenylethylamine

 

C1

(R)-dichlorprop (R)-phenylethylamine

Scheme 2. Formation of diastereomers of dichlorprop and (R)-phenylethylamine.

comparison with the retention times of the di-
astereomeric amides of authentic (R)—dichlor-
prop and (RS )-dichlorprop.

2.6. High pressure liquid chromatography
(HPLC)

Dichlorprop was reacted with cell extracts or
purified enzyme as described for the activity
assay, and the reactions were stopped with 5
mM EDTA. Aliquots were injected onto a Nu-
cleodex a-PM column (4.0 mm id. x 200 mm
length; Machery-Nagel, Germany). The eluent
was 30% phosphate buffer (50 mM, pH 3) and
70% methanol at a flow rate of 0.7 ml min“.
Species eluting from the column were moni-

tored with UV detection at 230 nm.

2.7. DNA hybridization, amplification and se-
quencing

Total genomic DNA was isolated from A.
denitrificans [23] and digested with BamHI.
Following separation on a 0.8% agarose gel, the
DNA was transferred [23] to a Hybond N mem-
brane (Amersham). Probes for tfd/lRASC (a l-kb
PstI fragment [18]) and for tfdAJMPm (an 801-
bp StyI fragment [24]) were labeled with digox-
igenin according to the GeniusTM System in-
structions (Boehringer-Mannheim Biochemi-
cals). The membrane was blotted with the probes

at high stringency (68°C, 50% formamide) or
low stringency (68°C, no formamide) using the
protocol from the same manual.

A. denitrificans DNA was used as template
DNA for PCR amplification with TVU and
TVL primers which amplify a 360-bp intemal
fragment of both RASC and JMP134 tfdA [25].
The product was purified by using the GFX
PCR DNA and gel band purification kit (Phar-
macia Biotech) and was sequenced in both di-
rections by using dye terminator chemistry at
the MSU-DOE—PRL Plant Biochemistry DNA
Sequencing Facility with TVU and TVL as
primers. The nucleotide and amino acid se—
quences of the amplified product (not including
the primer sequence) were aligned to those of
JMP134 and RASC using protocols in the GCG
suite of programs [26].

2.8. Data analysis

Data were fitted to straight lines, exponential
decays, or square hyperbola using the program
KaleidaGraph (Synergy software). As well,
progress curves of 2,4-DCP production vs. time
were fitted to Eq. (1) by non-linear regression
using the same program.

A, = At(1 _ e~k(inact)t) (1)
In this equation, A, is the absorbance at time t,

A, is the final absorbance, and kinnct is the rate
of inactivation.

156

 

RE. Suari at al. /.lournal a/‘Il’lolecular Catalysis B: Enzymatic 6 (1990) 421—428 425

3. Results
3. l . 7721A stereospecificity

The stereospecificity of purified deA from
R. eutropha JMP134 was investigated using two
methods. The first approach used derivatization
with (R)-phenylethy1amine followed by GC
analysis to separate and quantitate the two di-
astereomers. During the reaction of racemic
dichlorprop with deA, the percentage of S
relative to R enantiomer decreased over time,
suggesting that this enzyme uses the S isomer
(Fig. 1). The second procedure used to assess
stereospecificity involved a colorimetric assay
for 2,4-DCP release. 2,4—DCP was produced by
deA acting on racemic dichlorprop but not on
(R)-dieh101prop (Fig. 2), a result in agreement
with the GC results for this enzyme. The en—
zyme lost activity over time due to oxidative
inactivation events that are minimized by the
presence of ascorbic acid [27]. Based on initial
rates, (5)-dichlorprop utilization (0.81 lLlllOl
min‘I mg‘I protein) was 6% of that observed
for 2,4-D decomposition (l3 panol min" mg"|
protein) under saturating substrate concentra—
tions.

 

100
80
60 _

40

20 _ \\\\\“ql
.. ‘\\I“J
.441111_LA_JJJ_AIIllLLl‘LlJlJJLAlllLlJl

0 5 10 15 20 25 30 35
time (min)

 

 

 

 

0

% (8)—dichlorprop relative to (R)—dichlorprop

Fig. l. Stereoselectivity of dichlmprop decomposition catalyzed
by deA. T'l'dA from JMP134 (50 pg) was incubated with racemic
dichlorprop (100 uM) and ascorbic acid, ferrous ion, and OI-KG
using standard concentrations for the indicated time periods. Fol-
lowing extraction, the remaining dichlorprop was converted to the
(R)-phenylethylamine diastereomers and analyzed by GC/elec—
tron capture detection. Data were fitted to an exponential decay
with a decay rate of 0.096i0.011 min _ '.

 

 

 

 

70'
60 '
E 50 I/AT/‘fi
g 40 / '
CD
a 30
N-
20
10
0 = —I = +
_10 lLitlLaiileLIIliLLliiiLJAJiJ
0 5 10 15 20 25 30

time (min)

Fig. 2. 2,4—DCP production by deA acting on dichlorprop. Tl'dA
from JMP134 (10 pg ml“ I) was incubated with a racemic
mixture of dichlorprop (A) at 2 mM or with (R)—dichlmprop at
the same concentration (fl). 2,4—DCP was measured colorimetri-
cally. Data for the racemic dichlorprop were fitted to Eq. (I)
(kmm = 0.131 i0.004 min"| ), while data for the (R)-dichlorprop
were fitted by linear regression (slope of *0.02 uM min" ' ).

3.2. Herbicide-degrac‘ling activities in cell ex—
tracts

Cell extracts were examined from JMP134
and two additional phenoxyacid herbicide de-
graders: B. cepacia RASC, a 2,4-D degrader
with a deA that is 80% identical to that of R.
eutropha JMP134 [l8], and a strain of A. deni—

 

 

 

 

_ 350
E
g 300,
2- Z
3% 250:-
“E’S :
“1'3 200:—
SE 3
2?; 150:-
:e =
a? 100.-
O -
u_ I
0 50
9
z. o LillaiiIIlixilnarillilnllirilnini

 

O 100 200 300 400 500 600 700
substrate (IIM)

Fig. 3. Kinetic analysis of A. (lenitrij‘icans cell extract activities
towards (R)—mecoprop and (1%)—dichlorprop. Cell extracts of A.
(lenitrificmzs were examined for their abilities to decompose
(R)-dichlorprop (ti) and (R)-mecoprop (9). Initial rates calcu-
lated from the phenolic product release (based on 6 min time
points, triplicate samples) were fitted to the Miehaclis—Menten
equation to yield values of Km = 91¢ 16 'iM and Vnm = 351 i 21
iiM min “I 111ng protein for (R)-dichlorprop and Km = 52:4
IiM with V”m =247—I_—6 |.LM min‘l mg"1 protein for (R)—
Inecoprop. Mecoprop velocity data were conected for ~1%
contamination with a phenolic compound presumed to be 4—
chloro-Z—methylphenol.

157

426 R. E. Saari er al. / Journal of Molecular Catalysis B: Enzymatic 6 (1999) 421-428

 

 

 

 

 

60 _
E» so: __ ‘ A
b
0 4O _-
i Z D
‘3 305—
0 .—
g 20}-
: >
g 10;-
o‘- _l 1 L L In J 1 1 l 1 I I
O 5 10 15 20
time(min)

Fig. 4. Stereospecificity of dichlorprop-degrading activity of A.
denitrificans cell extracts as measured by chiral HPLC. Concen-
trations of (5)-dichlorprop (A). (R)-dichlorprop (O), and 2,4-
DCP (0) were measured at selected time points after incubation
of A. denitrificans cell extracts (50 ul) with racemic dichlorprop
(0.1 mM) and the stande reaction components. Dichlorprop was
quantitated by using chiral HPLC and 2.4-DCP was quantitated
colorimetrically. Data for the (5)-dichlorprop were fit to a straight
line of slope 0.05 uM min" ' , for (R)-dichlorprop to exponential
decay with an exponent of —0.067 min‘ '; and for 2.4-DCP to
Eq. (1) with an apparent inactivation rate of 0.16 min".

trificans isolated for its ability to degrade (R)-
mecoprop [14]. Cell extracts of IMP134 and
RASC exhibited highest activity towards 2,4«D,
but showed ~ 20% of this activity towards
dichlorprop. We attribute the cell extract activ—
ity towards phenoxy herbicides in both these
strains to deA, since TnS disruption of the tfdA
gene in either strain will prevent the conversion
of 2,4-D to 2,4-DCP by intact cells [5,18]. By

contrast to the specificity of the abovemen-
tioned strains, A. denitnficans extracts were
most active with phenoxypropionates including
mecoprop, dichlorprop, 2-(2,4-dibromophen~
oxy)propionate, and 2-(4-chlorophenoxy)pro-
pionate, with only minor activity towards the
phenoxyacetates 2,4-D and 2—chloro-4-methyl-
phenoxyacetic acid (~13 fold less than the
activity towards dichlorprop). Cell extracts of
A. denitrificans demonstrated a slightly higher
me towards the dichlorprop than towards
mecoprop, the substrate on which it was iso-
lated, but slightly less affinity (higher K m) (Fig.
3). The A. denimficans activity was dependent
on a-KG and ferrous ion, and was rapidly lost
(in 2 min) in the absence of ascorbic acid.
These cofactor requirements are characteristic
of or~KG-dependent dioxygenases.

3.3. Stereospecificity of activity in cell extracts

The stereospeeificity of herbicide decomposi-
tion by cell extracts was measured using an
HPLC column with a chiral stationary phase,
which permitted direct separation without prior
derivatization of the enantiomers of dichlorprop.
In agreement with the results using purified
enzyme, cell extracts of JMP134 degraded the S
enantiomer. Similar results were obtained with

150 160 170 180 190 200
RAAYDALPRDLQSELEGLRAEHYALNSRFLLGDTDYSEAQRNAMPPVNWPLVRTHAGSGRK
IIIII II:I:::II:IllllIlllIIlI:IlIIIIII:IIIII|II:IIIIIIIIIIIII
A . deni tri fi cans RAAYDDLPEDFKKELQGLRAEHYALNSRFI LGDTDYSESQRNAMP PVSWPLVRTHAGSGRK
llllIIIIIIIIllIllttlIllllzlllllllI:tll|l|lllll|lIII:IIIIIIIII

JMP134

RASC RAAYDDLPEDFKKELQGLRAEHYALH S RF I LGDTEYS ESQRNAMPPVSWPLI RTHAGS GRK
170 180 190 200 210 220
2 l O 2 2 0 2 3 0 2 4 O 2 5 0
JMP13 4 FL FI GAHASHVEGLPVAEGRMLIAELLEHATQREFVYRHRWNVGDLVM

IIIIIIII:I=II lltlllllllllIlIlllI:IlIIl Izllllll

A. deni tri fi cans FLFI GAHAGHIEGRPVAEGRMLLAELLEHATQRKFWRHSWKVGDLVM
IIIIIlllzlllllllllllltllllllllll Illllllllllllll

RASC FLFIGAHASHIEGRPVAEGRMLLAELLEHATQPKFVYRHSWKVGDLVM
230 240 250 260 270

Fig. 5. Comparison of an A. denitrificans deA-like sequence with the enzymes from RASC and JMP134. The deduced sequence encoded
by the PCR amplification product arising from A. denitrificans DNA was compared to the indicated portions of the RASC and JMP134
deA sequences. Identical amino acids are indicated by a solid vertical bar and similar amino acids by a dashed bar.

158

R.E. Saari et al. / Journal of Molecular Catalysis B: Enzymatic 6 (1999) 421 —428 427

RASC extracts (data not shown). By contrast,
A. denitrificans extracts degraded (R)-dichlor-
prop (Fig. 4) with concurrent 2,4-DCP produc-
tion.

3.4. Molecular biological comparison of strains

The similarity in cofactor requirements among
the three strains raised the question of whether
the A. denitrificans activity was due to a gene
related in sequence to those which encode the
two deA enzymes in the other species. A.
denitrificans genomic DNA hybridized at low
stringency to a probe for tfdAJMPm and at high
stringency to a segment of tfdARASC (data not
shown). Amplification of A. denitrificans DNA
with primers specific for tfdA yielded a 360-bp
DNA fragment with a sequence that was 94%
identical to that from RASC and 78% to that
from JMP134, in a 327-bp overlap. The trans-
lated sequence (109 amino acids long) had 95%
identity with deA from RASC and 86% iden-
tity to that from JMP134 (Fig. 5).

4. Discussion

4.1. R. eutrophus JMP134 deA oxidizes the S
isomer of phenoxypropionates

a—KG dependent dioxygenases typically ex—
hibit stereoselectivity in the hydroxylations they
catalyze (e.g., Refs. [28—31]); thus, it was not
surprising to find that R. eutrophus JMP134
deA oxidizes a single enantiomer (the S iso—
mer) of dichlorprop. Based on this result with
phenoxypropionate, the enzyme likely catalyzes
stereochemically selective oxidation at the pro-
R position when acting on the prochiral 2,4—D.

4.2. Related gene products can degrade oppo—
site stereoisomers

The above studies demonstrate that the herbi-
cide-degrading activity of A. denitrificans ex-
hibits opposite stereochemistry than that found

in strains JMP134 or RASC; yet, each of] these
activities are a-KG-dependent and the enzymes

are likely to be genetically related. The present

work extends the conclusions derived from stud-
ies with a strain of Sphingomonas herbicidouo-
rans that is capable of degrading both enan—
tiomers of meCOprop [32]. In that strain, two
a—KG-dependent dioxygenases appear to be pre-
sent, each specific for a separate enantiomer of
the substrate [33]. Based on our findings, it is
probable that the two S. herbicidouorans en—
zymes are closely related in sequence. The abil-
ity for closely related enzymes to recognize
opposite stereoisomers has precedent. For ex-
ample, recent crystallographic characterization
of two tropinone reductases demonstrate the
presence of almost identical overall protein fold-
ing, but the closely related enzymes exhibit
opposite substrate stereoselectivities due to dif-
ferences in a few charged residues at the active
site [34].

4.3. Biotechnological implications

The work described here may have biotech-
nological applications. An S isomer selective
dichlorprop-degrading enzyme, such as deA,
may be used to facilitate the preparation of the
desired R isomer, while a gene encoding an R
isomer selective enzyme, such as that in A.
denitrificans, may be useful for development of
herbicide-resistant transgenic plants. Also, in
understanding the role of microorganisms in the
environmental fate of phenoxypropionates, it is
useful to have analytical techniques such as
those used in this research capable of distin-
guishing the enantiomers.

Acknowledgements

We thank Peter Chapman for suggesting that
dichlorprop could be used to probe the stere—
ospecificity of deA, Paul Loconto of the Engi-
neering Research Labs of MSU for assistance in
setting up the GC method and suggesting the

159

 

428 RE. Saari et al. /Journul ofMo/ecular Catalysis 8: Enzymatic 6 (1999) 421—428

use of electron capture detection, and Hilary
Lappin-Scott for providing us with the meco-
prop-degrading strain of A. denitrificans. This
work was supported by the National Science
Foundation (MCB9603520), the National Sci—
ence Foundation—Science and Technology Cen-
ter Grant DEB9120006, and the Michigan State
University Agricultural Experiment Station.

References

[1] MA. Loos, in: RC. Kearney, D.D. Kaufman. (Eds) Herbi-
cides: Chemistry, Degradation and Mode of Action, V01. 1,
Marcel Dekker, New York, 1969, p. 1.

[2] IUPAC, Pure Appl. Chem. 69 (1997) 1335.

[3] E. Yabuuchi, Y. Kosako, I. Yano, H. Hotta, Y. Nishiuchi,
Microbiol. Immunol. 39 (1995) 897.

[4] F. Fukumori, R.P. Hausinger, J. Bacteriol. 175 (1993) 2083.

[5] W.R. Streber, KN. Timmis, M.H. Zenk. J. Bacteriol. 169
(1987) 2950.

[6] RP. Hausinger, F. Fukumori, D.A. Hogan, T.M. Sassanella,
Y. Kamagata, H. Takami, RE. Wallace, in: K. Horikoshi, M.
Fukuda, T. Kudo (Eds). Microbial Diversity and Genetics of
Biodegradation, Japan Scientific Press, Tokyo, 1996, p. 35.

[7] P. Ludwig, W. Gunkel, H. Hiinerfuss, Chemosphere 24
(1992) 1423.

[8] R. Martens, Pestic. Sci. 9 (1978) 127.

[9] M. Négre, M. Gennari, V. Andreoni, R. Ambrosoli, L. Celi,
J. Environ. Sci. Health B 25 (1993) 545.

[10] A.E. Smith, A.J. Aubin, Can. J. Soil Sci. 70 (1990) 343.

[11] A.E. Smith, Bull. Environ. Contam. Toxicol. 34 (1985) 656.

[12] A.E. Smith, J. Agric. Food Chem. 33 (1985) 483.

[13] A.E. Smith, J. Agric. Food Chem. 25 (1977) 893.

[14] V.A. Tett, A.J. Willetts, H.M. Lappin-Scott, FEMS Micro-
biol. Lctt. 14(1994) 191.

[15] BR. Lyon, Y.L. Cousins, DJ. Llewellyn, ES. Dennis,
Transgen. Res. 2 (1993) 162.

[16] L. Feng, l.R. Kennedy, Biotech. Bioeng. 54 (1997) 513.

[17] RH. Don, J.M. Pemberton. J. Bacteriol. 145 (1981) 681.

[18] Y. Suwa, A.D. Wright, F. Fukumori, K.A. Nummy, R.P.
Hausinger, W.E. Holben, L.J. Forney, Appl. Environ. Micro-
biol. 62 (1996) 2464.

[19] R.J. King, K.A. Short, R.J. Seidler. Appl. Environ. Micro-
biOl. 57 (1991) 1790.

[20] F. Fukumori, RP. Hausinger, J. Biol. Chem. 268 (1993)
2431 1.

[21] R. Stanier, N. Pelleroni, M. Douderoff, J. Gen. Microbiol. 43
(1966) 159.

[22] B. Blessington, N. Crabb, S. Karkee, A. Northage, J. Chro-
matogr. 469 (1989) 183.

[23] J. Sambrook, E.F. Fritsch, T. Maniatis. Molecular Cloning: A
Laboratory Manual, 2nd edn., Cold Spring Harbor Labora-
tory Press, Cold Spring Harbor, NY, 1989.

[24] WE. Holben, B.M. Schroeter, V.G.M. Calabrese, R.H. Olsen,
J.R. Kukor, V.O. Biederbeck, A.E. Smith, J.M. Tiedje. Appl.
Environ. Microbiol. 58 (1992) 3941.

[25] DA. Hogan, D.H. Buckley, CH. Nakatsu, T.M. Schmidt,
R.P. Hausinger, Microb. Ecol. 34 (1997) 90.

[26] Wisconsin Package Version 9.0, Genetics Computer Group
(GCG). Madison, WI.

[27] RE. Saari, RP. Hausinger, Biochemistry 37 (1998) 3035.

[28] J.E. Baldwin, R.A. Field, C.C. Lawrence, K.D. Merritt, C.J.
Schofield, Tetrahedron Lett. 34 (1993) 7489.

[29] J.E. Baldwin, K.D. Merritt, C.J. Schofield, S.W. Elson, K.H.
Baggaley, J. Chem. Soc., Chem. Commun. (1993). 1301.

[30] S. Englard, J.S. Blanchard, CF. Midelfort. Biochemistry 24
(1985) 1110.

[31] E. Leete, D.I-I. Lucast, Tetrahedron Lett. 38 (1976) 3401.

[32] C. Zipper, K. Nickel, W. Angst, H.-P. Kohler. Appl. Envi-
ron. Microbiol. 62 (1996) 4318.

[33] K. Nickel, M.J.-F. Suter, H.-P.E. Kohler. J. Bacteriol. 179
(1997) 6674.

[34] K. Nakajima, A. Yamashita, H. Akama, T. Nakatsu, H. Kato,
T. Hashimoto, J. Oda, Y. Yamada, Proc. Natl. Acad. Sci. 95
(1998) 4876.

160

 

 

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"‘tiiitittiiiit e

5