Conformational Dynamics Control Catalysis in Disparate Systems:
Structural Insights from DNA Repair and Antibiotic Biosynthetic Enzymes
by
Jeremy Wayne Setser
B.S., Chemistry (2008)
The University of Akron
OF TECHNOLOGY
1
JUN 3 0 2014
LIBRARIES
Submitted to the Department of Chemistry
in Partial Fulfillment of the Requirements for the Degree of
Doctor of Philosophy
at the
MASSACHUSETTS INSTITUTE OF TECHNOLOGY
June 2014
C 2014 Massachusetts Institute of Technology. All rights reserved.
Signature of Author .........
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Department of Chemistry
May 9,2014
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C ertified by ........................................................................................................................
.
Catherine L. Drennan
Professor of Chemistry and Biology
Howard Hughes Medical Institute Investigator and Professor
Thesis Supervisor
A ccepted by ................................
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Robert W. Field
Chairman, Departmental Committee on Graduate Students
This doctoral thesis has been examined by a
Committee of the Department of Chemistry as follows:
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Assistk1't Professor Elizabeth M. Nolan
Committee Chairman
Pfizer-Laubach Career Development Assistant Professor of Chemistry
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Professor Catherine L. Drennan
Research Supervisor
Professor of Chemistry and Biology
Howard Hughes Medical Institute Investigator and Professor
Signature redacted
Professor Leona D. Samson
Committee Member
Professor of Biological Engineering and Biology
Uncas and Helen Whitaker Professor
American Cancer Society Research Professor
2
Conformational Dynamics Control Catalysis in Disparate Systems:
Structural Insights from DNA Repair and Antibiotic Biosynthetic Enzymes
by
Jeremy Wayne Setser
Submitted to the Department of Chemistry
on May 9, 2014 in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy in
Biological Chemistry
ABSTRACT
Chemical reactions allow biological systems to function. The majority of these biochemical reactions
occur due to the work of protein catalysts known as enzymes. These biocatalysts are often thought of as
pre-formed, static 'locks' that bind, and subsequently transform, their substrate molecule 'keys'.
However, scientists are increasingly finding that dynamic movements of enzymes are a crucial aspect of
catalysis. One such example of a system that relies on conformational flexibility is the human DNA repair
protein alkyladenine DNA glycosylase (AAG). To efficiently repair DNA, AAG must search the millionfold excess of unmodified DNA bases to find a handful of DNA lesions. Such a search can be facilitated
by the ability of glycosylases, like AAG, to interact with DNA using two affinities: a lower-affinity
interaction in a searching process, and a higher-affinity interaction for catalytic repair. We have captured
crystallographic snapshots of AAG bound to DNA in both high- and lower-affinity states. These
depictions reveal several significant and unexpected protein structural rearrangements, providing
molecular insight into the DNA-searching process adopted by AAG. By combining these new insights
with existing biochemical and structural data, we are able to relate AAG to the big picture question of
how DNA binding proteins find their binding sites in the vast expanse of the genome.
In another study, a member of a biosynthetic pathway for antibiotic natural products, called kutznerides,
was shown to be dependent on conformational changes. The enzyme in question, KtzI, uses a bound
flavin cofactor, reducing equivalents from NADPH, and molecular oxygen to install a hydroxyl group on
the side-chain nitrogen of the amino acid L-ornithine, which is subsequently incorporated into the
kutzneride scaffold. KtzI was structurally characterized after being subjected to various chemical and
environmental factors, capturing the enzyme in several states along its catalytic trajectory. These states
suggest that a novel conformational change of both the protein backbone and the flavin moiety must take
place in order to complete the enzymatic cycle of KtzI. This drastic rearrangement was also shown to be
chemically interchangeable in the protein crystal, suggesting that these dynamic motions are catalytically
relevant.
Thesis Supervisor:
Title:
Catherine L. Drennan
Professor of Chemistry and Biology
Howard Hughes Medical Institute Investigator and Professor
3
ACKNOWLEDGEMENTS
I am indebted to so many people for so many things.
First, I would like to say thank you to Cathy for accepting me into your group, and believing I
could succeed. Joining a completely foreign research area was daunting enough to me; I cannot imagine
having the amount of trust needed to believe I could learn from the ground up. It really has been an
amazing experience over the past six years. I would also like to thank Liz Nolan, Leona Samson, and
Sarah O'Connor for their scientific insights and guidance during this time. A large part of my graduate
experience was built by the support of my fellow lab members, to whom I owe many thanks.
Graduate school is not easy. Experiments fail almost exclusively. Without the support of the
Drennan Lab members, past and present, this would have been a crippling undertaking. Thank you all. To
Nozomi Ando - I could say that I would have made it through the final years of graduate school without
my lab BFF, but I would probably be lying. Thank you for spending countless hours listening and
laughing with me. Your support and friendship bolstered me when I was at my most disillusioned. To
Peter Goldman - You were the first person I talked to in the Drennan Lab, and made me understand that a
regular dude could succeed in our lab. This realization was a large part of why I joined in the first place.
Even if the utter breakdown of your body ruined our chances at volleyball supremacy, you made our lab a
fun place to be. To Marco Jost - I was reluctant to accept the 'hot shot German' who knew everything
about crystallography coming into our lab, but you won me over eventually. I have no problem getting
your advice now, because I have come to understand you as the 'hot shot German' who knows everything
about everything. To Yan Kung and Christine Phillips Piro - Thank you for teaching me what it meant to
be a Drennan Lab member. To Mishtu Dey and Danny Yun - Thank you for your ever-patient guidance.
Ultimately, friends and family are the reason I made it through graduate school. I love you all. To
Emily Setser - Even though you will always be Emily Lippert to me, I am excited every time I remember
we are married. You have been my most important source of support for over six years. Thank you for
being the best thing that has happened to me. I could not have done this without you. I love you clown.
To Jeff Setser - Thank you for always being there to provide a brother's love when I needed it most. To
Diana Setser - I could not have asked for a better Sister-in-law, and the joy I have observed in your
household always reminds me that things will be OK. To Nola, June, and Oden Setser - Thank you for
being constant reminders that the future will be bright and joyful. To my Dad, Jim Setser - Thank you for
constantly making sure I was healthy and happy. To Mom and Dad Lippert - Thank you for always
treating me as your own Son. To Josh Slaga - We have come a long way since 8 th grade. Thank you for
handling all best friend-related duties for the last 15 years. To Alyssa Larson - The free stuff line at Sid
Pac may not have offered up any useful items, but it did provide me with a best friend here at MIT. Thank
you for being a constant source of support for five years and counting. To all CGSC members past and
present - Always remember that we actually make an impact around here. Keep it up. To all my other
friends here and elsewhere - Thank you for your support and all the fun times had at TGIF, the Muddy,
around Boston, in Ohio, and around the country; It has all been instrumental in getting me to this point.
Finally, this thesis would not have been possible without my Mother..She supported me in every
endeavor, in any way that she could...even if this meant letting her 'baby' move 650 miles away from
home. My Mother passed away in the middle of graduate school, and the period of time before, during,
and after this tragedy was the worst experience of my life. I was extremely lucky to have an advisor as
understanding as Cathy, as I spent much of my Mother's last months at her bedside. This time allowed me
to support and comfort her in what ways I could, and to spend what would end up being our last quality
time together. 'Grateful' does not even begin to express the amount of appreciation I have for my
Mother's wonderful influence. 'Sorrow' falls incredibly short of describing what I feel in her absence.
Therefore, I would like to dedicate this thesis in honor of the kind, strong, generous, and above all, loving
woman that was my Mother.
4
For Judy Ann Setser.
I love you Mom.
5
6
Table of Contents
ABSTRACT
ACKNOWLEDGEMENTS
3
4
Chapter 1. The Dynamism of DNA-binding and Flavin-dependent Enzymes
I.
I.1I
1.111
I.IV
SUMMARY
INTRODUCTION
DNA repair by human alkyladenine DNA glycosylase (AAG)
Natural product biosynthesis with FAD-dependent N-hydroxylases
FIGURES
Figure 1.1
The damage of DNA bases.
Figure 1.2
Base excision repair (BER) in humans.
Figure 13
Introduction to the flavin and nicotinamide cofactors.
Figure 1.4
General mechanism of flavin-dependent monooxygenases.
I.V
REFERENCES
13
13
15
16
19
23
Chapter 2. Structural Basis for the Inhibition of Human Alkyladenine DNA
Glycosylase (AAG) by 3,N 4-Ethenocytosine-containing DNA
11.1
SUMMARY
11.11 INTRODUCTION
11.111 RESULTS
AAG binding studies
Catalytic ability of AAG for eC containingDNA
Inhibition of AAG by cC containing DNA
Overall structure of the JI79AAG-EC DNA inhibitorcomplex
Protein-DNA interactions
Metal ion Mn2 , in the A79AAG-EC:G structure
Active site architectureof JI79AAG-EC DNA complex
II.IV DISCUSSION
II.V MATERIALS AND METHODS
J79AAG plasmid construction, creation of mutants, andprotein preparation
AAG protein expression andpurification
Preparationof oligonucleotidesand 32P-labeling
Gel mobility shift assays
DNA glycosylase assays
Competition DNA glycosylase assays
Crystallizationof the I79AAG-EC:G complex
Data collection and structure determination
II.VI ACKNOWLEDGEMENTS
27
28
30
34
37
42
7
TABLES & FIGURES
Table 11.1
Table 11.2
Table 11.3
Figure 11.1
Figure 11.2
Figure
Figure
Figure
Figure
Figure
II.3
11.4
11.5
11.6
11.7
Figure 11.8
Figure 11.9
Figure 11.10
43
Dissociation constant (Kd) values measured using gel shift assays; and 50% inhibitory
concentration (IC50) for the inhibition of A79AAG activity on EA:T 25-mer, measured
using competition DNA glycosylase assay at 37'C, in the presence of increasing
concentration of cold competitor 13-mer duplexes.
Data collection and refinement statistics of the A79AAG-DNA complex.
List of oligonucleotide primers used for the creation of A79AAG mutants by PCR based
site directed mutagenesis.
Biochemical characterization of AAG variants with oligomers containing etheno lesions.
Gel results of DNA glycosylase assays for truncated A79AAG on EA:T and EC:X
(X=G/A/T/C) 13-mer DNA duplexes used for crystallization.
Structure of A79AAG with EC inhibitor DNA.
Comparison of the overall structures of the A79AAG-DNA complexes.
Cation site in A79AAG structures.
Interaction of Tyr 162 and putative Mn 2 ' binding site in the structure of A79AAG-cC:G.
Active site architecture of AAG.
AAG binding pocket.
Activation of the leaving group by protonation.
A wall-eyed stereoview of the active site of AAG.
II.VII REFERENCES
58
Chapter 3. Searching for DNA Lesions: Structural Evidence for Lower- and
Higher-Affinity DNA Binding Conformations of Human
Alkyladenine DNA Glycosylase
III. SUMMARY
111.11 INTRODUCTION
111.111 RESULTS
Structural overview of asymmetric unit
A79AAG pseudo-duplex structure
A79AAG lower-affinity structure
III.IV DISCUSSION
III.V MATERIALS AND METHODS
AAG plasmidconstruction andproteinpreparation
Crystallization of A79AAG with single-strandedEC DNA
Data collection and structure determination
III.VI ACKNOWLEDGEMENTS
TABLES & FIGURES
Table 111.1
Figure 111.1
Figure 111.2
Figure 111.3
Figure III.4
Figure 111.5
Figure 111.6
8
61
62
64
67
70
72
73
Data collection and refinement statistics for A79AAG-DNA complex.
DNA adducts to which AAG binds with high affinity: Lesions (A) EC and (B) EA and (C)
one-base loop structures.
Structures of A79AAG bound to EC DNA.
Wall-eyed stereoview of the disordered loops of the lower-affinity A79AAG structure in
the context of the crystal lattice.
A79AAG shows no affinity for ssFC 13-mer by gel shift.
A79AAG structural comparisons.
Tyr162 contacts in lower-affinity A79AAG.
Figure 111.7
Figure 111.8
Figure 111.9
Comparison of the lower-affinity A79AAG structure with the high-affinity
A79AAG-EA:T structure.
Wall-eyed stereoviews of electron density in the lower-affinity A79AAG structure.
Proposal for how AAG can recognize DNA with two different affinities.
III.VIIREFERENCES
84
Chapter 4. Crystallographic Evidence for Drastic Conformational Changes
in the Active Site of a Flavin-Dependent N-hydroxylase
IV.I SUMMARY
IV.II INTRODUCTION
IV.III RESULTS
Quaternarystructure
Active site of reduced KtzI
Active site of oxidized KtzI
In crystallo conformationalchanges
IV.IV DISCUSSION
IV.V MATERIALS AND METHODS
Proteinexpression andpurification
Reconstitution of KtzI for crystallization
Crystallizationof KtzI
Re-reduction of oxidized KtzI crystals
Re-oxidation of reduced KtzI crystals
Data collection, structure determination,and structuralanalysis
IV.VI ACKNOWLEDGEMENTS
TABLES & FIGURES
Table IV.1
Table IV.2
Table IV.3
Table IV.4
Table IV.5
Table IV.6
Table IV.7
Table IV.8
Figure IV.1
Figure IV.2
Figure IV.3
Figure IV.4
Figure IV.5
Figure IV.6
Figure IV.7
Figure IV.8
Figure IV.9
Figure IV.10
87
88
91
97
106
112
113
Structures of KtzI and their respective resolution.
Data collection and refinement statistics for KtzI-FADred-NADP'-L-orn.
Data collection and refinement statistics for KtzI-FADox-NADP*-L-orn.
Data collection and refinement statistics for KtzI-FADred-ox-NADP'-L-orn.
Data collection and refinement statistics for KtzI-FADred-NADP'-Br.
Data collection and refinement statistics for KtzI-FADox-Br.
Data collection and refinement statistics for KtzI-FADox-red-NADP*-Br.
List of ordered residues and those modeled as alanine in KtzI structures.
Kutzneride scaffold cryptically incorporates the product of KtzI.
Sequence alignment of L-orn N-hydroxylases.
Proposed kinetic mechanism for L-orn N-hydroxylases.
Crystals of reconstituted KtzI.
Structure of KtzI in reduced and oxidized states.
Tetrameric assembly of L-orn N-hydroxylases.
Wall-eyed stereoview of the electron density for bent isoalloxazine ring.
Wall-eyed stereoviews of the active sites of reduced L-orn N-hydroxylases.
Wall-eyed stereoviews of the active sites of oxidized KtzI.
Structurally-based proposal for L-orn N-hydroxylase mechanism.
II.VII REFERENCES
138
9
Chapter 5. Outstanding Questions for AAG and KtzI
V.1
V.II
V.111
SUMMARY
Repairing DNA with AAG
Antibiotic biosynthesis with KtzI
FIGURES
Figure V.1
Figure V.2
Figure V.3
V.IV
Sequence alignment of our N-terminal truncation mutant (A79) and full-length (FL)
alkyladenine DNA glycosylase (AAG).
Kinetic mechanism for hydroxylation in (a) bold and (b) cautious flavin-dependent
monooxygenases.
Diverse flavin conformations are found in (a) KtzI and (b) para-hydroxybenzoate
hydroxylase (PHBH).
REFERENCES
Curriculum Vitae
10
143
143
145
149
152
153
11
12
Chapter 1. The Dynamism of DNA-binding and Flavin-Dependent Enzymes
I.I
SUMMARY
Chemical reactions are at the heart of life. Molecules combine and change to create new entities,
and life has emerged from, and mutated in response to, these chemicals. Many chemical reactions
are not favored, with some likely to occur only on the timescale of millions of years. To accelerate
these reactions in biological systems, enzymes are used. These biological catalysts function by
binding a substrate and hastening its transformation into a product. Many enzymes depend on
dynamic movements to catalyze their chemical reactions, and this dissertation will examine two
such cases. In Chapters 2 and 3, an enzyme that repairs DNA in humans will be discussed. The
subject of Chapter 4 will be a member of an assembly line of enzymes responsible for synthesizing
an antibiotic in a soil bacterium. These dissimilar systems are linked by their reliance on
conformational flexibility.
1.11
INTRODUCTION
Proteins comprise a significant portion of the cellular makeup, up to 55% in Escherichia
coli (1). These entities perform functions as wide-ranging as acting as a cellular garbage
disposal, in the case of proteolytic species like the 26S proteasome, to determining which cellular
entities are produced by binding DNA sequences, in the case of transcription factors like NF-KB,
to synthesizing vital components of the cell, like the over 30 enzymes involved in the
biosynthesis of cholesterol. Dysfunctions in any of these pathways are linked to disease in
humans, underlining their practical importance for our health and well-being. The proteins that
function to take a hypothetical substrate 'S', to its product 'P', are known as enzymes, and are an
extremely active area of research.
Enzymes use their amino acids, and in some cases, the help of organic or inorganic
species known as cofactors, to accelerate chemical reactions, thus acting as biological catalysts.
These biocatalysts have traditionally been viewed as having a fixed structure that is tuned to
precisely fit its substrate molecule, stemming from their initial likening to a "lock and key" by H.
Emil Fischer in the late
1 9 th
century. In more recent years, however, proteins have been
understood to be dynamic, and an emphasis has been placed on studying both small-scale (short
timescale) and large-scale (long timescale) movements, as this conformational flexibility is often
vital to a protein's function (reviewed in (2)). Various techniques including nuclear magnetic
resonance (NMR), small-angle X-ray scattering (SAXS), and fluorescence methods like those
13
based on Fbrster resonance energy transfer (FRET), are well-suited to study large-scale
conformational changes, as these techniques occur while the protein is in solution, mimicking the
freedom found in the cellular milieu (2). X-ray crystallography, which requires a protein to be
trapped in a crystalline lattice (often frozen at 100K), is less useful in this context, as the
protein's degrees of freedom are limited. What this method does provide, however, is a much
higher-resolution picture than the other techniques mentioned above. Therefore, X-ray
crystallography tends to trade the vital information on conformational flexibility for more
detailed molecular-level understanding. Indeed, a crystal structure provides very little in the way
of dynamic information. As we will show in this thesis, however, a combination of crystal
structures can illustrate the flexibility of a protein. In the following chapters, we describe the
ability to get the best of both worlds, obtaining information on protein dynamics by combining a
series of atomic-level snapshots, using two enzymes with very dissimilar roles as case studies.
In Chapters 2 and 3, we analyze the human DNA repair protein alkyladenine DNA
glycosylase (AAG). In Chapter 4, we will examine the L-ornithine N-hydroxylase from
Kutzneria sp. 744, KtzI. These two enzymes function in very different worlds. AAG repairs
damaged DNA in humans, and KtzI is a member of a host of enzymes that synthesize natural
product defenses for a soil actinomycete. Through our structural work, however, we have found
that these systems are linked through their reliance on conformational changes. For AAG, we
have snapshots of the enzyme both bound to an inhibitor DNA base (Chapter 2) and interacting
with DNA in a nonspecific manner (Chapter 3). These structures allow us to observe the
differences between these higher- and lower-affinity complexes, with most alterations found in a
region that is highly flexible in the absence of a bound DNA base. By compiling our data with
that of the field in general, we propose this dynamic region to be vital for AAG's ability to
search for its damaged-DNA substrates. For KtzI, we have determined several structures of this
enzyme along its reaction path, which indicate that a highly mobile active site, including
movements of the protein backbone and the flavin cofactor, is required for catalysis (Chapter 4).
We have been able to recapitulate these proposed conformational interchanges in the protein
crystal, showing that the required conformational flexibility is both chemically competent, and
able to occur in the "rigidified" environment of the crystalline lattice. Together, these studies
show large-scale dynamics to be a crucial aspect of enzyme catalysis, and further, that this
14
information can be obtained by a collection of X-ray crystallography snapshots put together in a
'flip-book'-type methodology.
1.111
DNA repair by human alkyladenine DNA glycosylase (AAG)
Damaged DNA must be repaired to allow faithful replication of genetic information for
healthy cell division and, thus, for life to occur. Ultraviolet light, ionizing radiation, xenobiotic
chemicals such as alkylating agents, and endogenous metabolites such as reactive oxygen and
nitrogen species (RONS) or methylating agents (e.g. S-adenosylmethionine, AdoMet) can harm
nucleic acids (3). The most common manifestations of DNA damage are known as DNA lesion
bases (4), where a canonical DNA base (Figure 1.1, top) is chemically altered (Figure 1.1,
bottom). These lesion bases occur at a rate of 10,000 per cell per day (4, 5), and affect base
pairing interactions and DNA energetics, causing transition (A<-+G; C<-+T) and transversion
(A,G-*C,T) mutations and, as errors are compounded, cell death (5, 6). Most DNA lesions are
repaired via base excision repair (BER), an endogenous pathway that removes and replaces the
damaged base in a series of steps (3). BER is highly conserved in all kingdoms of life (7), and in
humans these steps are catalyzed by the sequential action of a DNA glycosylase, AP
endonuclease I, DNA polymerase P, and DNA ligase I/III ((8, 9); Figure 1.2).
Alkyladenine DNA glycosylase (AAG) is a monofunctional DNA glycosylase involved
in the human BER process. Monofunctional glycosylases, which lack inherent endonuclease
activity, initiate BER by binding specific lesion bases and catalyzing the cleavage of the bond
between the base and its ribose sugar (N-glycosidic bond), releasing the free base and leaving
behind an apurinic/apyrimidinic (AP) site (Figure 1.2, 1). The repair of the AP site is completed
by the action of three subsequent enzymes as mentioned above (Figure 1.2, 2-4), which finally
restores the DNA to its undamaged state. AAG has been of particular interest due to its
involvement in remediating lesions brought on by chronic inflammatory diseases like ulcerative
colitis (10-12), which predispose individuals to cancer. Further, AAG represents an interesting
biochemical problem as it recognizes a wide variety of lesion bases (13-15), including substrates
that it can excise, such as 1,M-ethenoadenine (EA) and 7-methylguanine (7-meG) (Figure I.1),
and those that it cannot remove, including the inhibitory 3,N 4-ethenocytosine lesion (EC) (Figure
1.1). This biochemical puzzle led to a collaboration between members of the Drennan and
15
Samson groups here at MIT, and together we were able to biochemically and structurally
characterize AAG.
In one study (Chapter 2, (16)), we examined the ability of AAG to bind EC with
high-affinity, while being unable to excise this lesion, and determined the molecular structure of
this AAG-EC inhibitory complex. These data allowed us to propose a chemical rationale for the
inability of AAG to excise the eC lesion. We were also able to obtain a fortuitous
crystallographic snapshot of this enzyme with a self-assembled, "pseudo-duplex" piece of DNA.
In this structure, one AAG molecule in the asymmetric unit recognizes a EC lesion in a manner
similar to the high-affinity depiction described in Chapter 2, while the other AAG monomer is
making only nonspecific contacts with DNA, and thus is in a lower-affinity state. Taking these
and other studies together, we were able to provide evidence for a mechanism by which AAG
can find its lesion base substrates in the million-fold excess of undamaged DNA bases using a
flexible DNA binding region (Chapter 3, (17)).
I.IV
Natural product biosynthesis with FAD-dependent N-hydroxylases
Flavin-containing enzymes, or flavoenzymes, have been an intense area of study for
nearly a century (18). From the initial discovery of a yellow pigment in cow's milk in 1879 (19)
(which we now know to be riboflavin or vitamin B2 (Figure I.3a)), to the elucidation of flavin's
chemical structure (20, 21) and the first description of a flavoenzyme (22, 23) in the 1930s, this
field has grown and matured to the point that today over 18,000 articles found on PubMed use
the term "flavin". Flavoenzymes utilize either the flavin mononucleotide (FMN) or flavin
adenine dinucleotide (FAD) forms of flavin (Figure I.3a), and these cofactors can be used to
perform a wide variety of reactions. This chemical diversity is due to the reactive isoalloxazine
ring of the flavin, which can adopt three oxidation states differing by 1-electron, making it one of
the few species in biology that can mediate between both 1- and 2-electron transfers (Figure
I.3a). Herein, we will concentrate on a class of FAD-dependent enzymes known as hydroxylases
or monooxygenases, and more specifically a recently discovered sub-class that act upon the
amino acid L-ornithine (L-orn).
The FAD-dependent hydroxylases use molecular oxygen and reducing equivalents from
nicotinamide adeninine dinucleotide (phosphate) (NAD(P)H; Figure I.3b) to install a hydroxyl
(-OH) group on a nucleophilic substrate (as outlined in Figure 1.4). The catalytic cycle of these
16
enzymes is initiated by hydride transfer from the Pro-R C4 position of the nicotinamide of
NAD(P)H to the N5 position of the oxidized flavin isoalloxazine ring. Molecular oxygen can
then add to the reduced isoalloxazine via a radical-mediated reaction,
generating a
C4a-hydroperoxy species. The hydroxylated product is created after nucleophilic attack by the
bound substrate on the distal oxygen of the C4a-hydroperoxy, leaving a C4a-hydroxyflavin
intermediate. Finally, dehydration of the hydroxyflavin readies this cofactor for another round of
catalysis. Although this general mechanism is conserved in the majority of monooxygenase
enzymes (reviewed in (24-27)), there are intriguing differences observed in the N-hydroxylase
class. Herein, we explore the L-orn N-hydroxylase from Kutzneria sp. 744, called KtzI. This
enzyme is involved in the biosynthesis of cyclic peptides that have antimicrobial and antifungal
activity, which made it of great interest to both the Walsh and Drennan Labs. We have been able
to structurally characterize KtzI in several states along its reaction path, and by pairing these
snapshots with the biochemical and structural data already available for this enzyme-class, we
propose a structurally-based reaction mechanism that includes never-before-seen conformational
changes of both the protein backbone and the flavin cofactor. Further, we were able recapitulate
these conformational changes in the protein crystal, displaying their chemical competence. These
results are presented in Chapter 4.
17
18
FIGURES
Figure 1.1. The damage of DNA bases. Canonical DNA bases (top) are acted upon by a variety of
endogenous and exogenous sources, some of which cause damage. These damaged, 'lesion bases'
(bottom) are chemically altered (in red), commonly by alkylation (7-meG, 3-meT) or through interaction
with peroxide and aldehyde species (EA, EC). Endogenous pathways must repair damaged bases such that
mutagenesis and cytotoxicity are minimized.
NH 2
N
0
0
CH3
N
N
N
IN
DNA
K
N
DNA
DNA
Cytosine
(C)
Thymine
(T)
0
OH3
CH
N
N
A
N
NH 2
Guanine
(G)
(A)
2
NH
DNA
Adenine
NH
+N
N
3
N
CH 3
NH
NA
N
N
N
N
DNA
1,M-ethenoadenIne
(sA)
N
DNA
N
N
O
N
0
NH 2
7-methylguanine
(7-meG)
DNA
3-methylthymine
(3-meT)
DNA
3, A-ethenocytosine
(CC)
19
Figure 1.2. Base excision repair (BER) in humans. The major BER pathway in humans begins with the
action of a monofunctional DNA glycosylase like AAG (1). After lesion (red circle) removal, the
apurinic/apyrimidinic (AP) site left behind is recognized by AP endonuclease I, which clips the
phosphodiester DNA backbone leaving a 3'-hydroxyl group and a 5'-deoxyribosephosphate (dRP) moiety
(2;(28)). The dual-function DNA polymerase $ attaches a new template base (in blue) to the 3'-OH and
removes the 5'-dRP, leaving a free 5'-phosphate (3;(29)). To complete the repair, the nick left behind is
sealed using DNA ligase I or III (4;(9, 30)).
5
3'
4 4
19 9t
4
96-1
I
-
4
9
3'
4
I
9
I
51
I
1. DNA glycosylase
5'
3'
5'
2. AP Endonuclease I
dRP
5'
OH
3'
3'
3. DNA Polymerase
p
32
tPO
51
oH
4 4
9 9
3'
4
9
3'
5'
T
4. DNA Ligase 1/11t
5'
3'
20
4
4 4 4
9 9 9
4 4
4
3'
5'
Figure 1.3. Introduction to the flavin and nicotinamide cofactors.
a)
b)
nicotinamide adenine dinucleotide (phosphate)
flavin adenine dinucleotide
(FAD)
NAD(P)'
0
flavin mononucleotide
(FMN)
N
NNH
2
p0 1 0
OP.-. 0
0-
riboflavin
(vitamin B2)
NAD(P)H
(oxidized)
H
0
N
(reduced)
NH 2
N
H
H
0
H.)
(2L)
N
NH 2
K)
N
O-P=O
0
0
nicotinamide
HO
"OH
NH 2
Sa
N
4
Isoalloxazine
3
7a
quinone
O
H. jj
(1 e.
N-
N
KN
O-P==O
N0
0-
FAD
(oxidized)
R
OH 0
N
(2e-)
OH OH
0
21
81959a101y~
0
1
"OH
Hi
0-
N
0
O=P-0-
NH
N
NH
H
(1
e)
0
semiquinone
FADH-
R
I
H
NNyO
NH
H
0
hydroquinone
FADH 2
(reduced)
21
Figure 1.4. General mechanism of flavin-dependent monooxygenases.
R
R
'N
N
N
N
NH
N
N
H
0
R
N
o
N
NANHN
02N
HD
:0: 0
:0:
H
CH
I-
:0:
N
N
N
0H
H
A
H
I~
I
N
)aN
0
~~HO ..,
Nu:J
22
Nu:
N
*z
X
I0
N
HOt 0
N
0
NH
I.V
REFERENCES
1.
Moran, U., Phillips, R., and Milo, R. (2010) SnapShot: key numbers in biology., Cell 141,
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Hofseth, L., Khan, M., and Ambrose, M. (2003) The adaptive imbalance in base excision-repair
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Lee, C.-Y. I., Delaney, J. C., Kartalou, M., Lingaraju, G. M., Maor-Shoshani, A., Essigmann, J.
M., and Samson, L. D. (2009) Recognition and Processing of a New Repertoire of DNA
Substrates by Human 3-Methyladenine DNA Glycosylase (AAG), Biochemistry 48, 1850-1861.
O'Brien, P. J. (2006) Catalytic promiscuity and the divergent evolution of DNA repair enzymes,
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O'Brien, P. J., and Ellenberger, T. (2004) Dissecting the broad substrate specificity of human
3-methyladenine-DNA glycosylase, J Biol Chem 279, 9750-9757.
Lingaraju, G. M., Davis, C. A., Setser, J. W., Samson, L. D., and Drennan, C. L. (2011)
Structural basis for the inhibition of human alkyladenine DNA glycosylase (AAG) by
3,N4-ethenocytosine containing DNA, J Biol Chem 286, 13205-13213.
Setser, J. W., Lingaraju, G. M., Davis, C. A., Samson, L. D., and Drennan, C. L. (2012)
Searching for DNA lesions: structural evidence for lower- and higher-affinity DNA binding
conformations of human alkyladenine DNA glycosylase., Biochemistry 51, 382-390.
Massey, V. (2000) The chemical and biological versatility of riboflavin., Biochem Soc Trans 28,
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Blyth, A. (1879) The composition of cow's milk in health and disease., J Chem Soc 35, 530-539.
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lumi-lactoflavins), Ber Deut Chem Gesell 67B, 1460-1462.
Karrer, P., Sch6pp, K., and Benz, F. (1935) Synthese des flavins IV (Synthesis of flavins IV),
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24
Theorell, H. (1935) Reindarstellung der Wirkungsgruppe des gelben Ferments. (Purification of
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yellow enzyme and its effects), Biochem Z 266, 377-411.
Palfey, B. A., and McDonald, C. A. (2010) Control of catalysis in flavin-dependent
monooxygenases., Arch Biochem Biophys 493, 26-36.
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monooxygenases, a diverse class of oxidative biocatalysts., J Biotech 124, 670-689.
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function of p-hydroxybenzoate hydroxylase., Arch Biochem Biophys 433, 297-311.
Dym, 0., and Eisenberg, D. (2001) Sequence-structure analysis of FAD-containing proteins.,
Protein Sci 10, 1712-1728.
Robson, C. N., and Hickson, I. D. (1991) Isolation of cDNA clones encoding a human
apurinic/apyrimidinic endonuclease that corrects DNA repair and mutagenesis defects in E. coli
xth (exonuclease III) mutants, Nucleic Acids Res 19, 5519-5523.
Matsumoto, Y., and Kim, K. (1995) Excision of deoxyribose phosphate residues by DNA
polymerase beta during DNA repair, Science 269, 699-702.
Fan, J., and Wilson, D. M. (2005) Protein-protein interactions and posttranslational modifications
in mammalian base excision repair, Free Radical Biol Med 38, 1121-1138.
25
26
Chapter 2. Structural Basis for the Inhibition of Human Alkyladenine DNA
Glycosylase (AAG) by 3,N-Ethenocytosine-containing DNA
This research was originally published in the Journal of Biological Chemistry. "Gondichatnahalli
M. Lingaraju, C. Ainsley Davis, Jeremy W. Setser, Leona D. Samson, Catherine L. Drennan.
Structural Basis for the Inhibition of Human Alkyladenine DNA Glycosylase (AAG) by
Journal of Biological Chemistry. 2011;
3,N 4-Ethenocytosine-containing
DNA.
286(15):13205-13213" C the American Society for Biochemistry and Molecular Biology.
11.1
SUMMARY
Reactive oxygen and nitrogen species, generated by neutrophils and macrophages in chronically
inflamed tissues, readily damage DNA, producing a variety of potentially genotoxic etheno base
lesions; such inflammation-related DNA damage is now known to contribute to carcinogenesis.
While the human alkyladenine DNA glycosylase (AAG) can specifically bind DNA containing either
1,N 6-ethenoadenine (FA) lesions or 3,N 4-ethenocytosine (EC) lesions, it can only excise eA lesions.
AAG binds very tightly to DNA containing 8.C lesions, forming an abortive protein-DNA complex;
such binding not only shields EC from repair by other enzymes, but also inhibits AAG from acting
on other DNA lesions. To understand the structural basis for inhibition, we have characterized the
binding of AAG to DNA containing EC lesions and have solved a crystal structure of AAG bound to
a DNA duplex containing the EC lesion. This study provides the first structure of a DNA glycosylase
in complex with an inhibitory base lesion that is induced endogenously, that is by exposure to
environmental agents such as vinyl chloride. We identify the primary cause of inhibition as a
failure to activate the nucleotide base as an efficient leaving group, and demonstrate that the higher
binding affinity of AAG for sC versus i A is achieved through formation of an additional hydrogen
bond between Asn169 in the active site pocket and the 02 of 8C. This structure provides the basis
for the design of AAG inhibitors currently being sought as an adjuvant for cancer chemotherapy.
27
II.I
INTRODUCTION
Genotoxic etheno (e)-lesions such as 3,N 4-ethenocytosine (EC) and 1,N-ethenoadenine
(eA) are endogenously generated when DNA is attacked by reactive aldehydes. These reactive
compounds are generated as by-products of lipid peroxidation that is induced by reactive oxygen
and nitrogen species (RONS). Neutrophils and macrophages generate large quantities of RONS
in tissues undergoing chronic inflammation (1,
2), and it is widely accepted that such
inflammation increases the risk of colon cancer in ulcerative colitis (UC) and Crohn's disease
patients, and increases the risk of liver cancer in Wilson's disease and Hemochromatosis patients
(1,
3). In fact, increased levels of F-lesions in the DNA of tissues undergoing chronic
inflammation have been reported for each of these diseases (4). Depending on the type of DNA
polymerase, EC mispairs with A, T or C during DNA replication, resulting in both transition and
transversion mutations (5). In contrast, EA primarily gives rise to A:T to T:A transversion
mutations (6). These mutagenic E-lesions are generally removed via the base excision repair
(BER) pathway, initiated by lesion-specific DNA glycosylases that cleave the N-glycosidic bond
between the damaged base and the deoxyribose sugar (5, 7). In humans several DNA
glycosylases can excise eC, namely thymine DNA glycosylase, methyl-CpG binding domain
protein and single strand monofunctional uracil DNA glycosylase (5). In contrast, there is only
one DNA glycosylase known to excise EA lesions in humans, alkyladenine DNA glycosylase
(AAG) (5, 8).
AAG (also known as MPG and ANPG) has been previously characterized by
crystallography. The crystal structures of an N-terminally truncated, but catalytically active,
construct of AAG (A79AAG), both bound to a pyrrolidine abasic site mimic (pyr) and bound to
an EA containing piece of DNA, suggest a mode by which AAG recognizes a wide range of
lesions, while still discriminating against undamaged bases (9, 10). When substrate is bound,
AAG can excise the damaged base through acid-base catalysis (11). A putative catalytic water
molecule, revealed by crystallographic studies, is proposed to act as a nucleophile, as it is ideally
positioned to attack the N-glycosidic bond present between the EA base and the deoxyribose
sugar (10). This water molecule is also in contact with the side chain of Glul25, which is
proposed to be the catalytic base responsible for activating the water for nucleophilic attack.
Consistent with this proposal, introduction of a E125Q mutation completely abolishes AAG
activity (9-11). The identity of the general acid is unknown. Interestingly, in the structure of
28
A79AAG bound to FA:T containing DNA, the FA lesion remained intact (10). Later experiments
showed that the catalytic activity of A79AAG is significantly inhibited in the presence of a
variety of divalent metal ions including Mn 2 *, Zn 2 *, Ca2+, Cd 2 *, Ni2+and most importantly, Mg 2 +,
which was contained in the crystallization buffer (9, 10, 12, 13). However, the structures did not
show any Mg 2+ ions bound (9, 10).
In addition to repairing cA, AAG can repair other RONS, and alkylation induced DNA
damage,
including
lesions
hypoxanthine
(Hx),
1,N 2-ethenoguanine,
8-oxoguanine,
3-methyladenine (3-meA), 7-methylguanine (7-meG) and 3-methylguanine (2, 8). However, in
spite of this broad substrate specificity, there is a growing list of lesions to which AAG can bind
while failing to excise the lesion. In addition to FC containing DNA (14), this list now includes
3-methyluracil, 3-ethyluracil, 3-methylthymine and 3-methylcytosine (15). This binding without
cleavage can result in the formation of stable abortive complexes between AAG and damaged
DNA. The EC-AAG abortive complex has been shown to inhibit AAG glycosylase activity in
vivo in human cells, and to result in replication blockage (14), increasing the genotoxicity of cC
lesions in vivo. In tissues undergoing chronic inflammation, which have higher cC content (4),
the formation of AAG-EC abortive complexes may significantly diminish the repair of other
AAG substrates, ultimately resulting in the accumulation of various DNA lesions in addition to
cC. Interestingly, in ulcerative colitis patients, the colon epithelium undergoing chronic
inflammation was found to have increased AAG expression, perhaps indicating an adaptive
response triggered by increased levels of DNA damage (16). This adaptation might compensate
for the AAG hijacked by cC lesions.
In order to understand the structural basis for the inhibition of AAG by EC containing
DNA, we have carried out biochemical and crystallographic studies on AAG, using a truncated
form of the protein (A79) that was previously shown to have full catalytic activity (11, 15, 17).
Our crystal structure of A79AAG bound to a DNA duplex containing cC:G (G paired opposite
EC), in combination with previous (11) and current (this work) biochemical analysis, suggests
that the failure of AAG to activate the leaving group (cC) by protonation is likely the primary
reason for its inability to remove cC from the DNA. This structure also shows that a divalent
metal ion, Mn2 , can bind to the base opposite the cC lesion, changing its sugar pucker, and
providing the first structural framework for considering the molecular basis for metal ion
inhibition of AAG.
29
11.111 RESULTS
AAG binding studies
Th, binding affinity of A79AAG to the EA:T (EA paired opposite T) lesion containing
25-mer and 13-mer duplexes was compared with the binding affinity to cC:G (EC paired opposite
G) duplexes, using gel mobility shift assays (Figure 11.1, A and B; and Table 11.1). As shown in
Table II.1, when either FA or EC lesions are present in a given sequence context, A79AAG
consistently binds the FC:G duplex with ~2-fold higher affinity compared to that of the EA:T
duplex. In addition, A79AAG binds the
EA:T
25-mer duplex (Kd = 20 ± 2 nM) with ~2-fold
higher affinity, compared to the EA:T 13-mer duplex used for crystallization (Kd = 46 ± 6 nM).
Correspondingly, A79AAG also binds the eC:G 25-mer duplex (Kd
=
13 ± 2 nM) with -2-fold
higher affinity, compared to the eC:G 13-mer duplex (Kd = 21 ± 3 nM). These results indicate
that the binding affinity of A79AAG to DNA containing the same lesion varies depending on the
length of the DNA duplex. The binding studies also show that in a given sequence context,
A79AAG binds EC:G duplex with higher affinity compared to that of the eA:T duplex.
Catalytic ability of AAG for cC containing DNA
Following our binding studies, we tested the DNA glycosylase activity of both full-length
and A79AAG on EA and PC residues present in the 25-mer oligonucleotide duplexes (Figure
II.lE). As shown in the representative gel, both full-length and A79AAG robustly removed EA
from the EA:T 25-mer duplex. However, the activity of both full-length and A79AAG on
PC
residues from the same duplex was completely absent. In contrast, the positive control, E. coli
Mismatch Uracil DNA Glycosylase (MUG) (Trevigen) shows robust catalytic activity on PC
contained in an cC:G 25-mer duplex (Figure II.lE). Further, we tested the activity of A79AAG
on PC containing 13-mer oligonucleotide duplexes used for crystallization, with cC paired
opposite different bases. The results show A79AAG does not have activity on cC:G, cC:A,
EC:C, or cC:T 13-mer duplexes (Figure 11.2).
Inhibition of AAG by eC containingDNA
We employed competition assays to measure the inhibition of A79AAG activity on cA:T
25-mers using cA:T and
EC:G
13-mer competitor DNA oligonucleotides (Figure II.1, C and D;
and Table II.1). The catalytic activity of A79AAG on labeled EA:T 25-mer duplex was measured
30
at 37*C in the presence of an increasing concentration of the aforementioned cold competitors
(Figure II.1C). The cleavage data were fitted to equation (2) (Materials and Methods) in order to
calculate the 50% inhibitory concentration (IC 50 ) for each cold competitor used in the experiment
(Figure II.1D). The results obtained correlate with the binding measurements described above.
As shown in Table 11.1, the EA:T 13-mer DNA duplex binds approximately 4-fold weaker (IC 50
= 163 nM) than the cC:G 13-mer (IC 50 = 39 nM).
Overall structure of the A 79AAG-EC DNA inhibitor complex
The crystal structure of A79AAG-E-C:G at 2.2
A resolution was determined
by molecular
replacement using the structure of A79AAG-pyr:T complex as a search model (9). Difference
electron density maps calculated in the absence of DNA show interpretable electron density for
DNA backbone and for the EC base in the active site pocket of AAG. In comparison to the
previously published A79AAG-DNA complexes (9, 10), the A79AAG-eC inhibitor complex
crystallized in a different space group (P1). The final model of A79AAG-EC:G has been refined
to the R factor of 23.9
(Ree =
28.4) (Table 11.2).
The overall structure of A79AAG bound to an EC DNA lesion includes the insertion of
Tyrl62 into the DNA and the flip of the FC nucleotide into the enzyme active site (Figure 11.3).
The two copies of this inhibitor-protein complex in the asymmetric unit are quite similar to each
other, with a root mean square deviation (RMSD) for all alpha carbon atoms of approximately
0.6
A,
and to that of the A79AAG-cA:T substrate complex (10), also having a RMSD of
approximately 0.6
A. With the
exception of some loops and two disordered regions (Figure II.4),
the only notable difference in our inhibitor structure is that the octahedrally coordinated Na'
metal ion modeled in the structure of the A79AAG-eA:T complex was found to be absent.
Instead, this site is occupied by the N-terminal amino group of residue 80 of a symmetrically
equivalent A79AAG molecule (Figure 11.5). Similar to the Na' ion, the N-terminal amine in this
position also interacts with the main chain carbonyls of Metl49, Gly174, Ala177, Serl7l and the
side chain of Ser172.
Protein-DNA interactions
In both molecules in the asymmetric unit of the A79AAG-C:G structure, the DNA
duplex is largely B-form and is bent away from the protein by about 220, with the bend primarily
31
centered on the flipped EC nucleotide (Figure II.3A). For the most part, all protein-DNA
interactions are similar between the A79AAG-EC:G inhibitor complex and the A79AAG-FA:T
substrate complex (Figures 11.3 and 11.4). Tyr162 makes the most important protein contact,
given that it is inserted into the DNA duplex, replacing the EC lesion, and forming Van der
Waals contacts with the opposite base, G19 (Figure 11.3). A potential steric clash of Tyr162 with
G19 is prevented by shifting this opposite base out of the minor groove, leaving it without a base
pairing partner (Figure 11.3, B and C). This sliding rearrangement, in which G19 is orphaned in
terms of hydrogen bonding, is disruptive and presumably has an energetic penalty associated
with it. The deoxyribose of the orphaned base does make favorable Van der Waals interactions
with Metl64 (Figures II.3B and II.6C), counteracting some of these energetic costs. As was the
case in the A79AAG-EA:T substrate complex, no direct hydrogen bonding interactions are
present between the protein and the base opposite the DNA lesion, indicating that specific
recognition of this base is not required for the binding of AAG to DNA.
Metal ion Mn2" in the z79AAG-EC:G structure
The A79AAG-FC:G complex was crystallized in the presence of 200 mM MnCl 2 , a
condition under which AAG's ability to excise EA from a DNA duplex is impaired (12, 13). We
find electron density consistent with a metal ion near the base opposite the EC lesion (Figures
II.3C and 11.6), at a distance of approximately 16
A to the
AAG active site (Cl' of EC). Mn 2 ,
refines well in this electron density with no positive or negative difference electron density. In
contrast, refinement of a water molecule or a sodium ion (also present in the crystallization
buffer) leads to positive difference electron density, suggesting that the correct atom in this site is
heavier than water and sodium, consistent with Mn 2 ,. Anomalous difference density is also
present at both sites in the asymmetric unit at approximate sigma levels of 8 and 5 for chain A
and chain B respectively, consistent with the presence of Mn 2 1 ions (Figure II.6A). At the
wavelength of data collection (k = 1.116
A),
sodium would not give rise to an anomalous signal,
ruling out the other metal ion contained in the crystallization buffer as being present in this site.
The refined Mn 2 , is coordinated to the 06 of G19 (the base opposite SC), to the N7 of A 18, and
to three water molecules (Figure II.3C). The coordination of Mn 2' appears to induce a significant
change in the phosphodiester backbone in the region around G19, resulting in a change in the
sugar pucker to a C2'-exo conformation (Figure II.6B).
32
Active site architectureof z179AAG-EC DNA complex
The PC base lesion is recognized and stabilized by hydrogen bonds, along with Van der
Waals interactions (Figures 11.7 and 11.8). Similar to the recognition of FA (Figure II.7B), eC is
stacked between Tyr127 on one side, and His136 and Tyr159 on the other side (Figure II.7A).
Tyr159 makes edge-to-face contact with the lesion base. The specificity to discriminate between
the EC lesion and undamaged cytosine appears to be achieved through a hydrogen bond donated
from the main chain amide of His136 to the N' of the PC base. This interaction is similar to the
damage-specific recognition of -A (versus undamaged adenine) by AAG, which is made through
a hydrogen bond donated by the main chain amide of His136 to the N6 of EA. An additional
hydrogen bond is also observed between the carboxamide nitrogen of the side chain of Asn169
and the 02 of EC. Mutants of Asn169, namely Asnl69Leu and Asnl69Ala, show respective
~2-fold and ~4-fold reduced affinity for the EC:G 25-mer oligonucleotides, compared to that of
wild type A79AAG (Table 11.1), indicating that the additional hydrogen bond donated from
Asn169 to
FC
contributes to the increased affinity of A79AAG for EC inhibitor DNA.
Interestingly, in the A79AAG-FC:G inhibitor complex, the putative catalytic water molecule
(proposed to act as a nucleophile) occupies the same position as in the A79AAG-eA:T substrate
complex (Figure 11.7). Also in agreement with the structure of the A79AAG-EA:T substrate
complex, this water molecule interacts with Glul25, Arg 182 and main chain carbonyl oxygen of
Val262. Glul25, which is proposed to activate the water molecule for nucleophilic attack (10,
11), is held in its position through a hydrogen bond donated from Tyr127 to its carboxyl group.
The side chain of Arg 182 further contacts the 3'-phosphate of the EC nucleotide.
In comparison to the recognition of the EA substrate, the EC inhibitor recognition induces
slight changes in the active site, mostly with respect to DNA backbone positions (Figure II.7C).
Since FC is smaller than EA, the DNA backbone must be pulled farther into the active site such
that EC is recognized with optimal molecular interactions. In a modeling exercise in which the
DNA backbone is held rigid, AAG fails to form optimal hydrogen bonding interactions between
both the main chain amide of His 136 and the N4 of FC, and the side chain of Asn 169 and the 02
of EC. Perhaps as a result of this slight repositioning of the DNA backbone, the side chain of
Arg 182 adopts a different conformation (Figure II.7C). The side chain of Arg 182 still interacts
with the 3' phosphate of the EC nucleotide and still hydrogen bonds to the putative catalytic
water molecule. Overall, however, comparison of the active site of the A79AAG-eC inhibitor
33
complex with that of the A79AAG-EA:T substrate complex shows that, with the exception of
Arg182, all the residues involved in lesion recognition and catalysis maintain similar
orientations.
II.IV DISCUSSION
AAG plays an important role in the maintenance of genomic integrity, presumably
through its ability to recognize, bind and excise a wide-range of DNA base lesions. It was
therefore surprising that AAG also has the ability to recognize and bind a number of DNA base
lesions that it is incapable of excising, in particular the -C lesion. Moreover, the tight binding of
AAG to EC leads to the inhibition of its catalytic activity and, in addition, is known to shield EC
from ABH2-mediated direct reversal repair (18). In order to understand the structural basis for
the inhibition of AAG by eC containing DNA, we solved the crystal structure of A79AAG bound
to a 13-mer EC:G (EC paired opposite G) duplex.
Given that AAG can bind EC containing DNA, we anticipated that the lack of activity
may be a result of one or more of the following factors: (i) AAG might fail to flip EC into its
active site, or it might flip EC into an alternative binding pocket that lacks the appropriate
catalytic residues; (ii) the binding mode of EC in the active site might not favor accommodation
of the water molecule thought to act as a nucleophile in the reaction; (iii) the side chain of the
putative catalytic base (Glul25) might adopt a nonproductive conformation that fails to activate
the putative catalytic water molecule; (iv) AAG might be unable to protonate EC, failing to
activate it for departure.
The crystal structure shows that AAG successfully flips the EC inhibitor into the same
active site pocket that binds the EA substrate, ruling out the first possibility. We also find that the
putative catalytic water molecule is present in the inhibitor complex, ruling out the second
possibility. Furthermore, as was observed in the structure of the A79AAG-EA:T substrate
complex, this water molecule is in contact with Glul25, as would be required for its activation.
The water molecules are ideally positioned to attack the N-glycosidic bond. Thus, we infer that
AAG's inability to remove EC is unlikely to be due to a problem with nucleophilic activation or
attack, ruling out the third possibility from our list.
We next examined the fourth possibility, that the failure to excise EC is due to a problem
with leaving group activation. In a previous biochemical study, O'Brien and Ellenberger
34
measured the pH-rate profiles for AAG's excision of neutral Hx and CA lesions, and its excision
of the positively charged 7-meG lesion under single turnover conditions (11). They found that
the pH-rate profiles for FA and Hx excision follow a bell-shaped curve, indicating that for the
excision of neutral lesions, AAG uses the action of both a general acid and a general base
(Glul25). The general base can activate a catalytic water molecule, while the general acid is
expected to facilitate the protonation of neutral lesions, making the lesion base a better leaving
group (11). In contrast, the pH-rate profile for the excision of 7-meG shows only a single
ionization corresponding to a general base, suggesting that leaving group activation of 7-meG is
not necessary because the base is already positively charged. To help pinpoint the site of
protonation, the activity of AAG on Hx was compared to its activity on 7-deaza-Hx, and
although AAG greatly enhances the rate of Hx excision (~10'), the same lesion with N7 changed
to C7 is not cleaved by AAG, directly implicating the involvement of the N7 position in catalysis
(See Figure II.9B for numbering) (11). While this study was unable to identify a specific residue
as the general acid, the crystal structure of a A79AAG(E125Q)-EA:T substrate complex shows a
water molecule in contact with the equivalent position to N7 of Hx, that is N7 of EA (Figure
II.9A) (10), raising the possibility that a protein-bound water molecule could be responsible for
protonation. Once protonated, the AAG active site is designed to stabilize the protonated form of
the base through a hydrogen bond between N7H of FA and the backbone carbonyl oxygen of
Ala134 (Figure II.9A). Given these findings on the catalytic significance of protonation of the
N7 of Hx and EA, it is important to consider the equivalent position in the EC base. A
superposition shows that unlike FA, cC has a carbon (C5) in the position equivalent to N7, and
thus cannot be protonated at that site (Figure II.9A). Therefore, as opposed to our findings with
respect to possibilities one through three, it appears that AAG's failure to cleave eC could be due
to an inability to activate the EC leaving group by protonation. Since AAG is reported to bind
and not cleave a number of different pyrimidine lesions, including 3-methyluracil, 3-ethyluracil,
and 3-methylthymine (15), this mechanism of inhibition may be broadly applicable.
Given that AAG cannot repair EC lesions, it is interesting that AAG binds this lesion so
tightly. The molecular basis for the approximate 2-fold higher affinity of AAG for the EC:G
duplex (compared to the substrate cA:T duplex) can be attributed to an additional hydrogen bond
formed between the carboxamide side chain of Asn 169 and the 02 of EC. Mutation of Asn169 to
residues that cannot maintain this hydrogen bond (Leu and Ala) completely abolished this 2-fold
35
binding effect (Table II.1), suggesting that this one hydrogen bond is chiefly responsible for the
higher affinity of FC containing DNA. Thus, in addition to its previously proposed role of
serving to help discriminate between damaged and undamaged guanine (19), Asn169 appears to
play a role the recognition and binding of pyrimidine DNA lesions. It is important to note that
Asn169 is strictly conserved among AAG-related glycosylases (9).
Although inhibition of AAG by divalent metal ions (Mg 2 l, Mn 2 +, Zn 2 *, Ca 2 *, Cd 2 + and
Ni 2 ) has been well-documented (12, 13), no previous crystal structure displayed electron density
consistent with such an ion, even though these cations were used in the crystallization buffers (9,
10). Here we find density consistent with the presence of Mn 2 , in position to coordinate the base
opposite the EC lesion (G19) (Figures II.3C and 11.6). Binding of Mn 2 . to this site appears to
influence the pucker of the sugar yielding a C2'-exo configuration (Figure II.6B). This
occurrence is the first time that this sugar pucker has been observed in an AAG structure. In our
Mn 2 +containing structure, the binding of -C to the active site is nearly identical to that observed
in the A79AAG-EA:T structure. Thus, the inhibitory effect of Mn 2 +does not appear to be due to a
large conformational change in the active site, although the electrostatics of the active site could
be affected by the presence of the positively charged ion approximately 16 A away. Other
dynamic movements of protein or DNA that are important in catalysis might also be affected by
the Mn 2. coordination. While binding of divalent metal cations to protein and DNA is common,
it is intriguing that we find this divalent metal bound to such an important site in this
protein-DNA complex. We now have a physical model for the influence of divalent cations on
AAG activity that can be tested.
With a molecular view of the AAG-FC abortive complex in hand, it is interesting to
consider what the physiological benefits in forming this complex might be. Abortive complexes
between alkyltransferase-like (ATL) proteins and a lesion that it cannot excise, 0 6-alkylguanine,
have recently been observed (20). ATLs are known to interact with proteins in another DNA
repair pathway, nucleotide excision repair (NER), suggesting that ATLs may function to present
alkylated DNA to NER proteins for repair (20, 21). Given the difficultly of finding a single
damaged DNA base in the midst of the genome, it makes sense that once found, some DNA
repair proteins may be designed to 'hand-over' lesion bases that they cannot themselves repair to
an alternative repair pathway. While it is too early to infer directly from ATLs, preliminary data
do show that AAG can interact with human NER proteins hHR23A and hHR23B (22).
36
Obviously, more studies are necessary to establish if AAG--C abortive complexes interact with
NER proteins, resulting in EC repair, but this idea is intriguing and, given the recent studies on
ATL, not without precedence. Originally identified by its ability to excise alkylated 3-meA and
7-meG lesions (8, 23), AAG's role in DNA repair is far more complex than once thought. Given
the importance of repairing RONS generated DNA damage for tissues undergoing chronic
inflammation, a complete understanding of AAG's physiological function is essential.
ILV
MATERIALS AND METHODS
A 79AAG plasmidconstruction, creation of mutants, and protein preparation
Constructs of full-length (FL) and a truncated form of AAG (with 84 residues at the Nterminus deleted) were cloned into pET19b-PPS vectors for protein expression (UniprotID:
P29372). These constructs encoded the wild-type AAG sequence with an N-terminal loX
histidine tag followed by the precision-protease cleavage site (PPS). The precision protease
treatment to remove the histidine tag leaves behind four extra amino acids (GPHM) from the
expression vector at the N-terminus of AAG. For FL-AAG, this cleavage site results in a GPHM
sequence prior to residue 1, and for the truncated form of AAG, this cleavage site results in the
addition of residues Gly8O, Pro81, His82 and Met83 prior to residue 84. We refer to this
truncated protein construct as A79AAG, although the actual AAG protein sequence starts at
Thr84. It should be noted that the AAG protein from previous structural studies was also referred
to as A79AAG (9, 10). However, in those former studies, all residues contained in the construct
are
of the
wild type
sequence.
For the
creation
of A79AAG-Asnl69Leu
and
A79AAG-Asn169Ala mutants, PCR based site directed mutagenesis was performed with the
primers shown in Table 11.3 and successful incorporation was confirmed by DNA sequencing
(MWG Biotech). A79AAG protein expression and purification was done similarly to previously
described protocols (11, 24) and is outlined below.
AAG protein expression andpurification
Both full-length and A79AAG and its mutants (Asnl69Leu, Asnl69Ala) were expressed
and purified using a similar protocol. Each pET19b-PPS AAG plasmid construct was
transformed into BL21(DE3) cells by heat shock and plated onto LB (Luria Bertani) agar plates,
supplemented with 100 [tg/mL of ampicillin and incubated at 37'C overnight. A single colony
37
was used to inoculate 250 mL of LB broth supplied with 100 tg/mL of ampicillin and grown at
37*C overnight. This starter culture was used to inoculate 6 L of LB broth supplied with 100
Rg/mL of ampicillin. The culture was grown at 37*C, until the cells reached an optical density at
600 nm (OD600 ) equal to 1.0. The cultures were cooled to room temperature, and protein
expression was induced with 0.5 mM IPTG for 6-8 hr. The cells were centrifuged at 4*C and
4000 rpm for 30 min and resuspended in 150 mL of Buffer A (20 mM Potassium Phosphate pH
7.0, 500 mM NaCl, 10% v/v glycerol, 1 mM DTT) supplied with 4 tablets of EDTA-free
protease inhibitor mixture and frozen at -80'C. The cells were thawed and lysed by sonication.
The cell lysate was centrifuged at 18,000 rpm for 45 min and the supernatant was loaded onto a
HisPrepT M FF 16/10 Ni-affinity column (Amersham Biosciences), which was pre-equilibrated
with 10 column volumes (CV) of Buffer A. The first wash was done with 10 CV of Buffer A,
followed by a wash with 10 CV of Buffer A containing 40 mM imidazole. The protein was
eluted by creating a 10 CV imidazole gradient from Buffer A and Buffer B (Buffer A supplied
with 500 mM imidazole). Upon elution, the N-terminal 1oX histidine tag was cleaved from the
protein by precision protease (GE Healthcare Biosciences) treatment at 16*C for 12-14 hr. This
sample was diluted to the final NaCl concentration of 100 mM and loaded directly onto a
HiTrapT M SP FF ion-exchange column (Amersham Biosciences). The unbound protein was
eluted using 10 CV of Buffer C (20 mM HEPES pH 7.5, 100 mM NaCl, 10% glycerol, 5 mM
DTT). The bound protein was eluted using a gradient between Buffer C and Buffer D (Buffer C
supplemented with IM NaCl), and concentrated by centrifugation at 3500 rpm at 4*C using
Amicon 10-KDCO (kilodalton cut off) ultrafilters. Further purification was done by gel filtration
using Buffer E (20 mM HEPES pH 7.5, 100 mM NaCl, 10% glycerol and 5 mM DTT) and a
Superdex"
75 gel filtration column (Amersham Biosciences). The final purified protein in
Buffer E was almost 99% pure as evidenced by the SDS-PAGE analysis. The purified protein
was concentrated using Amicon 10-KDCO ultrafilters and the amount of protein was estimated
using the extinction coefficient method by UV absorption at 280 nm.
Preparationof oligonucleotides and 32P-labeling
DNA oligonucleotide substrates (Integrated DNA Technologies) were dissolved in TE
buffer (10 mM Tris-HCl pH 8.0 and 1 mM EDTA) and quantified by the extinction coefficient
method using UV absorption at 260 nm. For the DNA glycosylase and binding studies, the
38
lesion-containing strand was labeled on the 5' end with
1
2
P-yATP (Perkin Elmer) using
polynucleotide kinase (PNK) (New England Biolabs) at 37*C for 30 min, followed by heat
inactivation of PNK at 70'C for 15 min. Duplex oligonucleotides were created by annealing the
32 P-labeled strand
with its complementary strand. The unincorporated 3 2 P-yATP was separated
from
32 P-labeled
oligonucleotides
using
Sephadex
G-25
quick
spin
columns
(Amersham-Pharmacia).
Gel mobility shift assays
The binding affinity of A79AAG to different DNA oligonucleotides was measured using
gel mobility shift assays. DNA oligonucleotides were
32 P-labeled
and purified using Sephadex
G-25 quick spin columns (as outlined above). Binding reactions were set up as solutions
containing 1X binding buffer (50 mM HEPES-NaOH pH 7.5, 100 mM NaCl, 1 mM EDTA,
9.5% v/v glycerol, 50 [tg/mL BSA and 5 mM DTT), 2 nM
32 P-labeled
oligonucleotide and
0-1000 nM of the purified A79AAG protein. The reaction samples were incubated on ice for 30
min and the products were resolved using 6% native-PAGE in 0.5 X TBE buffer at 110 V for 3
hr at 4'C. The extent of complex formation was quantified and analyzed by phosphorimaging.
Dissociation constant
(Kd)
values were calculated by fitting the binding data to the binding
equation (equation 1) using GraphPad Prism (GraphPad software, Inc.).
Y =Bmax*X/(Kd+X)
(1)
where Y is the specific binding, Bmax is the maximal binding, X is the concentration of protein
and Kd is the dissociation constant. Each experiment was repeated at least three times and the
data represent the average of at least three independent experiments.
DNA glycosylase assays
DNA glycosylase assays were set up as solutions containing IX glycosylase assay buffer
(50 mM Tris-HCl pH 7.8, 100 mM NaCl, 1 mM EDTA, 50 gg/mL BSA and 5 mM DTT), 2 nM
32 P-labeled
oligonucleotide and 25 nM of either the purified full-length or A79AAG enzyme.
The reactions were carried out at 37*C. Aliquots (10 [tL) from particular time points were mixed
with piperidine to the final concentration of 0.2 M and heated at 75*C for 15 min. The piperidine
39
treatment cleaves all abasic (AP) sites resulting in single strand breaks at the region of AP-sites.
This procedure was followed by the addition of one sample volume of 90% formamide buffer
with dye markers. The samples were heated at 75*C for 15 min and the products were resolved
using 20% denaturing Urea-PAGE in IX TBE buffer at 450 V for 2 hr. The extent of substrate
cleavage was quantified and analyzed by phosphorimaging.
Competition DNA glycosylase assays
Competition DNA glycosylase assays were performed to measure the inhibition of
A79AAG activity on aA containing duplex DNA substrate by EA and PC duplexes. The reactions
were set up as solutions containing IX glycosylase assay buffer, 1 nM
32P-labeled
EA:T (T paired
opposite EA) 25-mer oligonucleotide duplex DNA (5'-GCA ATC TAG CCA EAGT CGA TGT
ATG C-3'), 5 nM of the purified A79AAG enzyme and increasing concentrations of competitor
DNA (0-3000 nM). The reactions were carried out at 37'C for 30 min. After incubation, NaOH
was added to a final concentration of 0.2 M, followed by heating at 75*C for 15 min. Similar to
piperidine treatment, hot alkali treatment with NaOH cleaves all AP sites and creates DNA
single strand breaks at the AP sites. Upon cooling, one sample volume of 90% formamide buffer
with dye markers was added into the reaction mixture. The samples were heated at 75 0 C for 15
min before loading, and the products were resolved using 20% denaturing Urea-PAGE in IX
TBE buffer at 450 V for 2 hr. The extent of substrate cleavage was quantified and analyzed by
phosphorimaging. The experiment with each competitor was repeated at least three times. In
order to calculate the IC 50 (50% inhibitory concentration), the competition data were fitted to the
sigmoidal dose response curve (equation 2) using GraphPad Prism.
Y= Ymin + (Ymax -Ymin) / (1 +
1 0 LogIC50-X)
(2)
where X is the logarithm of competitor concentration, Ymax and Ymin are the maximum and
minimum values of % AAG activity (Y) and LogIC50 is the logarithm of IC50 .
Crystallization of the A79AAG-EC:G complex
The eC:G DNA duplex was prepared by annealing the EC containing 13-mer
crystallization oligonucleotide (5'-GAC ATG
40
ECTT
GCC T-3') with its complementary strand
that contained G opposite PC (5'-GGC AAG CAT GTC A-3'). The A79AAG-EC complexes were
prepared by mixing equimolar ratios of A79AAG and FC:G 13-mer DNA duplex at the final
protein-DNA complex concentration of 0.3 mM in the complex buffer (20 mM HEPES-NaOH
pH 7.5, 100 mM NaCl, 0.1 mM EDTA, 5% v/v glycerol and 1 mM DTT). The complex was
incubated on ice for 15 min and used for crystallization. The crystals were obtained by the
hanging drop vapor diffusion method, upon mixing 1 pL of complex and 1 pL of the reservoir
solution (100 mM sodium cacodylate pH 6.0, 200 mM manganese chloride and 20%
polyethylene glycol (PEG)-3350) over 0.5 mL of the reservoir solution, followed by incubation
for 2 days at 22*C. The crystals formed as plates, which were mounted directly from the hanging
drop to the center of loops larger than the crystal size and flash frozen in liquid nitrogen.
Data collection and structure determination
The X-ray diffraction data for the A79AAG-EC:G complex were collected at the
Advanced Light Source (Berkeley, CA) on beamline 12.3.1 at lOOK to 2.2 A resolution. These
data were processed using Denzo/Scalepack (25) and the data statistics are given in Table 11.2.
The structure of the A79AAG-EC:G, with 2 molecules in the asymmetric unit, was determined
by molecular replacement using PHASER (26) and the AAG protein coordinates from the
A79AAG-pyr:T complex structure (PDB ID 1BNK (9)) as a search model. A 2mFOI-DIFe!
electron density map (all maps having coefficients from 4ligma weighting) contoured at 1R, and
a mIFI-DIFel electron density map contoured at 3o, calculated in the absence of DNA, showed an
interpretable electron density map for the DNA backbone. Another round of refinement was
performed upon partial fitting of the DNA (7 nucleotide pairs starting at 5'- 4ATG(pyr)TTG10 3' and the complementary strand 5'- 16CAAGCAT22 -3'). The resulting calculated 2mlF.I-DIFcl
(1 Y) and mIFOI-DIFel (3y) electron density maps showed interpretable electron density for the EC
base in the active site pocket of AAG, as well as continuous electron density for building the
remaining DNA backbone. Upon fitting the missing DNA portions into the electron density,
initial models were subjected to restrained refinements using Refmac 5.4 (27, 28). Topology and
parameter files for the FC lesion were generated using XPLO2D (29). Initial refinement included
simulated annealing in CNS (30). Iterative rounds of positional and B-factor refinement of the
A79AAG-eC complexes were performed with the guidance of calculated 2mFO1-DIFeI,
mIF!-DlFel electron density maps (generated by Refmac 5.4), and 2mFOI-DIFcI composite
41
omit-maps (generated by CNS), using the model building program Coot (31). Anomalous
difference maps were calculated in CNS (30) using the native data (X = 1.116
A) and the phases
from the final A79AAG-EC:G model. Additional rounds of refinement using TLS parameters
and non-crystallographic restraints were very effective in improving the quality of the fit. At
each stage, the progress of model building was judged by following the change in R factors. The
final model of A79AAG-C:G complex converged at an R factor of 23.9 (Rfe = 28.4) (Table
11.2). The final model was evaluated using PROCHECK (32) and Rampage (33). For residues
80-298 of the A79AAG-EC:G structure, the following have no electron density and are therefore
not included in the model: the residues 203-207, 265-268 and 295-298 in chain A; the residues
205-206, 265-266 and 294-298 in chain B. Due to a lack of interpretable electron density for the
side chains of some residues in the structure of A79AAG-EC:G (Arg201, Leu249 and Glu253 in
chain A), these residues were modeled as alanines. Each protein molecule in the asymmetric unit
has a DNA 13-mer associated with it. One nucleotide of each of these duplexes is disordered
(A26).
II.VI ACKNOWLEDGEMENTS
This work was supported by the US National Institutes of Health (grants P30-ES002109,
GM65337, GM65337-03S2, ES002109, CA055042 and CA092584). C.L.D is a Howard Hughes
Medical Institute Investigator. L.D.S. is an American Cancer Society Research Professor. J.W.S.
is supported by a Repligen KIICR Graduate Fellowship. The Advanced Light Source is
supported by the Director, Office of Science, Office of Basic Energy Sciences, of the U.S.
Department of Energy under Contract No. DE-AC02-05CH 11231. The atomic coordinates and
structure factors (code (3QI5)) have been deposited in the Protein Data Bank (www.rcsb.org).
Author contributions: G.M.L. performed the DNA binding, activity, and inhibition studies.
G.M.L. and L.D.S. designed the experiments. G.M.L., C.A.D., J.W.S., and C.L.D. carried out the
crystallography experiments and the structural analysis. G.M.L., J.W.S., L.D.S., and C.L.D.
wrote the manuscript.
42
TABLES & FIGURES
Table 11.1. Dissociation constant (Kd) values measured using gel shift assays; and 50% inhibitory
concentration (IC 50 ) for the inhibition of A79AAG activity on gA:T 25-mer, measured using competition
DNA glycosylase assay at 37'C, in the presence of increasing concentration of cold competitor 13-mer
duplexes.
IC501 (95% confidence interval (nM)
A79AAG
Oligonucleotide
Wild type
sA:T 25-mer
20 ± 2
Wild type
eC:G 25-mer
13 ± 2
Wild type
EA:T 13-mer
46 ± 6
163 (152-174)
Wild type
eC:G 13-mer
21 ± 3
39 (38-41)
Asnl69Leu
EC:G 25-mer
31± 4
AsnI69Ala
eC:G 25-mer
47 ± 6
Kd' ±
SD 2 (nM)
1: Average of at least three independent experiments.
2: Standard Deviation
43
Table 11.2. Data collection and refinement statistics of the A79AAG-DNA complex.
A79AAG- C:G
Space group
P1
Cell constants
a= 41.23
A,
b= 41.22
A, c= 82.14
A
a= 81.230, P= 88.4', y= 89.15*
12.3.1
Beamline
Wavelength (A)
1.116
Resolution (A)
41
-
2.20
No. total observations
71943
No. unique observations
26278
Completeness (%)
c
96.6 (95.2)
<I/I(1)>
17.1 (4.9)
Rsym (%) a,c
5.8 (19.2)
Model refinement
23.9
Rwork (%) b
Rfree (%)
28.4
b
B-factors (A )
Protein
DNA
Water
Mn 2 , ion
2
15.2
17.9
14.1
chain A: 27.9 chain B: 44.0
RMSD bonds (A) d
RMSD angles (0) d
Number of atoms
Protein
0.007
1.1
3257 (2 mol./asu)
DNA
Water
Mn 2 , ion
1024
232
2
Ramachandran plot (%)
Most favored
89.7
Additionally allowed
10.0
Generously allowed
0.3
1
a: Rsym= IIhkI - <IhkI>I Y IIkl,
where I is the intensity of a reflection hkl and <I> is the average over
symmetry-related reflections of hkl.
b: Rork = YIF 0 - FeI/YIFOI in which F and Fe are the observed and calculated
0
structure factor amplitudes,
respectively. Rfre is calculated from 5% of the reflections not used in the model refinement.
C: Values in parenthesis correspond
to the highest resolution shell.
d: RMSD, root mean square
deviation.
44
Table 11.3. List of oligonucleotide primers used for the creation of A79AAG mutants by PCR based site
directed mutagenesis. The codons corresponding to the mutated amino acid are underlined.
A79AAG
mutants
Primer sequences (5'-3')
Forward- GTACTTCTGCATGGCCATCTCCAGCC
Asnl69Ala
Reverse- GGCTGGAGATGGCCATGCAGAAGTAC
Forward- GTACTTCTGCATGCTCATCTCCAGCC
Asnl69Leu
Reverse- GGCTGGAGATGAGCATGCAGAAGTAC
45
Figure 11.1. Biochemical characterization of AAG variants with oligomers containing etheno
lesions. (A) Representative gels showing the results of gel mobility shift assays for A79AAG binding to
eA:T (FA paired opposite T) and cC:G (eC paired opposite G) 25-mer DNA duplexes. 2 nM 1 2P-labeled
oligonucleotide was incubated with an indicated concentration of A79AAG in the lx binding buffer, and
the resulting protein-DNA complexes were resolved using 6% native-PAGE. The top band corresponds to
the protein-DNA complex and the bottom band corresponds to the free DNA. (B) Graphical
representation of A79AAG binding to eA:T 25-mer (n), eC:G 25-mer (0), eA:T 13-mer (e) and eC:G
13-mer (o) oligonucleotide duplexes as measured by gel shift mobility assays. Error bars indicate the
standard deviation of at least three independent trials. (C) Representative gels showing the results from
competition DNA glycosylase assays. The activity of A79AAG on labeled EA:T 25-mer duplex is
inhibited in the presence of indicated concentrations of eA:T 13-mer and eC:G 13-mer unlabeled
competitors. 2 nM 32 P-labeled eA:T 25-mer duplex oligonucleotide was incubated in lx glycosylase assay
buffer with 5 nM A79AAG enzyme and increasing concentrations of cold competitors at 37*C for 30 min.
Following hot alkali treatment, the products were resolved using 20% denaturing Urea-PAGE. The top
band represents the uncleaved DNA substrate and the bottom band represents the cleaved product. (D)
Graphical representation of the data from competition DNA glycosylase assays, looking at the inhibition
of A79AAG activity on labeled eA:T 25-mer duplex substrate by eA:T 13-mer (e) and EC:G 13-mer (o)
unlabeled competitors. Error bars indicate the standard deviation of at least three independent trials. (E)
Gel results of DNA glycosylase assays for full-length AAG and truncated A79AAG on eA:T and eC:G
25-mer DNA duplexes with E. coli Mismatch Uracil DNA Glycosylase (MUG) as a positive control.
Single strand breaks are observed in this Urea denaturing gel when piperdine treatment cleaves AP sites
(bottom band). When AAG fails to cleave the lesion base, no AP site is formed and the oligomer remains
intact (top band).
46
A
B
eAT 25-mer
Substrate
A79AAG (nM)
0
eC:G 25-mer
5 10 25 50 100 250 500 0
1 2.5
5
10 25
50
100 250
100N
1A
7V
W,
4w
0i
C
4ao2iO2O 400
D
Competitor
Competitor (nM) -
0
eA:T 13-mer
cC:G 13-mer
25 50 100 200 300 400 500 600 5 10 20 40 60 80 100 200
e
EIpw
sS
400
700 000
Sgo
1000
IA7SAAG) nM
Ift
a
* 301
10
o
4to0
jConii.Itkorj nM
E
FL -AAG
A79AAG
Substrate
tA:T 25-mer
EA:T 25-mer
Time (min)
0
30
60
S.A*
0
30
FL-AAG
MUG
60
A79AAG
EC:G 25-mer
EC:G 25-mer
0
0
30
60
30
60
EC:G 25-mer
0
30
60
0-Ao
47
Figure 11.2. Gel results of DNA glycosylase assays for truncated A79AAG on 6A:T and cC:X
(X=G/A/T/C) 13-mer DNA duplexes used for crystallization.
EA:T 13-mer
Time (min)
A79AAG
48
60
-
30
+
60
+
EC:G 13-mer
60
-
30
60
+
+
EC:A 13-mer
60
30
60
-
+
+
EC:C 13-mer
EC:T 13-mer
60
-
30
60
+
+
60
30
60
-
+
+
Figure 11.3. Structure of A79AAG with sC inhibitor DNA. (A) Overall structure of the A79AAG-EC:G
complex; the protein is in green ribbons with Tyr162 in stick form. The DNA is colored as follows:
carbon, yellow; oxygen, red; nitrogen, blue; and phosphorus, orange. (B) Schematic illustration of the
interactions between the amino acid side chains (3 letter code) and main chains (mc) of the AAG protein,
and EC containing DNA in the structure of the A79AAG-EC:G complex. Hydrogen bonds are indicated by
a solid line and van der Waals interactions by a dashed line. The disordered A26 nucleotide is in dashed
lines. (C) Intercalation of Tyrl62 (carbons in green) in the A79AAG-EC:G structure, and coordination of
metal ion Mn2 (orange sphere) to the A18 and G19 of the DNA strand opposite the EC lesion (carbons in
yellow), and to three water molecules (red spheres). Distances (dashed lines) are measured in Angstr6ms
(A). The electron density for the metal ion site is shown in Figure II.6A. Other non-carbon atoms are
colored as in (A).
B
3'
G14
C1
G15 |C1
16
GlO
ArgI97
Tyr165
T9
A17
m219
AI|
TS
#Igle
mc163
G19-T 162
Met164-
=c220
182
Asn169
Tr12
vC7
?98
HisI36 Tyr159 Tyr157
Gly163-
C
20
G6
A21
T5
mc136
Lys229
mc143
T22
A4
G23
C3
m=145
T24
AI162\
Te
G6
C2
A18
2.3
2.3
p
1 C20
.2
3.
- I
A2
G1
5f
G19
49
Figure 11.4. Comparison of the overall structures of the A79AAG-DNA complexes. The
superposition of AAG-C:G (green) and AAG-EA:T (gray) (PDB ID 1F4R; (10)) complexes with the
carbons of the flipped EC nucleotide colored yellow, and the Tyr162 side chain indicated with an arrow.
Regions that show differences with respect to disordered loops are indicated by dashed ellipses. The
region 1 (left) corresponds to the residues Gly265-Gly268; and region 2 (right) corresponds to
the
residues Leu249-Pro254.
Tyr1 62
50
-
Figure 11.5. Cation site in A79AAG structures. (A) In the structure of the A79AAG-FA:T substrate
complex (PDB ID 1F4R (10)), a Na' ion (purple sphere) is coordinated by AAG (carbons in gray) and a
water molecule (red sphere). (B) In the structure of A79AAG-FC:G, Na' is replaced by the N-terminal
NH 3 ' of Gly80 from a symmetry related molecule (carbons in cyan) in interacting with the A79AAG
molecule (carbons in green). All the residues in cyan, Gly80, Pro8l and His82 (along with Met83 not
shown in the figure), are part of the precision protease cleavage site sequence and do not represent the
wild type AAG sequence. All non-carbon atoms colored as in Figure 11.3. For both (A) and (B), hydrogen
bonds are indicated by dashed lines with distances given in Angstr6ms (A).
B
A
Ser1 71
Ser171
Sery72
Ser 729
2
Ala177
.
4
2.6
+
2.6
Met149
Met1 49
%9 3 2
Ala 177
Gly80
,'2.8
Pro81
His82
Glyl74
Gly1 74
51
Figure 11.6. Interaction of Tyr162 and putative Mn 2 ' binding site in the structure of A79AAGEC:G. (A) 2mFI-DIFeI omit electron density map contoured at la (gray) is drawn around the Mn 2 , ion
(orange sphere) and the DNA bases (A18 and G19) that coordinate the metal ion. An anomalous
difference electron density map calculated with the native dataset (k = 1.116 A) contoured at 8a
(magenta) shows a strong positive peak for an anomalous scatterer such as Mn 2 , (chain A site depicted).
Water molecules that coordinate the Mn 2 , ion are not shown. Carbons are in yellow and non-carbon atoms
are colored as in Figure 11.3. (B) Difference in the sugar pucker and conformation of DNA backbone near
the Mn 2 1 ion in the structure of A79AAG-EC:G compared to A79AAG-EA:T (carbons in gray) (PDB ID
IF4R (10)). All other atoms are colored as in (A). (C) The Tyr162 intercalation site is near a Mn 2 + ion and
Metl64 (sulfur atom in yellow), which both interact with G19 by using hydrogen bonding and Van der
Waals interactions respectively. Protein carbons are in green and all other atoms are colored as in (A).
52
A
G6
A121
B
*
0
A18
C2
8T19
C2'-exo
f'
C2'-end~
18
C
e66
1644
G1
53
Figure 11.7. Active site architecture of AAG. (A) Active site of the A79AAG-EC:G inhibitor complex
with DNA and amino acid carbons in yellow and green respectively and a putative catalytic water
molecule as a red sphere. Hydrogen bonds are indicated as dashed lines. A stereoview of the electron
density of this interaction is shown in Figure 11.10. (B) Active site architecture of the A79AAG-EA:T
substrate complex (carbons in gray) showing the amino acids interacting with a flipped FA base and the
DNA backbone (carbons in yellow) (PDB ID lF4R (10)). (C) Comparison of the active site architecture
of A79AAG-EC:G (protein carbons colored green, DNA carbons colored yellow and catalytic water
colored red) and A79AAG-FA:T (all carbon atoms and catalytic water colored gray). The movement of
the DNA backbone in the A79AAG-EC:G complex due to the binding of EC is indicated by a black arrow.
All non-carbon atoms are colored as in Figure 11.3.
A
GkuI25
Arg182
=i262
Lu80
*Tyr57
Tr159
B
Glu125
Tyr7
Asn169
Arg82
LU180
Ty157
tA
yr59
C
Tyr'127
4G025
Ser286
Arg182 .A
0 tC-3'
54
HOW13
Figure 11.8. AAG binding pocket. A side (A) and top (B) view of EC in the A79AAG-EC:G structure
with Van der Waals surfaces shown in gray spheres. (C) 2mIF.I-DIFI omit electron density map contoured
at 1o (gray) is drawn only around EC:G containing DNA and a putative catalytic water molecule (red
sphere). All atoms are colored as in Figure 11.3.
B
A
Asn169
Leu180
CysI67
Asn169
Leul W
GIu125
Cys178
Tyr159
Tyr1 27
6C
CW
Met149
His136
His1 36
Ata 134.
Al134'
Ala1 35
Ala135
C
Tyr157
Asn 169
Leul80
Aris136
GW01259
'I
C
Arg182
Tyr162-
55
Figure 11.9. Activation of the leaving group by protonation. (A) Superposition of the structure of the
A79AAG-EC:G inhibitor complex (amino acid carbons in green; DNA carbons in yellow; and water as a
red sphere) with that of the A79AAG-E125Q-EA:T substrate complex (amino acid carbons, DNA carbons,
and waters are in gray) (PDB ID lEWN (10)). All other non-carbon atoms are colored as in Figure 11.3.
The hydrogen bonding contacts are indicated as black dashed lines. A magenta dashed line represents a
putative hydrogen bonding contact that could stabilize a protonated state of LA. An arrow indicates a
water molecule that could protonate N7 of LA. (B) Schematic illustration of the recognition of EA (top)
and eC (bottom) by active site residues and of the structure of Hx (right) with each base numbered to
correlate with the text. Arrows indicate the proposed protonation site on EA (N7) (top), and the
corresponding position in EC (C5) and Hx (N7).
A
HJs
36
Ala134
N7(rA
C5(4C
OeC sense.
Asn169
B
NH.
Aa169
<A
NH
I
2
Hx
DNA
Asnl89
EC
56
Figure 11.10. A wall-eyed stereoview of the active site of AAG. 2mIFI-DIFeI omit electron density map
contoured at 10 (gray) is around EC:G containing DNA, active site protein residues and a putative
catalytic water molecule. Atoms are colored as in Figure 11.3.
Tyr27
Tyr127
pw4G u125
G u125
Asn169
Arg1 82
_EC
Asn 169
Arg1 82
His136,/
IC
Tyrl59
His136,
Tyrl 5
57
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59
60
Chapter 3. Searching for DNA Lesions: Structural Evidence for Lower- and
Higher-Affinity DNA Binding Conformations of Human Alkyladenine DNA
Glycosylase
Reprinted with permission from "Searching for DNA lesions: Structural Evidence for Lowerand Higher-Affinity DNA Binding Conformations of Human Alkyladenine DNA Glycosylase.
Jeremy W. Setser, Gondichatnahalli M. Lingaraju, C. Ainsley Davis, Leona D. Samson,
Catherine L. Drennan. Biochemistry. 2012; 51(1):382-390." C 2012 American Chemical
Society.
111.1
SUMMARY
To efficiently repair DNA, human alkyladenine DNA glycosylase (AAG) must search the
million-fold excess of unmodified DNA bases to find a handful of DNA lesions. Such a search can be
facilitated by the ability of glycosylases, like AAG, to interact with DNA using two affinities: a
lower-affinity interaction in a searching process, and a higher-affinity interaction for catalytic
repair. Here, we present crystal structures of AAG trapped in two DNA-bound states. The
lower-affinity depiction allows us to investigate, for the first time, the conformation of this protein
in the absence of a tightly bound DNA adduct. We find that active site residues of AAG involved in
binding lesion bases are in a disordered state. Furthermore, two loops that contribute significantly
to the positive electrostatic surface of AAG are disordered. Additionally, a higher-affinity state of
AAG captured here provides a fortuitous snapshot of how this enzyme interacts with a DNA adduct
that resembles a one-base loop.
61
III.I
INTRODUCTION
Genomic DNA is under constant attack from endogenous and exogenous sources with
most damage occurring in the form of DNA base lesions (1). While approximately 10,000 of
these lesions occur daily (1, 2), most do not go on to harm the cell because they are repaired by
endogenous pathways. One of the most prevalent DNA repair pathways is known as base
excision repair (BER), which is initiated when a DNA glycosylase hydrolyzes the N-glycosidic
bond of a lesion base. In humans, the abasic site produced by a monofunctional glycosylase is
repaired by the subsequent action of AP endonuclease I, DNA polymerase P, and DNA ligase I
or III (3, 4).
Human alkyladenine DNA glycosylase (AAG) is one of the monofunctional glycosylase
enzymes responsible for initiating BER. AAG catalyzes the removal of a diverse group of purine
lesions, including those caused by damage from alkylation (3-methyladenine, 3-methylguanine,
and
7-methylguanine)
and
reactive
oxygen
and
nitrogen
species
(hypoxanthine,
1,N 6-ethenoadenine (EA) and 1,N 2-ethenoguanine) (5, 6). Removal of these lesions is paramount
as they can cause cytotoxicity and mutagenesis (7). To access lesion bases, AAG, like most
glycosylases, uses the canonical nucleotide-flipping mechanism wherein the nucleoside with the
damaged base is flipped out of the double helix and into the active site while a protein residue
intercalates the DNA, effectively substituting for the flipped base. This nucleotide-flipping has
been observed in the crystal structure of a catalytically active N-terminal truncation mutant of
AAG (denoted A79AAG) in which the protein is bound to DNA containing substrate FA (8).
This structure shows that Tyr162 of AAG intercalates DNA while the lesion fits snugly into the
binding pocket. This tight interaction observed structurally is supported by the nanomolar
affinity of AAG for its substrates in vitro (9-11). Interestingly, AAG also binds with high affinity
to DNA containing lesions that it cannot excise, such as inhibitor 3,N 4-ethenocytosine (FC) (5, 9,
12). Structural studies show that EC is also flipped out of the DNA into the active site of AAG
and that an extra hydrogen bond between AAG and cC accounts for the two-fold higher affinity
for the inhibitor over the substrate DNA (9). Finally, AAG can also bind with high affinity to
DNA with a base loop structure, shielding it from repair and leading to frameshift mutations
(13). These highly specific interactions (outlined in Figure 111.1) between AAG and DNA are
even more intriguing when one considers the massive search that must be undertaken by DNA
glycosylases to find damaged DNA bases in the human genome.
62
Given the ~101o nucleotides in the human genome and the -10' lesions per cell per day
(1, 2), there are approximately one million normal bases for every lesion present in DNA. Even
for an abundant protein like AAG (-2 x 10' molecules per nucleus (14)), each enzyme would
have to inspect tens of thousands of normal bases before finding one lesion to excise. Such a task
would be seemingly impossible if it involved a strict three-dimensional search, where proteins
float through the cell in a stochastic hunt for a scarce number of lesions. In order to limit the
search space, it has been proposed that DNA binding proteins could nonspecifically bind and
track along DNA in a one-dimensional search (15, 16). Recently, protein 'sliding' on DNA was
observed directly in single-molecule fluorescence studies for a number of enzymes including
several glycosylases (17-19). Such a nonspecific search has been indirectly observed for AAG
using kinetic assays in which the ability of the enzyme to excise two lesions contained in one
piece of duplex DNA was examined (20, 21). Kinetic data are also available that indicate AAG is
able to search both strands of substrate DNA and avoid obstacles using a 'hopping' mechanism
(20). While the ability to slide or hop along DNA requires a lower-affinity and nonspecific
complex between protein and DNA, base excision requires high-affinity and specific
interactions. Thus, one would expect AAG, and related enzymes, to have differential modes of
DNA binding. Evidence in support of this idea is available for other glycosylase systems,
including crystal structures of a functional homologue of AAG from Escherichia coli (AlkA)
(22, 23), as well as a crystal structure (24) and single-molecule data (17) for human
8-oxoguanine DNA glycosylase (reviewed in (25)).
In our current structural studies, we have captured two novel states of AAG. One
structure shows AAG making only nonspecific contacts with DNA, depicting a 'lower-affinity'
or 'searching' protein-DNA complex. The other shows a higher-affinity complex in which AAG
is bound to two pieces of single stranded DNA each containing an *C lesion (sseC) in an
arrangement that resembles a single base loop structure. By comparing these structures to each
other, and to previously solved structures of AAG bound with high affinity to double-stranded
DNA (dsDNA) (8, 9, 26), we can investigate the molecular basis for the differential affinities of
this DNA repair protein for DNA and explore the recognition events involved in identifying
DNA lesions.
63
111.111 RESULTS
Structuraloverview of asymmetric unit
The structure of A79AAG in the presence of stoichiometric amounts of single-stranded
SC-containing DNA (ssEC) was determined to 2.0
A resolution
by molecular replacement using
the previously solved structure of A79AAG bound to pyrrolidine-containing DNA (abbreviated
A79AAG-pyr:T) (PDB ID 1BNK, (26)) as a search model. The final structure, with two
molecules of A79AAG in the asymmetric unit, has been refined to an R factor of 21.9 (Rfree =
26.5) (Table 111.1). Instead of observing two A79AAG molecules each bound to one ssFC DNA,
we obtained two different and unique structures of this protein. During crystallization, the ssEC
oligonucleotides formed a self-complementary pseudo-duplex, which is specifically recognized
by a single molecule of A79AAG in the asymmetric unit (Figures III.2A and III.2B, in orange).
We will refer to this interaction as the pseudo-duplex structure. Although the other molecule of
A79AAG in the asymmetric unit is also interacting with an EC-containing DNA strand, it only
makes nonspecific contacts with the phosphodiester backbone and leaves the EC lesion
untouched (Figures III.2B and 111.3). This nonspecific protein-DNA interaction will be referred
to as the lower-affinity structure (Figures III.2A and III.2B, in green). The 13-mer pseudo-duplex
piece of DNA that we observe crystallographically is highly unlikely to persist in solution, which
precludes traditional binding measurements. We have studied the binding of A79AAG to 13-mer
ssEC oligonucleotides by gel shift assays as previously described (9) and found no measurable
affinity (Figure 111.4). These same assays have shown high affinity A79AAG binding (Kd of 21 E
3 nM) for pre-annealed 13-mer doubled stranded FC oligonucleotides and this highly specific
interaction is depicted by a crystal structure with the same dsDNA (9). With these data in mind,
the molecules of A79AAG shown in Figure 111.2 must have affinities for their 13-mer
oligonucleotides that fall in the range from immeasurably weak, as observed for true single
stranded DNA, to the tight binding (Kd of 10-23 nM) measured for pre-annealed dsDNA (9).
Considering the green molecule (Figure 111.2) has only a few nonspecific contacts to the DNA,
whereas the orange molecule has many specific contacts and closely resembles the high-affinity
structures solved previously with dsDNA (8, 9, 26), these structures appear to represent lowerand higher-affinity states, respectively, and will be referred to as such.
64
J 79AAG pseudo-duplex structure
In the pseudo-duplex structure, the canonical nucleotide-flipping mechanism of DNA
glycosylases can be observed with Tyrl62 inserted into the DNA duplex while the lesion
nucleoside (EC7') from one ssEC strand is flipped into the enzyme active site (Figure III.2A
Panel I). Interestingly, the active site interactions with the FC lesion for this pseudo-duplex
structure are identical to those of A79AAG with dsDNA (abbreviated A79AAG-EC:G) (PDB ID
3QI5, (9)) and both structures share high overall similarity with a root-mean-squared deviation
(RMSD) between alpha-carbons of 0.43
A
(Figure 111.5). This pseudo-duplex structure is also
highly homologous to the structure of A79AAG with substrate lesion EA in dsDNA (abbreviated
A79AAG-FA:T) (PDB ID 1F4R, (8)), with an RMSD for a-carbons of 0.93
A.
Although nucleotide-flipping is observed in the pseudo-duplex structure, the interactions
surrounding the intercalated residue (Tyr162) are not identical to those previously observed in
the structure of AAG with SC-containing dsDNA. In the A79AAG-FC:G structure, a potential
steric clash of Tyrl62 with G19 (base opposite EC) is prevented by a shifting of G19 out of the
minor groove, leaving it without a base pairing partner (Figures III.2C and III.2E) (9). In
contrast, A2 of the pseudo-duplex structure avoids a steric clash with a sideways motion that
allows for hydrogen bonding to T8' of the opposite strand (Figure III.2D, III.2F). This sideways
motion also changes the orientation of the neighboring base G 1, such that it now hydrogen bonds
to T9' (Figure III.2D). Although Metl64 contacts the 'opposite bases' (G19 and A2) in both
structures, the orientation of the interaction is also different (Figures III.2E and III.2F).
A79AAG lower-affinity structure
The lower-affinity structure of A79AAG shows only nonspecific interactions (Figures
III.2A and III.2B, in green), with hydrogen-bonding contacts to the phosphodiester backbone by
the side chains of Argl82, Argl97, and Arg207 and the main chain amides of Ser219 and
Lys220 (Figures III.2B and III.6A). Interestingly, the protein residue that normally intercalates
substrate DNA, Tyr162, is contacting the pseudo-duplex by stacking with nucleotide A2' (Figure
III.2A Panel II and Figure III.6B). While this Tyr162 adopts a similar orientation as found for
intercalated Tyrl62 residues from the higher affinity complex structures (e.g. Figure III.7A vs.
III.7B), it has higher B-factors, indicating increased conformational flexibility (see Figure
III.8B). In addition to stacking with Tyrl62, A2' hydrogen bonds with T8, leaving eC7 orphaned
65
in terms of base pairing (Figure III.6C). This aC7 lesion also has no interaction with protein
residues (Figure 111.3).
Overall, the structure of lower-affinity A79AAG is similar to other structures of this
protein, including the pseudo-duplex structure described above (RMSD for a-carbons = 1.27
= 1.26
the A79AAG-EC:G complex (RMSD for a-carbons
A79AAG-EA:T (RMSD for a-carbons
lower-affinity
= 1.18
A)
A)
A),
(9), and the structure of
(8). While these RMSDs are low, the
structure has three distinct disordered regions when compared to the
A79AAG-EA:T structure (Figures III.7A and III.7B). Since there are no lattice contacts in this
area (Figure 111.3), we can attribute the disorder to the absence of bound nucleotide in the active
site of lower-affinity AAG. The residues that lack electron density in the low-affinity complex,
and are thus considered disordered, include Glul3l-Argl4l (Loop 1), Gly263-Lys273 (Loop 2),
and C-terminal residues after Asp289 (Loop 3) (Figure III.7A and III.7B).
Loop 1 contains
crucial active site residues, including Ala134-Hisl36, which form a snug pocket for lesion bases
(Figure III.7C). This binding pocket is only partially formed in the absence of nucleotide (Figure
III.7D). Disordered loops 2 and 3 are not involved in forming the active site but contribute to the
electrostatic potential of the protein (Figures III.7E-III.7H).
Electrostatic
surfaces for
A79AAG-aA:T
(after removing
the DNA)
and for
lower-affinity A79AAG are considerably different (Figures III.7E and III.7F) (All electrostatic
depictions were calculated using the Adaptive Poisson-Boltzman Solver (APBS) software
plug-in (27) for PyMOL (28)). The A79AAG-aA:T complex shows a continuous, and richly
positive, DNA-binding surface as would be expected for a protein that contacts DNA with high
affinity (Figures III.7E and III.7G). In contrast, the DNA-binding surface for the lower-affinity
structure is more neutral with disordered loops 1-3 disrupting positive patches observed in the
A79AAG-EA:T high-affinity complex (Figures III.7E-III.7H). Other regions of the protein show
little difference in the ordered electrostatic surface, such as the area near the intercalating residue
Tyr162 (marked with a star in Figures 111.7 and 111.9). Also, both structures display a positively
charged electrostatic surface that circles the protein molecule from the top of A79AAG to the
bottom (see middle view of Figures III.9A and III.9B), as well as a negative electrostatic region
located opposite the DNA binding surface (Figure 111.9).
66
III.IV DISCUSSION
DNA glycosylases are charged with the formidable task of locating and repairing
potentially harmful DNA lesions while avoiding the million-fold excess of normal, healthy DNA
bases. The difficulty of this searching process can be partially overcome by the formation of a
weak complex between protein and DNA, effectively creating a nonspecific, one-dimensional
search. However, to maintain fidelity and genomic integrity, the enzyme must also be able to
form a stronger, highly specific complex for lesion recognition and excision. Therefore, the
ability to adopt both low and high-affinity conformations appears advantageous. Here, we have
trapped a human DNA glycosylase, AAG, in both lower and higher-affinity complexes with
DNA (Figure 111.2), providing two snapshots of this enzyme that relate to this search process.
Interestingly, AAG has been shown to bind with high affinity to DNA damage that it can
repair (such as t-A lesions) as well as to damage that it cannot repair (such as EC lesions and
one-base loops) (9-11, 13). Crystallographic studies have provided molecular insight into how
AAG recognizes both EA and EC lesions within dsDNA (8, 9), but no structure of AAG bound to
a one-base loop or in a low affinity complex with DNA has been determined. The pseudo-duplex
structure that we present here appears to be the best representation available of how AAG could
bind to such a DNA loop structure. In a one-base loop, one nucleoside is 'looped out' of the
DNA, and the base opposite the 'looped out' base hydrogen bonds with an adjacent base instead
(Figure III.1C). This arrangement of bases resembles what we observe in the pseudo-duplex
structure and represents a major departure from the hydrogen bonding pattern of nucleotides
observed in other AAG-DNA complexes (e.g. Figure III.2C). The 'looped out' base is nicely
accommodated in the AAG active site with identical interactions to those observed previously
for an EC lesion (Figure III.5B) (9). In addition, the close resemblance of the pseudo-duplex
structure to previously solved structures of A79AAG bound to dsDNA (8, 9, 26) is consistent
with the idea that this structure represents a high-affinity complex between AAG and DNA. This
observation is in agreement with the high-affinity binding observed between AAG and a
one-base loop structure in vitro (13). A physiological rationale for why AAG binds to DNA
damage that it cannot repair remains to be determined: while tight binding of AAG to lesions it
can repair such as -A, can be beneficial to the cell (29), tight binding of AAG to base loop
structures shields them from repair, increasing mutation rates (13). As the physiological
67
significance of this behavior of AAG is elucidated, our work suggests a molecular basis for the
recognition of base loops by this human DNA repair protein.
Excitingly, our crystallization conditions have also yielded the first nonspecific or
lower-affinity depiction of AAG, providing insight into a conformation of the protein likely
responsible for inspecting DNA for damage. Although the top side of the active site, including
the position of the putative catalytic water, agrees well with high-affinity lesion-bound structures
(Figure III.5B), the residues comprising the active site floor are disordered (Figure 111.7). This
observation of a partially ordered active site suggests an order of events for the binding of AAG
to DNA in which a lesion base is first identified by a more dynamic state of the protein, and is
later recognized with high-affinity as the active site pocket closes around the nucleotide-flipped
lesion. Our structural studies are consistent with fluorescence-based kinetic assays, which have
provided evidence for a two-state lesion recognition process for AAG, where the active site
experiences changes in environment prior to nucleotide flipping (10, 30). This initial state
observed kinetically has been likened to the initial recognition complexes suggested for other
glycosylases (25, 31).
With the lesion flipped into the active site, an intercalating residue (Tyrl62 for AAG)
maintains the double helical DNA structure. An interesting point of discussion in the DNA repair
literature is whether intercalating residues play an active or passive role in lesion recognition. In
other words, whether the intercalating residue directly interrogates base pairs (active), or the
success of the search relies on the intercalating residue filling the gap left behind by a flipped
lesion (passive). Two recent structural studies on the glycosylases MutM and the functional
homologue of AAG from E. coli, AlkA, have provided conflicting answers to this question. In
both studies, the glycosylases were linked to undamaged DNA in a stable complex using
disulfide crosslinks (22, 31), and the position of the intercalating residue was evaluated. For
MutM, the intercalating residue (Phe 114) is fully inserted into the DNA duplex, buckling the
bases with which it interacts, as the protein simultaneously bends the DNA, suggestive of an
active interrogation mode (31). In contrast, the structures of AlkA with undamaged DNA show
snapshots of a glycosylase in a more passive interrogation mode, with the intercalating residue
(Leul25) situated completely outside of a double-helix, which maintains all base-stacking
interactions and remains mostly linear (22). In our lower-affinity structure, the intercalating
residue of AAG, Tyrl62, has increased flexibility, but still maintains the same average position
68
for its sidechain as is found in the higher-affinity structures (e.g. Figure III.7A vs. III.7B).
Tyr162 is also still involved in a stacking interaction with a nucleotide (A2') even when
intercalation is not possible (Figure 111.6). This observation suggests that Tyrl62 is capable of
making both lower and higher-affinity interactions with DNA, possibly playing roles both in a
lower-affinity 'searching' process and in a higher-affinity 'recognition' process. Consistent with
an ability to form different types of interactions, the Tyri62 loop is flexible, displaying
approximately two-fold higher B-factors than average for this crystal structure (Figure 111.8).
Just as residues in the active site of AAG are disordered in the absence of a tightly bound
DNA lesion, residues that contribute to the positive electrostatic surface are also disordered
(loops 2 and 3 in Figures 111.3 and 111.7).
The highly positively-charged and complementary
surface of AAG that binds DNA with high affinity (Figures III.7E and III.7G) is disrupted in the
low-affinity structure (Figures III.7F and III.7H). Loops 2 and 3 are not pre-ordered, ready to
bind with high affinity to a DNA lesion. Instead, they are highly mobile, suggesting that they
could play an active role in interrogating DNA.
In terms of interrogating DNA, there is strong evidence that the searching process of
DNA binding proteins is not a strictly linear scan of DNA. A single-molecule study of eight
different DNA binding proteins, including three glycosylases, found that the movements of these
proteins along DNA was better described by a rotation-coupled sliding mechanism (17).
Such
movement would orient the enzyme so that its binding surface always faces the axis of the DNA
double helix. In essence, these proteins circle the DNA while diffusing along it. The electrostatic
potential surface calculated for AAG is consistent with this rotation-coupled search mechanism.
In both higher and lower-affinity AAG complexes, a positive electrostatic surface is found to
wrap around the protein (Figures III.9A and III.9B). This surface could be used to 'roll' or 'rock'
back and forth along the negatively charged DNA backbone while the presence of a negatively
charged electrostatic cap on the opposite face of AAG (red in Figure 111.9) would maintain
correct orientation for lesion recognition. 'Hopping', another DNA search method, has been
established for AAG through the use of kinetic assays (20). Hopping, or short-range dissociationassociation events, allows AAG to search both DNA strands simultaneously and avoid obstacles,
such as a DNA-encasing endonuclease like EcoRI, that may be present along the search path
(20). Rotation-coupled sliding and hopping are not mutually exclusive, and we consider both in
the proposed search mechanism for AAG that is outlined in Figure III.9C.
69
In the initial search, we propose that AAG closely resembles the lower-affinity structure,
interacting with DNA nonspecifically through its positive electrostatic surface. Incorrect
orientation of AAG would be avoided due to the negative electrostatic patch opposite the active
site (Figure III.9B). The positive surface that wraps around AAG would promote a rotationcoupled sliding search of the DNA, while still allowing for the hopping events described above.
As a lesion is recognized, disordered regions of AAG, including the active site pocket, become
more ordered (Figure 111.7). After nucleotide-flipping,
AAG
adopts
a higher-affinity
conformation such as the pseudo-duplex structure (Figure III.2A, in orange) or dsDNA structures
published previously (8, 9, 26). Here, previously disordered loops are completely ordered to
display the full potential of a continuous electrostatic surface for binding DNA, Tyr162 is fully
inserted into the DNA, and a base lesion is bound tightly in the AAG active site. This lesion
recognition complex would interact very strongly with the DNA, halting the search by AAG. In
cases where the lesion is a substrate, base excision would follow. After base-lesion release, the
active site and other loops of AAG would become partially disordered, decreasing the extent of
an ordered DNA binding surface, ultimately leaving AAG in its lower-affinity, nonspecific
searching state once again. In cases where the lesion cannot be repaired, AAG would remain
fixed in its higher-affinity state, providing a rationale for the abortive AAG-EC complexes
observed in vivo (12).
The two novel structures of human AAG presented here help provide a molecular
understanding of this intriguing DNA repair protein, both in terms of understanding how AAG
can recognize different types of DNA damage, such as base lesions and one-base loops, and how
it may search the genome for DNA damage. With recent literature describing an expanded
ability of AAG to both repair base lesions (5) and identify DNA damage that it cannot repair (5,
9, 13), this study provides important insight into the molecular basis of AAG interactions.
III.V MATERIALS AND METHODS
AAG plasmid construction and proteinpreparation
The A79AAG plasmid was constructed as described previously (9). Briefly, 84 residues
at the N-terminus of the protein were truncated in this construct, and four extra residues from a
PreScission Protease cleavage site (GE Healthcare) (Gly80, Pro8l, His82, and Met83) were left
70
behind after histidine-tag cleavage. Therefore, Thr84 begins the wild type AAG sequence but
four residues precede Thr84 such that Gly80 is now the N-terminus. We refer to this truncated
protein construct as A79AAG. It should be noted that the AAG protein from previous structural
studies was also referred to as A79AAG (8, 26). However, in those studies, all residues
contained in the construct are of the wild type sequence. The expression and purification of the
A79AAG protein was performed as described previously (see Chapter II.V, 9).
Crystallizationof A79AAG with single-strandedeC DNA
An equimolar ratio of A79AAG and 13-mer single-stranded EC-containing DNA (sseC)
(5'-GAC ATG ECTT GCC T-3') were mixed to form a protein-DNA complex concentration of
0.3 mM in the complex buffer (20 mM HEPES-NaOH pH 7.5, 100 mM NaCl, 0.1 mM EDTA,
5% v/v glycerol and 1 mM DTT). The complex was incubated on ice for 15 min and used for
crystallization. Crystals were obtained by the hanging drop vapor diffusion method upon mixing
1 pL of protein-DNA complex and 1 pl of reservoir solution (100 mM BIS-TRIS pH 5.5, 200
mM cesium chloride and 20% polyethylene glycol (PEG) 3350) over 0.5 mL of reservoir
solution. Crystals appeared after incubation for 14 days at 22*C. These crystals were
cryoprotected with precipitation solution supplemented with 10% glycerol and flash frozen in
liquid nitrogen prior to data collection.
Data collection and structure determination
X-ray diffraction data were collected at the Advanced Light Source (Berkeley, CA) on
beamline 12.3.1 at 100 K to 2.0
A resolution
and processed using Denzo/Scalepack (32) (Table
111.1). The structure, with two molecules in the asymmetric unit, was determined by molecular
replacement in PHASER (33) using the coordinates from the A79AAG-pyr:T complex structure
(PDB ID 1BNK, (26)). Refinement was carried out in CNS (34) and Refmac 5.4 (35, 36), using
topology and parameter files for the FC lesion generated by XPLO2D (37). Additional rounds of
refinement using TLS parameters and non-crystallographic symmetry restraints were very
effective in improving the quality of the fit. Model building was performed using the program
Coot (38), and figures were prepared using PyMOL (28). The final model converged to an R
factor of 21.9
(Rfree
= 26.5) (Table 111.1) and was evaluated using PROCHECK (39) and
composite omit maps. As was observed in our previously solved structure using this protein
71
construct (9), the positively charged N-terminus of both molecules of A79AAG in the
asymmetric unit occupies what was initially identified as a sodium ion site by Ellenberger and
co-workers (8). Although this coordination of the N-terminus is common to our A79AAG
structures, the packing of the molecules in the current study allowed AAG to crystallize in a
novel space group (P4 3). The following residues of the total sequence of 80-298 lack electron
density and are therefore not included in the model: 201-208, 265-266 and 294-298 in chain A
(pseudo-duplex structure); 131-141, 199-206, 253-254, 263-273 and 290-298 in chain B (loweraffinity AAG). Due to a lack of interpretable electron density for the side chains of some
residues in the structure (Leu200, Leu209, Glu250 in chain A; Prol44, Ile161, Met164, and
Arg212 in chain B), these residues were modeled as alanines. Nucleotides G 1' and CI '-T 13' of
the strand containing the nucleotide-flipped FC lesion, and T9-T13 of the pseudo-complement
strand are disordered.
III.VI ACKNOWLEDGEMENTS
This work was supported, in part, by the US National Institutes of Health Grants P30-ES002109
(to C.L.D. and L.D.S.), GM65337 (to C.L.D.), GM65337-03S2 (to C.A.D.), and CA055042 and
CA092584 (to L.D.S.). C.L.D. is an investigator of The Howard Hughes Medical Institute.
L.D.S. is an American Cancer Society Research Professor. J.W.S. is supported by a Repligen
KIICR Graduate Fellowship. The Advanced Light Source is supported by the Director, Office of
Science, Office of Basic Energy Sciences, of the U.S. Department of Energy under Contract No.
DE-AC02-05CH 11231. The atomic coordinates have been deposited with the Protein Data Bank
(http://www.rcsb.org) under the PDB accession code 3UBY. Author contributions: G.M.L.
performed protein expression and purification, crystallization, and data collection. C.A.D. carried
out the data processing, molecular replacement, and initial refinement. G.M.L. and C.A.D.
started the structural analysis, while J.W.S. carried out the final rounds of refinement and
completed the structural analysis. G.M.L. and L.D.S were involved in study design, J.W.S. and
C.L.D. wrote the manuscript, and all authors edited the manuscript.
72
TABLES & FIGURES
Table 111.1. Data collection and refinement statistics for A79AAG-DNA complex.
Space group
P43
a = b =41.17, c = 262.55
a= P= Y= 90
ALS 12.3.1
Cell constants (A)
(0)
Beamline
1.116
Wavelength (A)
66 - 2.00 (2.07 - 2.00)
Resolution (A)
No. total observations
95966
No. unique observations
26998
Completeness (%)
92.4 (89.6)
c
<I/(C)>'
14.8 (7.4)
Rsym (%) a, c
7.4 (15.8)
Model refinement
Rwork (%) b
21.9
Rfr,,ee (%)
26.5
b
B-factors (A2)
Protein
DNA
Water
35.6
48.9
22.3
RMSD bonds (A) d
RMSD angles
0.008
1.2
(0) d
Number of atoms
Protein
DNA
Water
Ramachandran plot (%)
2965 (2 molecules/asu)
353
250
Most favored
91.0
Additionally allowed
9.0
Generously allowed
a: Rsym= Ihk
where I is the intensity of a reflection hkl and <I> is the average over
symmetry-related reflections of hkl.
b: Rwork = IF 0 - FeI/1JFI in which F and Fc are the observed
and calculated structure factor amplitudes,
0
respectively. Rfree is calculated from 5% of the reflections not used in the model refinement.
C: Values in parenthesis correspond to the highest
resolution shell.
d: RMSD, root mean square
deviation.
-
hk>1/X
hk,
73
Figure 111.1. DNA adducts to which AAG binds with high affinity: Lesions (A) EC and (B) EA and (C)
one-base loop structures.
A
B
EC
G
74
C
EA
T
A
AA
TT
it
Figure 111.2. Structures of A79AAG bound to EC DNA. (A) A79AAG crystallized in the presence of
ssEC DNA has two A79AAG molecules in the asymmetric unit: one that makes few contacts to DNA and
represents a lower-affinity complex (green) and one that makes multiple contacts to DNA and represents
a higher-affinity complex (orange). The two strands of ssFC DNA, which form a pseudo-duplex, are
shown as sticks with cyan carbons. Panel (I) displays Tyr162 (in orange stick) intercalating DNA while
the FC lesion is flipped into the active site. Panel (II) depicts the lower-affinity interaction between
A79AAG and DNA where Tyr162 (in green stick) stacks with nucleotide A2'. Atoms are colored as
follows: oxygen (red), nitrogen (blue), and phosphorus (orange). A blue star denotes the location of the
empty active site of lower-affinity AAG. (B) Schematic illustration of the interactions between the two
strands of ssEC DNA and amino acid side chains (3-letter code) and main chains (mc) of the A79AAG
molecules. Amino acid labels from the lower and higher-affinity (pseudo-duplex bound) A79AAG
molecules are colored green and orange respectively. Hydrogen bonds are indicated by a solid line and
Van der Waals interactions by a dashed line. DNA bases are shown as rectangles containing one-letter
codes and numbers that signify their respective positions in the oligonucleotide (5' to 3'). All DNA bases
contained in the nucleotide-flipped EC lesion strand are denoted with a prime symbol ('). Disordered
nucleotides are in dashed lines. (C) Nucleotide interactions near lesion in A79AAG-EC:G dsDNA (PDB
ID 3QI5 (9)) (yellow carbons). Relevant distances shown by dashed lines are given in Angstr6ms (A). (D)
Nucleotide interactions near lesion in pseudo-duplex A79AAG structure (cyan carbons). (E) Van der
Waals interactions with G19 in A79AAG-EC:G structure. (F) Van der Waals interactions with A2 in
pseudo-duplex A79AAG structure.
A
3'
B
C12'
C11'
G10'
C7
r1
T9'
5'
Tyr1 62
C
A
T8'
-
C3
G6'
A4
T5'
D
A17
2.
GI
9
G19
C
T5
'3.
T8
T8
2
A18
p
G6
A4'
Arg207
Arq 197
3.6T8
G6
C3'
nc219
rnc220
C3
Arg82
.T r162
A21
-
Gil
EC7
E
T9
-Met164
5,
A2
C11
G19'
Tr 162
Tyr162
12
T13
3'
75
Figure 1113. Wall-eyed stereoview of the disordered loops of the lower-affinity A79AAG structure
in the context of the crystal lattice. The asymmetric unit of our structure is represented as in Figure 111.2
with lower-affinity A79AAG and the higher-affinity pseudo-duplex structure in green and orange cartoon,
respectively, with the pseudo-duplex piece of DNA between them in stick form with cyan carbons.
Molecules that make up the rest of the crystal lattice are in gray with protein in cartoon and DNA in stick
form. The disordered loops of lower-affinity A79AAG are visualized by aligning the A79AAG-EA:T
structure (PDB ID 1F4R (8)) and displaying these loops as pink cartoon. Regions that become disordered
in the absence of a bound DNA adduct are circled and labeled 1-3. This numbering scheme matches that
of Figure 111.7. With this view, one can see that these loops would have ample space in the crystal packing
and the disorder observed in these residues is not due to crystal contacts. The EC base that is not bound in
an AAG active site (in stick form with yellow carbons (FC7)) is found to have no interactions with protein
(or DNA) in the crystal lattice. Non-carbon atoms are colored as in Figure 111.2.
76
Figure III.4. A79AAG shows no affinity for ssEC 13-mer by gel shift. Gel mobility shift assays were
performed as previously described (9) using the indicated concentrations of A79AAG and 2 nM of
32
P-labeled ssEC 13-mer. The band at the bottom of the gel represents the ssEC 13-mer in solution. A clear
band shift is not observed as the amount of A79AAG is increased and the smearing of the gel in the later
lanes is indicative of some low-affinity interactions. This same assay performed with 13-mer duplex DNA
containing the EC lesion shows a very clear band shift with a Kd of -20 nM (9).
Substrate
A79AAG (nM)
sseC 13-mer
0
5
10 25
50 100 250 500 1000
77
Figure I.5. A79AAG structural comparisons. (A) A wall-eyed stereoview
of A79AAG bound to
dsDNA (protein in blue ribbon, DNA in yellow sticks) and pseudo-duplex
DNA (protein in orange
ribbon, DNA in cyan sticks) shows striking similarities in binding mode. The
intercalating Tyr162 (in
sticks) is denoted with an arrow and the EC lesion is colored such that carbon
atoms match the rest of the
respective DNA. All Non-carbon atoms are colored as in Figure 111.2. (B) Active
site overlay of A79AAG
bound to EC-containing dsDNA (protein carbons in blue) and pseudo-duplex
EC DNA (protein carbons in
orange) with the lower-affinity structure (protein carbons in green). The putative
catalytic water molecule
that is present in all AAG structures is denoted as a sphere and colored to match
the protein carbon atoms
of its respective structure. DNA carbons and all other non-carbon atoms are
colored as in (A). Hydrogen
bonding is indicated as dashed lines.
A
TyrB62
Tyr162
GIu2
Leu180
Asn169
Arg 182
Tyr157
H20
Hi3
Smc262
T159
78
His1 36
Figure 111.6. Tyr162 contacts in lower-affinity A79AAG. (A) Hydrogen bonding contacts (dashed
lines, distances in A) for lower-affinity A79AAG (green) with pseudo-duplex DNA (carbons in cyan),
and non-carbon atoms colored as in Figure 111.2. (B) Van der Waals radii for protein and DNA are shown
in gray spheres with all other representations and colors as in (A). (C) The same depiction as in (B) with
the orientation changed slightly to show relevant distances (in A) as depicted by dashed lines and to draw
attention to the rotation of FC7 out of the pseudo-duplex.
A
Tyr162
3
A2'
:2.5
02.8
Arg 197
Lys220
2.9
2.9
Ser219
Arg 182
B
CC3'
G6
3
s3.
&C7
T8
7%
C3'
:3.7
43
79
Figure 111.7. Comparison of the lower-affinity A79AAG structure with the high-affinity A79AAGEA:T structure. (A) A79AAG (purple cartoon) bound to an FA lesion (stick form with cyan carbons)
with active site residue His 136 and intercalating Tyr 162 represented in stick form with purple and yellow
carbons respectively (PDB ID 1F4R, (8)). Non-carbon atoms are colored a in Figure 111.2. Regions that
become disordered in the absence of a bound DNA adduct are circled and labeled 1-3. This numbering
scheme matches that of Figure 111.3. (B) Lower-affinity AAG (green) with Tyr162 (yellow). Loops 1-3
and other atom colors are as in (A). (C) Binding pocket for the EA lesion is shown with Van der Waals
surfaces for protein residues (gray spheres) and EA lesion (cyan sphere). (D) Disrupted binding pocket in
lower-affinity AAG with Van der Waals surfaces colored as in (B). (E) Electrostatic representation of
A79AAG-EA:T calculated in the absence of DNA where surfaces in blue are more positive, those in red
more negative, and those in white are near neutral. The position of Tyr162 is denoted with a yellow star.
(F) Electrostatic representation of lower-affinity AAG with colors and symbols as in (E). (G) Same
depiction as in (E) but with substrate DNA modeled (orange cartoon). (H) Same electrostatic depiction as
in (F) aligned and superimposed with a cartoon model (purple) of A79AAG-EA:T. Disordered regions
that affect electrostatic potential are circled and represent the same loops as in (A) and (B).
80
B
A
is 3
Tr162,-
Tyr162
2
2
c3
Leul8O
Leu180
Asn169
Cys167
Asn169 Cys167
Glul25
Glu125
Tr159
Tyr127 -
Tyr127 y'
Tyrl59
His136
Ala134E
E
F
G
H
A
d4
81
Figure 111.8. Wall-eyed stereoviews of electron density in the lower-affinity A79AAG structure. (A)
2F0 -F, electron density omit map (blue mesh) contoured at la around Vall06-Glyl19 shows
representative electron density. (B) 2F0 -F, electron density omit map (blue mesh) contoured at 10 shows
the region from Val158-Tyrl65. This Tyr162 loop is flexible, displaying broken electron density and
higher B-factors (2-fold higher than the chain average; this same loop in the high-affinity pseudo-duplex
EC structure shows average B-factors). The absence of these residues results in the appearance of positive
difference electron density, while refinement in the presence of the modeled residues does not yield
negative difference electron density. Thus, these residues are included in the model as depicted.
Non-carbon atoms are colored as in Figure 111.2.
A
B
82
Figure 111.9. Proposal for how AAG can recognize DNA with two different affinities. The
electrostatic representations from Figures III.7E and 1II.7F are displayed in (A) and (B), for
A79AAG-EA:T and lower-affinity A79AAG, respectively, with the same coloring and symbols as in
Figure 111.7. The orientation of the molecules start as in Figure 111.7, and are then rotated 1200
counterclockwise (in two 60* steps) along the vertical axis such that the continuous positive surface can
be visualized. (C) Cartoon depiction of the search on DNA by AAG where blue and red represent
positively and negatively charged surfaces respectively. Relevant steps are labeled and a lesion base is
denoted as a green line. The AAG catalysis complex is a darker shade of blue to represent the more
ordered positive surface visualized crystallographically ((A) vs. (B) above).
A
Ao
B
C
Rotation-coupled sliding
Hopping
OW
Lesion
Searching
Recognition
Lesion
Catalysis
83
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85
86
Chapter 4. Crystallographic Evidence for Drastic Conformational Changes
in the Active Site of a Flavin-Dependent N-hydroxylase
A version of this chapter will be submitted for publication using the same title and with the
following author list:
Jeremy W. Setser, John R. Heemstra Jr.2 , Christopher T. Walsh 2 , and Catherine L. Drennani,4
Departments of 'Chemistry, 3Biology, and the 4Howard Hughes Medical Institute,
Massachusetts Institute of Technology, 77 Massachusetts Avenue, Cambridge, MA 02139 USA
2
Department of Biological Chemistry and Molecular Pharmacology, Harvard Medical School,
240 Longwood Avenue, Boston, MA 02115 USA
IV.I
SUMMARY
The soil actinomycete Kutzneria sp. 744 produces a class of highly decorated hexadepsipeptides,
which represent a new chemical scaffold that has both antimicrobial and antifungal properties.
These natural products, known as kutznerides, are created via nonribosomal peptide synthesis
using various derivatized amino acids. The piperazic acid moiety contained in the kutzneride
scaffold, which is vital for its antibiotic activity, has been shown to derive from the hydroxylated
product of L-ornithine: L-N-hydroxy-ornithine. The production of this hydroxylated species is
catalyzed by the action of an FAD- and NAD(P)H-dependent N-hydroxylase known as KtzI. We
have been able to structurally characterize KtzI in several states along its catalytic path, and by
pairing these snapshots with the biochemical and structural data already available for this
enzyme-class, we propose a structurally-based reaction mechanism that includes novel
conformational changes of both the protein backbone and the flavin cofactor. Further, we were
able recapitulate these conformational changes in the protein crystal, displaying their chemical
competence and catalytic relevancy. Our series of structures, with corroborating biochemical and
spectroscopic data collected by us and others, affords mechanistic insight into this relatively new
class of flavin-dependent hydroxylases, and adds another layer to the complexity of flavoenyzmes.
87
IVII
INTRODUCTION
Natural products and their derivatives are vital for human health, as they comprise over
one-third of all FDA-approved small-molecule drugs (1). Many of these compounds are
antibiotics that are biosynthesized by microbes via nonribosomal peptide synthesis (NRPS)
pathways (2). NRPS pathways are designed such that various enzymes act in an assembly-line
fashion to build up peptide chains using a broad range of both canonical and derivatized amino
acids. Even with the chemical diversity present in currently available antibiotics, which is largely
generated from the variety of permutations available in NRPS and similar systems, there exists a
pressing need for new therapeutic candidates to combat drug-resistant infections (2). Therefore,
finding new molecular scaffolds with therapeutic benefit, and elucidating the biosynthetic
pathways necessary to construct such compounds, is imperative for human health and
well-being.
Kutznerides are a recently discovered class of antifungal antimicrobials produced by the
soil actinomycete Kutzneria sp. 744 (3, 4). These natural products are highly-decorated, cyclic
hexadepsipeptides (Fig. IV.la), which are constructed via NRPS. The gene cluster encoding this
NRPS pathway has been elucidated (Fig. IV.lc (5)) and the functions of many of its biosynthetic
components have been demonstrated in vitro (5-11). One of these enzymes, KtzI (Fig. IV.lc in
red), was originally annotated as a 47-kDa flavin-dependent lysine/ornithine N-monooxygenase
(hydroxylase) (5), and this proposed activity was further investigated biochemically (7).
KtzI was found to use a non-covalently bound FAD cofactor, reducing equivalents from
NADPH or NADH (albeit less efficiently), and molecular oxygen to install a hydroxyl group on
the side chain nitrogen of L-ornithine (L-orn), producing L-N 5hydroxy-ornithine (Fig. IV.lb), thus
making KtzI a "stand-alone" NRPS enzyme. Binding of substrate analogue L-lysine causes
oxidation of the NADPH cofactor, without subsequent production of L-hydroxy-lysine,
effectively uncoupling the reaction (7). This specificity for substrate, and the cofactor and
cosubstrate usage, is similar to previously characterized flavin N-hydroxylases (12-23).
However, KtzI does differ from these other systems in the fate of its hydroxylated product.
The hydroxyornithine product of KtzI is ultimately incorporated into the kutzneride
scaffold as the N-N bond-containing piperazic acid moiety (7) (Fig. IV. 1), which has been shown
to be crucial for the antibiotic activity of these molecules (3). The intramolecular cyclization
88
necessary to reach this final structure, that is the creation of a bond between N2 and N of
hydroxyornithine, is likely preceded by some further activation of the hydroxylamine, but no
candidate enzymes have yet been established. In any case, the hydroxylation catalyzed by KtzI is
not retained in the final piperazic acid product, and thus is known as a 'cryptic' modification.
This use of cryptic N-hydroxylation for antibiotic biosynthesis is a departure from the previously
characterized
members of the N-hydroxylase
family.
All other lysine and ornithine
N-hydroxylases investigated to date have their hydroxy-modifications carried on to the final
product, where these moieties are used as ligands in iron-scavenging molecules called
siderophores (12-23). This divergence in the overall role of the N-hydroxylating protein has no
observable effect on the chemistry involved, however, as these enzymes all catalyze the creation
of a primary hydroxylamine product. The L-orn-specific N-monooxygenases, which have been
characterized in more detail, also appear to follow a common kinetic mechanism.
KtzI shares high sequence homology (33% identity) with the L-orn N-hydroxylases from
Pseudomonasaeruginosa(PvdA) and Aspergillusfumigatus(SidA) (Fig. IV.2), which have been
characterized in detail both biochemically (7, 15-17, 20, 21, 24-29) and structurally (29, 30). In
the biochemical studies, PvdA and SidA were proposed to follow a similar reaction mechanism
as outlined in Figure IV.3. Common to most flavin monooxygenases, the oxidized FAD cofactor
is reduced by hydride transfer from the C4-pro-R position of the NADPH nicotinamide to the N5
position of the flavin isoalloxazine ring, completing what is known as the reductive half of the
reaction (Fig. IV.3, 143). The oxidative half of the reaction begins when molecular oxygen adds
to the reduced flavin species, creating the highly reactive C4a-(hydro)peroxy intermediate (Fig.
IV.3, 4b and 5). Hydroxylation occurs through nucleophilic attack by the bound substrate (here
the side chain amine of L-orn) on the activated flavin-hydroperoxide intermediate, producing
both the hydroxylated product and a hydroxy-FAD intermediate (Fig. IV.3, 546). The oxidized
flavin species is regenerated through the loss of water, and that with the concomitant dissociation
of end products completes the catalytic cycle (Fig. IV.3, 6-1). If the hydroperoxy-flavin
intermediate is not sufficiently protected or used improperly, it is lost as the toxic byproduct
hydrogen peroxide, thus uncoupling the reductive and oxidative halves of the reaction and
wasting the reducing equivalents provided by NADPH (Fig. IV.3, 'uncoupling'). Even though
the N-hydroxylases follow this canonical mechanism, they diverge in other specifics and create
89
what is effectively a hybrid between two flavin monooxygenase archetypes known as "cautious"
and "bold" monooxygenases.
The "cautious" (or Class A) monooxygenases, which are typified by the extensively
studied para-hydroxybenzoate hydroxylase
(PHBH; reviewed in (31-33)), have a stringent
substrate-specificity and also maintain tight regulation over the reductive half of their reaction
cycle (Fig. IV.3, 143). In these enzymes, the rate of FAD reduction is intimately linked to
substrate binding, such that the presence of the hydroxylatable target greatly enhances the
reaction rate, upwards of 10 5-fold for PHBH (34). This control point significantly decreases the
risk of uncoupling (Fig. IV.3, 'uncoupling'), as the addition of 02 to reduced flavin, yielding the
highly reactive C4a-hydroxperoxy-flavin (Fig. IV.3, 4b, 5), can only occur when the substrate is
present to accept its hydroxy-modification. On the other hand, in "bold" (or Class B)
monooxygenases, such as those of the Baeyer-Villiger monooxygenase (BVMO) and flavin
monooxygenase (FMO) enzyme families, the reduction of FAD by NADPH occurs just as well
with or without bound substrate, and thus they must utilize a completely different tactic to
protect against uncoupling. In these systems, the spent NADP' cofactor remains bound to the
protein
throughout
the
C4a-(hydro)peroxy-flavin
reaction
cycle,
protecting
both
the
reduced-
species from being quenched (reviewed in (31,
and
the
32)). "Bold"
monooxygenases also tend to have little substrate specificity, illustrated by the over 200 known
compounds that the mammalian FMOs will hydroxylate (35).
The N-monooxygenases, although formally grouped with the Class B monooxygenases
(32), have been found to carry traits from both classes described above. They have the narrow
substrate specificity of the "cautious" enzymes, but show no sign of substrate-assisted reduction
of FAD and remain bound to NADP* throughout catalysis (15-17), which are both fundamental
characteristics of "bold" monooxygenases. Some structural insight into how catalysis is
controlled by these hybrid-monooxygenases has been provided by a series of recent crystal
structures (29, 30), which describe how cofactors and substrate are bound in the active site. In
this study, we build upon this structural knowledge by characterizing the N-hydroxylase KtzI in
never-before-seen states of this enzyme class, including the anaerobically reconstituted,
"pre-turnover" complex; and a view of the oxidized enzyme that suggests a drastic
conformational change, with a novel flavin movement, takes place. These disparate states were
shown to be chemically interchangeable in crystallo, demonstrating the chemical competence of
90
the conformational changes involved, and suggesting that these rearrangements are, in fact,
catalytically relevant. By combining our data with all other biochemical and structural data for
L-orn N-hydroxylases, we are able to propose a structurally-based reaction mechanism that
represents a paradigm shift for a flavin-dependent monooxygenase system.
IV.III RESULTS
The L-ornithine (L-orn) N-hydroxylase from Kutzneria sp. 744, KtzI, was structurally
characterized by X-ray crystallography, yielding over 30 structures under various conditions that
provide six unique snapshots of the enzyme (Table IV.1). The initial structure of KtzI was
obtained by reconstituting the purified protein with FAD and NADPH in an anaerobic
environment, followed by crystallization in NaBr-containing conditions. This structure, and all
other structures herein, is abbreviated to signify the molecules to which KtzI is bound and the
presumed redox state of its cofactors. Thus, the initial structure of this enzyme is denoted KtzIFADred-NADP'-Br, as it binds reduced FAD as evidenced by its colorless appearance (Fig.
IV.4a), oxidized NADP' due to the hydride transfer necessary to create the FADred species, and
a bromide ion in the substrate-binding pocket. This structure was determined to 2.4
A resolution
by molecular replacement using the protein coordinates from the functional homologue of KtzI
from Pseudomonas aeruginosa, PvdA (PDB ID 3S5W (30)). Two additional structures under
NaBr-containing conditions were obtained where the enzyme was reconstituted with FAD and
NADPH aerobically, and then either kept aerobic throughout (KtzI-FADox-Br; 2.1
A)
or
re-reduced after crystal growth with sodium dithionite under anaerobic conditions (KtzI-FADoxred-NADP'-Br; 2.6
A).
The oxidized flavin cofactor (FADox, yellow) is readily visible in the
KtzI-FADox-Br crystals (Fig. IV.4b), confirming the assignment of its redox state. The transition
from FADox (yellow; Fig. IV.4b) to FADred (colorless; Fig. IV.4a) (abbreviated FADox-red)
was readily apparent in the dithionite-reduced crystals. It is possible this 're-reduced' structure
has NADPH instead of NADP' bound to the enzyme, due to the chemical reduction by
dithionite, however this cofactor was found to bind in an identical manner to that of its
anaerobically reconstituted counterpart, KtzI-FADred-NADP*-Br (as detailed below), and thus
we have named it consistently for simplification. It was determined empirically that the
91
NaBr-containing condition occluded substrate binding due to a competing bromide ion, and thus
an alternative crystallization precipitant was pursued.
The reconstituted enzyme was found to crystallize using a replacement salt in the
precipitant (KSCN), which afforded substrate binding, and three additional structures. Two
structures were determined where KtzI was reconstituted with FAD, NADPH, and L-orn under
anaerobic conditions, and then either kept anaerobic (KtzI-FADred-NADP'-L-orn; 2.2
A;)
or
allowed to equilibrate with atmospheric oxygen, thus re-oxidizing the originally reduced crystals
(KtzI-FADred-ox-NADP -L-orn; 2.7
A).
The change from colorless (FADred; Fig. IV.4a) to
yellow (FADox; Fig. IV.4c) (abbreviated FADred-ox) was used to inform the assignment of the
flavin cofactor's redox state. One final structure was determined by following the same
reconstitution
protocol
above,
but
under
aerobic
conditions
throughout
(KtzI-FADox-NADP+-L-orn; 2.4 A), providing off-yellow crystals (Fig. IV.4c). These oxidized
structures are proposed to bind NADP+ due to this cofactor binding in a conformation that would
not (and does not) provide reduction of FAD (detailed below). The protein coordinates from the
initial refined model of KtzI (KtzI-FADred-NADP'-Br) were used as a search model in MR or
for rigid body refinement to solve all subsequent structures. Data processing and refinement
statistics can be found in Tables IV.2-IV.7.
Quaternarystructure
Even though KtzI was crystallized under a wide variety of conditions, it was found to
adopt the same homotetrameric assembly in each (Fig. IV.5a). Extensive interfaces exist between
each of the protomers such that ~25% of the available surface area is buried by these
interactions. These interfaces are conserved in the structures of the KtzI homologues PvdA (30)
and SidA (29), and the same tetramer is generated in these structures by crystal symmetry (Fig.
IV.6a). The buried surface areas are not quite as extensive for PvdA (17% buried) and SidA
(16% buried), largely due to a looser association at one of the interfaces (Fig. IV.6b). Although
the secondary structure at this interface is similar in all three homologues, the sequence
conservation in this region is relatively poor between KtzI and PvdA/SidA (residues 230-235,
260-270, 325-330 (KtzI numbering) in Fig. IV.2), and the arrangement of residues in KtzI causes
a helix to rearrange, yielding a clamping effect (Fig. IV.6b). In particular, the aforementioned
helix (residues 260-270 in KtzI) moves ~4-6
92
A closer
to its adjacent protomer, which allows a
cross-protomer hydrogen bond from Tyr270 to the 2'-phosphate of the NADP cofactor (Fig.
IV.6b). Given that this helix movement is exclusive to KtzI, the monomeric units of SidA and
PvdA are more related structurally to one another (rmsd ~1.1
A),
A),
than to our enzyme (rmsd ~1.5
despite the fact that overall sequence conservation between the proteins is not markedly
different (37% identity between SidA and PvdA vs. 33% between SidA/PvdA and KtzI). Indeed,
the sequence and structural conservation between all three proteins is apparent in the active site.
Active site of reduced KtzI
The fully-liganded, anaerobic complex of KtzI, KtzI-FADred-NADP'-L-orn, depicts this
enzyme in its reduced, "pre-turnover" state (Fig. IV.3, 4a and Figs. IV.5b and IV.5e). The FAD
and NADP cofactors are bound in an elongated conformation with the nicotinamide of NADP*
stacking on the Re-face of the flavin isoalloxazine ring. The nicotinamide cofactor is not in a
position to reduce the flavin, as its reactive C4 carbon is pointed away from the site of reduction
(N5 of the isoalloxazine ring). Instead, the carbonyl oxygen of the nicotinamide is oriented
toward this N5 position by a conserved glutamate residue (Glu212 in KtzI, Fig. IV.2), allowing
the carbonyl oxygen to hydrogen bond with the reduced N5-H group of FADred (Fig. IV.5b).
The flavin isoalloxazine ring is sandwiched between His51 and NADP', and is found to adopt its
bent or "butterfly" conformation (Fig. IV.5b and Fig. IV.7). There is evidence from structural
(36) and computational (37, 38) studies that this bend signals the fully reduced state of the
cofactor, which would be appropriate in this NADPH-reduced, anaerobic context. However, we
find bent isoalloxazine rings in all the structures presented herein, including those from formally
oxidized species where the flavin is surrounded by a completely different protein environment
(described in detail below), and thus believe it is unlikely that this phenomenon is chemically- or
environmentally-induced alone. It has been established for the flavin-containing reductase NrdI
that interaction with the photoelectrons produced during X-ray exposure can reduce the flavin
isoalloxazine ring, generating the bent conformation of the cofactor (39), and we propose that
this is likely to be a contributing factor in our structures, especially those from oxidized
preparations.
The amino acid substrate L-orn is specifically recognized in the active site by hydrogen
bonds to its carboxy and amino moieties, as well as to its sidechain amine (N5 ), by conserved
lysine, serine, and asparagine residues (Lys67, Asn245, Asn275, and Ser406 in KtzI; Fig. IV.5b
93
and Fig. IV.2). The carbonyl of Asn275 further hydrogen bonds with the 3'-OH of the
nicotinamide ribose of NADP* (Fig. IV.5b). The positioning of L-orn is such that the site of
hydroxylation (N5) is aligned with the C4a position of the isoalloxazine at a distance (5.9
A) that
would be amenable for catalysis after the addition of oxygen and subsequent creation of the
reactive C4a-hydroperoxy intermediate (Fig. IV.3, 5 and Fig. IV.5b, red dashed line). This highly
reactive center must be protected from bulk solvent, and this role is filled on one side of the
active site by the NADP' cofactor and a protein loop containing Asn275, Tyr276, and Ser277
('Tyr-loop'; Fig. IV.5e) and on the other by a neighboring protomer (Fig. IV.5b and Fig. IV.6c,
colored in wheat). Removal of these contacts on either side would result in an open active site
(Fig. IV.5e and Figs. IV.6c and IV.6d).
The reduced, pre-turnover state of KtzI was also crystallized wherein the substrate, L-om,
is replaced by a bromide ion from the precipitant solution (>1.0 M NaBr), and this structure is
denoted as KtzI-FADred-NADP'-Br (Fig. IV.8a). Even when reconstituted with a high
concentration of L-orn (31.8 mM), the bromide ion remained bound in the active site, and
bromides were found to occupy other sites all over the protein. However, these ions have no real
effect on the protein fold (rmsd = 0.3
A vs. KtzI-FADred-NADP'-L-orn),
nor on the arrangement
of cofactors and crucial active site residues (Fig. IV.8a vs. Fig. IV.5b). Further, the reduced
active sites of all the structurally characterized L-orn N-hydroxylases, save some minimal
fluctuations, are found to adopt identical conformations (PvdA: Fig. IV.8b (30)) and SidA: Fig.
IV.8c (29)) where the protective role served by: NADP', the Tyr-loop (Tyr285 in PvdA and
Tyr324 in SidA), and an adjacent protomer (Fig. IV.6), are all conserved.
Active site of oxidized KtzI
As mentioned above, KtzI was found to adopt the same homotetrameric assembly in each
structure, with minimal deviations between the respective states (rmsd = 0.1-0.3
A).
However,
one structure of this enzyme, that is where it was reconstituted aerobically with FAD and
NADPH and then crystallized in NaBr-containing conditions (denoted KtzI-FADox-Br), is most
different from the others with an rmsd of ~0.6
A. This divergence
is concentrated in the active
site where a drastic structural rearrangement has taken place. In this state, NADP* has vacated
the active site, Arg104 has swung in to hydrogen bond with Glu212, the isoalloxazine ring of
FAD has 'flapped' completely across the active site, and the Tyr-loop has inserted into the active
94
site interior, where Tyr276 stacks with the new conformation of the flavin ring (Figs. IV.5c and
IV.5f). The absence of bound NADP(H) is quite surprising as a large excess (20-fold vs. protein)
of NADPH was used during reconstitution. The conformational change from FADred to FADox
in KtzI shifts the isoalloxazine ring 6.5
A (N5-N5)
and rotates the angle between the flavin ring
and its ribityl tail by ~1370 (Fig. IV.5h), which is quite different from the change observed for
the prototypic Class A flavin hydroxylase, para-hydroxybenzoate hydroxylase (PHBH) (Fig.
IV.5i). The insertion of the Tyr-loop into the active site moves the Ca carbon of Tyr276 4.0 A,
such that this residue is now sandwiching the isoalloxazine ring with the help of His51, which
has rotated slightly to remain parallel with the flavin cofactor (Fig. IV.5c). The absence of
NADP' and L-orn has caused other conserved residues: Asn245, Asn275, Ser277, and Ser406, to
adopt new conformations with their sidechains pointed away from the active site (Figs. IV.5c and
IV.5f). These drastic rearrangements greatly alter the effective surface of the protein active site,
such that the isoalloxazine moiety is now open to solvent (Fig. IV.5f). Aside from this active site
reorganization, the rest of the protein topology remains largely the same (rmsd ~0.6
A),
with
most residues adopting identical conformations. KtzI was also reconstituted aerobically with
FAD, NADPH, and L-orn, and crystallized under KSCN-containing conditions, which yielded
yet another independent snapshot of this state of the enzyme.
Following the same aerobic reconstitution that produced the KtzI-FADox-Br structure,
but with the addition of L-orn and the use of KSCN in place of NaBr, an amalgamation of the
fully-liganded,
anaerobic complex (KtzI-FADred-NADP'-L-orn)
and KtzI-FADox-Br was
produced. This structure, denoted as KtzI-FADox-NADP*-L-orn (Figs. IV.5d and IV.5g), is
similar to the fully-liganded, anaerobic complex, in that it has FAD, NADP', and L-orn bound to
the protein and the active site residues adopt the same conformation as in the pre-turnover
complex (Fig. IV.5d vs. Fig. IV.5b). However, the bound flavin adopts the 'flapped',
FADox-conformation observed in KtzI-FADox-Br (Fig. IV.5d vs. Fig. IV.5c). This arrangement
appears as if the flavin of fully-liganded anaerobic structure was simply replaced with that of the
flapped, oxidized conformation, such that the NADP' moiety now stacks with the Si-face of the
isoalloxazine ring (Figs. IV.5d and IV.5g).
Both snapshots of the oxidized KtzI enzyme propose a completely novel conformational
change for a protein-bound flavin molecule, however, these structures are also quite different
from one another (Fig. IV.5c vs. Fig. IV.5d). This dissimilarity was quite surprising as the
95
method employed to crystallize these structures was identical aside from the salt used in the
precipitant (NaBr vs. KSCN) and a relatively modest pH change (7.5 vs. 8.5). This unanticipated
discrepancy, combined with the lack of precedence for either depiction, concerned us that some
crystallization artifact may have trapped non-relevant or dead-end states. In an effort to
understand the chemical relevancy of these respective states, manipulations were employed to
observe whether the supposed conformational changes could be recapitulated in the protein
crystal.
In crystallo conformational changes
In KtzI-FADox-Br, the 'flapped-flavin' state is accompanied by the departure of NADP'
and the insertion of a protein loop, resulting in the isoalloxazine ring being exposed to solvent.
Bromide ions, including the substrate-occluding ion observed in KtzI-FADred-NADP*-Br, are
also present in the active site of KtzI-FADox-Br (Fig. IV.9a), raising questions about how these
ions may affect, or possibly cause, this unusual structure. In an effort to establish the chemical
relevancy of this state, crystals were grown as usual and then subjected to chemical reduction by
NADPH and sodium dithionite under anaerobic conditions. The crystals showed a definitive
color change from yellow (oxidized flavin; Fig. IV.4b) to colorless (reduced flavin; Fig. IV.4a)
when either reductant was used, but structural characterization could only be carried out on the
sodium dithionite-reduced crystals. This limitation was due to the fact that a longer incubation
period (30 min) was required to fully reduce crystals with NADPH as compared to sodium
dithionite (<10 sec), and this longer time frame left the crystal too deteriorated for useful data
collection.
Amazingly, soaking the oxidized, KtzI-FADox-Br crystals with sodium dithionite under
anaerobic conditions provided a structure, denoted as KtzI-FADox-red-NADP*-Br, which is
identical
(rmsd
=
(KtzI-FADred-NADP'-Br)
0.15
A)
to
its
anaerobically
reconstituted
(Fig. IV.8d vs. Fig. IV.8a). This finding
counterpart
means that the
conformational changes necessary to turn KtzI-FADox-Br into its reduced, and NADP-bound
species, that is the 'flapping' of its isoalloxazine ring across the active site with concomitant
evacuation of the Tyr-loop and re-binding of the NADP' cofactor, were all able to occur in the
crystal.
96
In KtzI-FADox-NADP*-L-orn, the flapped, FADox conformation is observed such that
the isoalloxazine faces the protein exterior, but NADP', L-orn, and all active site residues appear
as in the fully-liganded anaerobic complex, creating the effect of an NADP cofactor slipping
behind the flapped-flavin conformation (Figs. IV.5d and IV.5g). Attempts to reduce these
crystals, even upon long incubations with NADPH or dithionite, proved futile as no color-change
was observed. However, we examined whether the opposite chemical transformation could occur
in the crystal, that is from the fully-reduced, anaerobic species (Fig. IV.5b) to its oxidized
counterpart (Fig. IV.5d) through the addition of oxygen.
Crystals of the KtzI-FADred-NADP+-L-orn complex were grown anaerobically and then
equilibrated aerobically, allowing the flavin cofactor to oxidize. This re-oxidized structure
(denoted KtzI-FADred-ox-NADP*-L-orn), is identical (rmsd = 0.13
A) to
that of the aerobically
grown, KtzI-FADox-NADP+-L-orn state (Fig. IV.9b vs. Fig. IV.5d). Excitingly, this observation
means that upon the addition of oxygen to the fully-liganded, anaerobic complex (Fig. IV.5b),
the FAD isoalloxazine was able to travel across the active site from its FADred state to arrive at
the FADox position observed in the aerobically reconstituted KtzI structures (Figs. IV.5d, IV.5g,
and IV.5h), all while contained in the crystal lattice. Clashing of the FAD and NADP cofactors
would likely occur in any FADred to FADox conformational change in KtzI, and thus the
nicotinamide cofactor would be expected to depart during this re-oxidation process, signaling
that more dynamic changes take place in between the equilibrium states that we have observed
crystallographically.
IV.IV DISCUSSION
The flavin-dependent N-hydroxylases are a class of enzymes that have been studied
almost exclusively in the context of their role in generating iron-chelating siderophores (12-23).
The L-orn-specific N-monooxygenase from Kutzneria sp. 744 (KtzI) does not follow this trend,
however, as it instead provides a building block for the biosynthesis of a new class of antifungal
antimicrobials called kutznerides (7) (Fig. IV.1a). Regardless of their overarching role, the
N-hydroxylases all catalyze the same reaction; using FAD, NAD(P)H, and molecular oxygen to
attach a hydroxyl group on the primary-amino side chain of their substrate (Fig. IV.lb). Further,
the L-orn N-hydroxylases from Pseudomonas aeruginosa (PvdA) and Aspergillus fumigatus
97
(SidA), which share high sequence homology with KtzI (Fig. IV.2), have been observed to
follow a common kinetic mechanism (15-17) (Fig. IV.3). Preliminary structural interpretation of
this kinetic mechanism has been made possible by recent crystal structures of PvdA (30) and
SidA (29). Here, we add to the structural reaction coordinate by providing novel depictions of
this enzyme class, showing KtzI in its anaerobically-reduced, fully-liganded, "pre-turnover"
state, and in two oxidized states that suggest a series of conformational changes occur. These
proposed rearrangements were shown to be chemically competent in crystallo, indicating that
they are an important part of the catalytic cycle for KtzI, and more generally the N-hydroxylase
family of enzymes.
The anaerobically-reduced KtzI-FADred-NADP'-L-orn structure (Figs. IV.5b and IV.5e),
and an accompanying depiction where a bromide ion from crystallization has displaced the
substrate (KtzI-FADred-NADP*-Br; Fig. IV.8a), are very similar to the aerobically-reduced
depictions of PvdA (30) (Fig. IV.8b) and SidA (29) (Fig. IV.8c). This similarity indicates that
reconstituting the enzyme in the absence of oxygen, as was done for KtzI, can be approximated
by chemical reduction of the aerobic state, and thus these structures can all be taken together as
independent validations of the pre-turnover state of this enzyme family. In this state, NADPH
has already transferred its hydride to the FAD moiety (Fig. IV.3, 2->3), such that FADred,
NADP*, and the L-orn substrate (or competing bromide ion) are bound in the active site. This
redox assignment can be confirmed by the bleached appearance of these crystals (Fig. IV.4a),
and the fact that the reactive C4 position of the nicotinamide moiety is pointed away from the
site of reduction (N5 of FAD; Fig. IV.5b), and thus is not positioned for hydride transfer. A
conserved glutamate residue (Glu212 in KtzI; Fig IV.2) instead poses the nicotinamide such that
its carbonyl oxygen can hydrogen bond with the newly-formed N5-H of reduced flavin (Fig.
IV.5b). The bound NADP' also makes up a large portion of the protein surface (Fig. IV.5e),
effectively sealing the active site with its presence. Therefore, in addition to its role in flavin
reduction, this cofactor acts to shield the FADred and FAD-OO(H) intermediates (Fig. IV.3, 3-5)
from bulk solvent, protecting these species from being quenched. Indeed, there is kinetic
evidence from PvdA (30) and SidA (15, 16) that NADP* remains bound throughout the reaction
cycle to perform this role. This protective function is further exemplified by the observation that
chemical reduction of SidA, followed by exposure to oxygen in the absence of NADP*, resulted
in immediate uncoupling through H20
98
2
production (15) (Fig. IV.3, 'uncoupling'), whereas the
reactive C4a-(hydro)peroxy intermediate is stabilized (in the absence of its hydroxylatable
target) by NADP' on the order of minutes in both SidA (15, 16) and PvdA (17). There is
evidence from kinetic isotope effects and computational studies that suggest the 2'-OH of the
ribose of NADP* may be directly responsible for stabilizing the flavin-peroxide intermediate,
and further, may act as the proton donor for creation of the C4a-hydroperoxy derivative (26).
Taking all these studies together, it is readily apparent that the nicotinamide cofactor is crucial to
the entirety of the reaction cycle, well beyond its initial task of reducing the flavin moiety, and
this multi-faceted involvement is, indeed, a hallmark of the Class B monooxygenase family (32).
In the N-hydroxylases, the protein scaffold acts as an additional barrier to quenching species.
As was mentioned for the structure of PvdA (30), if one considers the monomeric unit of
an N-hydroxylase, the active site is excluded from solvent by both NADP* (Fig. IV.5e) and the
bound substrate (Fig. IV.6d), which would be incongruent with the observation of a stabilized
flavin-hydroperoxide in the absence of L-om, as described above. Although the protein backbone
provides protection in the form of the conserved Tyr-loop (Fig. IV.5e), departure of NADP' or
L-orn would open the monomeric active site to solvent. This concern is alleviated, however, by
thinking of the protein in terms of the tetrameric assembly observed crystallographically (Fig.
IV.5a and Fig. IV.6a). The interfaces of this tetramer are quite extensive in all three homologues,
burying 16-25% of the available surface area, and one of these interfaces guards the active site
with its interaction (Fig. IV.5b, wheat-colored protomer and Fig. IV.6c). A tetramer is also
consistent with gel filtration chromatography data collected on SidA and PvdA (16, 21), alluding
to this species' relevance in solution. Therefore, it can be reasoned that the N-hydroxylase
tetramer is formed for the protection of reactive intermediates, and thus is vital for catalysis.
These enzymes, like those of the "cautious" monooxygenase family, also manage catalytic
specificity for their L-orn substrate, and this task is accomplished using a highly organized active
site.
L-om is secured in the binding pocket by hydrogen-bonds made from conserved residues
to all of its polar groups (Fig. IV.5b and Fig. IV.2), highlighting an environment designed to bind
this amino acid. As was shown in crystal structures of SidA, however, this site is also able to
accommodate L-lys and L-arg in a highly-similar binding mode (29). There is some evidence that
SidA can hydroxylate L-lyS to a certain degree (29), but studies have mostly found L-orn
N-hydroxylases to be highly specific for L-orn (7, 16, 20, 21, 24), and thus it is somewhat
99
surprising to find non-substrates binding in the active site. This discordance seems to indicate
that although different L-amino acids can bind similarly to L-orn, hydroxylation is controlled by
the angle and proximity of the amine group to the flavin-hydroperoxide, such that L-orn presents
the only species in prime position for catalysis. The inability of the L-orn N-monooxygenases to
hydroxyate L-lys effectively, with the large amount of uncoupling observed in the presence of
this amino acid, indicates that the extra methlyene unit of this molecule extends its amine side
chain too far into the active site for hydroxylation to occur, but resembles L-orn enough to trigger
oxidative uncoupling (Fig. IV.3, 'uncoupling'). Further insight into the oxidative half of the
reaction was provided by aerobic structures of KtzI.
When KtzI was reconstituted under aerobic conditions, and then crystallized using two
different precipitants, the result was two very distinct crystal structures (Figs. IV.5c and IV.5f vs.
Figs. IV.5d and IV.5g). Both states share an unprecedented flavin conformation, which we will
now refer to as the FADox(out) conformation, that is entirely different from the FADred 'in'
state. This 'out' conformation shifts the isoalloaxazine moiety completely across the active site,
such that it is now open to solvent (Figs. IV.5f and IV.5g). Aside from this similarity, the aerobic
structures are largely divergent: In KtzI-FADox-Br, the new flavin conformation is accompanied
by the dissociation of NADP', and the movement of conserved active site residues including
insertion of a 'Tyr-loop' (Asn275, Tyr276, Ser277) and swinging of Arg104 such that it takes the
place of NADP' by hydrogen bonding with Glu212 (Fig. IV.5c vs. Fig. IV.5b). In
KtzI-FADox-NADP'-L-orn, the 'flapped'
flavin resides in an active site that is largely
unchanged compared to the pre-turnover complex, with NADP', L-orn, and protein residues all
adopting similar conformations (Fig. IV.5d vs. Fig. IV.5b). This discrepancy led us to examine
the methods used to attain these two structures.
The most influential differences between the preparations of these dissimilar states
include the presence (KtzI-FADox-NADP*-L-orn) or absence (KtzI-FADox-Br) of substrate
during reconstitution, and the salt used in the precipitant solution (KSCN vs. NaBr).
Crystallization experiments using NaBr-containing conditions were performed with the same
reconstitution protocol used for the substrate-bound, KSCN-crystallized structures (i.e. with 31.8
mM L-om), but a competing bromide ion continually abrogated substrate binding (Fig. IV.9a),
likely due to its vast excess (-1 M vs. 31.8 mM). This lack of a direct comparison makes it
difficult to assess whether the presence or absence of substrate vs. bromide ions drives the
100
difference between these states. It is tempting to attribute the overarching difference between
these two snapshots to the presence or absence of a bound NADP', however, a structure of SidA
in the presence of FAD and L-orn (without NADP') adopted a similar conformation to the
reduced structures (Fig. IV.10, A (right) (29) vs. Fig. IV.5b), signaling that NADP' binding is
not the sole determining factor. Although the hydrogen bond switch from Glu2l2-NADP* (Fig.
IV.5b) to Glu2l2-ArglO4 (Fig. IV.5c) observed in KtzI, was also observed in the crystal
structures of reduced vs. oxidized states of SidA (Fig. IV.8c vs. Fig. IV.10, A (right)), no
precedence exists for the flavin conformational change suggested by our structures. Further, as
crystallographic characterization occurs on equilibrium states, our aerobic structures are likely
the end-product of some other intermediate state(s). Therefore, we felt it necessary to confirm the
chemical relevance of our structural snapshots through in crystallo characterization of KtzI.
KtzI-FADox-Br and KtzI-FADox-NADP'-L-orn crystals were subjected to anaerobic
reduction by NADPH (25 mM) and sodium dithionite (50 mM). In each case, reduction could be
monitored by eye as a color change from yellow (Fig. IV.4b) to clear (Fig. IV.4a), and this
change was only observed for KtzI-FADox-Br crystals. The KtzI-FADox-Br state (Figs. IV.5c
and IV.5f) was returned completely to its pre-turnover state by anaerobic reduction in crystallo
(Fig. IV.8d vs. IV.8a; rmsd = 0.15 A). This transformation displays the chemical competence of
this conformational change, and as reduction was elicited using both sodium dithionite and the
physiologically-relevant NADPH cofactor, provides evidence for the catalytic relevancy of this
dynamic motion. Indeed, sodium dithionite could be acting as a conduit for NADPH reduction of
FADox, as its reduction potential (-0.66 V) allows it to either reduce the leftover NADP' in the
crystallization drop (which then goes on to reduce FADox), or to reduce FADox directly. In any
case, the recapitulation of the reduced, pre-turnover state from this oxidized state using chemical
means provides compelling evidence for its general relevance. Further evidence for dynamic
motions being required for KtzI catalysis comes from the in crystallo re-oxidation of the
fully-liganded, anaerobic state.
KtzI-FADred-NADP'-L-orn crystals (Figs. IV.5b and IV.5e) equilibrated in atmospheric
oxygen fully recapitulated the structure of their aerobically reconstituted counterpart (Fig. IV.9b
vs. Fig. IV.5d; rmsd = 0.13
A).
This structure, which is named KtzI-FADred-ox-NADP'-L-orn,
began as an aerobic protein solution, was anaerobically reduced by NADPH and crystallized
anaerobically,
before finally being re-oxidized.
The enzyme has undergone
multiple
101
transformations during this process, and the snapshots we see are the end-products of these
dynamic motions. Indeed, by comparing the structures before (Fig. IV.5b) and after (Fig. IV.9b)
oxygen exposure, it becomes apparent that any movement of the flavin moiety to get from the
FADred to FADox(out) conformation would be sterically occluded by the bound NADP'. This
observation is compelling evidence for the existence of intermediate states as for these flavin
motions to occur, dissocation, and then reassociation, of NADP* would be required, which we
are unable to observe directly. Further, the L-orn bound in this structure is certainly not the same
as the L-orn bound initially, which would be expected to have been hydroxylated upon exposure
of the fully-loaded, pre-turnover enzyme to 02, but instead represents another molecule that has
re-bound sometime during the re-oxidation process. Given that this re-oxidized structure and its
aerobically reconstituted counterpart (Fig. IV.9b vs. Fig. IV.5d) are identical (rmsd = 0.13
A),
this same logic can be used to explain both of their unobserved intermediate states. Whether
these conformational changes between equilibrium states occur multiple times is unable to be
discerned from our studies; however, we believe our data make it clear that they must occur at
least once. Taken together, these observations lead directly to the question: why do these
conformational changes occur?
A conformational change in a flavin-dependent hydroxylase is not a new observation.
Indeed, "cautious" monooxygenases control faithful coupling of NADPH reduction to substrate
hydroxylation by movements of their flavin moiety (reviewed in (31, 32)). As mentioned
previously, "cautious" (or Class A) monooxygenases get their name from the fact that in these
enzymes, NADPH will reduce FAD only when substrate is bound, thus limiting oxidative
uncoupling (Fig. IV.3, 'uncoupling'). The structural basis for this phenomenon was first
proposed for para-hydroxybenzoate
hydroxylase (PHBH) (40), where substrate-coupled
reduction is controlled by a planar, hinge-like movement of the FAD isoalloxazine ring from the
protein interior ('in') to the protein exterior ('out') upon substrate binding (Fig. IV.5i), which
allows NADPH access to reduce FADox. Once reduction occurs, the flavin moves from this
'out' position back 'in' to protect its reduced N5-H from solvent. Although this precedence
exists for "cautious" enzymes, KtzI is the first Class B monooxygenase proposed to use flavin
conformational changes during catalysis, and these movements are even more drastic than those
of the Class A systems (Fig. IV.5h vs. Fig. IV.5i).
102
We know the conformational changes observed for KtzI are not correlated in the same
way as "cautious" enzymes, as L-orn N-hydroxylases, like "bold" monooxygenases, can just as
easily be reduced in the absence of substrate. Further, oxidized SidA and PvdA crystals (whose
structures resemble those seen for the reduced KtzI states) were reduced in crystallo without any
indication of a conformational change (29, 30), so flavin movement may not be strictly required
for the reduction step. To formulate the mechanistic implications of these conformational
changes, it is more pertinent to consider another trait N-hydroxylases share with "bold"
monooxygenases, that is, NADP' remains bound throughout the reaction cycle, and thus, only
dissociates after hydroxylation occurs. This observation means that there must be some signal for
NADP' to dissociate, such that another round of catalysis can occur. By combining our snapshots
of KtzI, with those of PvdA and SidA, we believe a structural reaction coordinate for Nhydroxylase enzymes (summarized in Fig. IV.l0) can be built that utilizes conformational
changes to control catalysis, which may provide further insight into the "bold" monooxygenase
family in general.
L-orn N-hydroxylases in their resting, oxidized flavin state, could exist in an equilibrium
between the FADox(out) conformation observed in KtzI-FADox-Br (with or without insertion of
the conserved Tyr-loop), and the FADox(in) of SidA-FADox-L-orn (Fig. IV.10, A). It may be
difficult to distinguish these two states spectroscopically, as the Tyr276-stacked, FADox(out)
conformation in our crystals of KtzI-FADox-Br (Fig. IV.10, A (left)) still retain the deep yellow
color of oxidized flavin (Fig. IV.4b), indicating that the signal is not significantly quenched or
altered. In contrast, the flavin signal does appear to be altered by NADP' stacking on the
FADox(out) conformation, as crystals of KtzI-FADox(out)-NADP'-L-orn (Fig. IV.10, E) take on
an off-yellow appearance (Fig. IV.4c). In any case, we have been able to show that our
KtzI-FADox-Br state can be transformed into its reduced counterpart, KtzI-FADred-NADP*-Br
(Fig. IV.10, B), by in crystallo reduction with NADPH, displaying the chemical competence of
the FADox(out)-Tyr-loop-inserted conformation. Therefore, it cannot be confirmed whether
either (or both) state(s) are physiologically relevant for NADPH reduction. A so-called 'sliding
mechanism' for NADPH reduction has been suggested for the Baeyer-Villiger monooxygenase
(BVMO) class of "bold" monooxygenases,
wherein the nicotinamide adopts different
conformations during and after reduction. This proposal is largely based on the observation of
varied binding modes of this cofactor in crystal structures (41, 42), and the indication from
103
kinetic studies that the reduction process happens in two phases, one of which is consistent with
an equilibration step for NADPH in the binding pocket (43). The sliding mechanism has also
been proposed for SidA (25), as biphasic reduction kinetics have been observed for both PvdA
(17) and SidA (15, 27), and there is evidence from a mutant of SidA (Ser257Ala) that
destabilizing a hydrogen-bonding interaction to the pyrophosphate of NADP', actually increases
the reduction rate, suggesting that a more dynamic nicotinamide ring is favorable for this step
(25). Although our data do not provide further information on this sliding mechanism directly,
the FADox(out) conformation we observe, and have shown to be chemically competent, adds an
extra layer to consider when thinking about the reduction step. Indeed, the FADox(out)
conformation may be a contributing factor to the complicated kinetics of reduction observed for
these enzymes. We are able to use structural information to comment more directly on the
equilibrium state following this transient reduction step.
After FAD reduction and NADP* binding, conserved residues have been observed to
rearrange in both KtzI (Fig. IV.10, A4B) and SidA (Fig. IV.10, A (right) vs. Fig. IV.8c), such
that Arg104 (Arg144) swings out of the NADP' binding, allowing Glu212 (Glu260) to hydrogen
bond to the carboxamide of NADP', orienting this cofactor to protect the N5-H of FADred.
Binding of L-orn to the pre-turnover, anaerobic state causes only minor rearrangements of
substrate-binding residues (Lys67, Asn245, Ser406), as our structures with bromide or substrate
bound in this site are fairly identical (rmsd = 0.3
A), and those of SidA without
substrate bound
look largely similar (29). The addition of molecular oxygen to this pre-turnover state, whether or
not substrate is bound, creates the reactive hydroperoxy-flavin (Fig. IV.3, 4b/5). This
transformation is also unlikely to cause any structural deviations, as the binding affinity for L-orn
has been shown to be similar in the reduced- vs. hydroperoxide-flavin forms of SidA (15), and
NADP' remains bound to both states (15-17), suggesting a similar active site architecture. Once
the hydroxyl group is picked up by L-orn, however, the enzyme must be recycled to its
resting-state in some way.
The hydroxylation step immediately precedes the departure of NADP' in biochemical
studies (15, 16, 30), and the structural transmission of this signal may be exemplified by the
post-turnover structure of PvdA (PvdA-FADox-NADP'-L-N 5OH-orn; Fig. IV.10, D). In this
structure, aerobic reconstitution of PvdA with FAD, NADPH, and L-orn (turnover conditions),
followed by crystallization, trapped the product complex in the crystal. The electron density for
104
the nicotinamide of NADP' is disordered in this state, such that this cofactor could not be
modeled effectively, indicating this moiety is dynamic in the product-bound state (30). We
propose that the act of taking L-orn (Fig. IV.10, C (inset)) to L-N 5OH-orn (Fig. IV.10, 1
'hydroxylation') causes a steric clash with NADP', which propagates to destabilize this cofactor
(Fig. IV.10, 2 'destabilization'). The destabilization of NADP' was not enough, however, to
actually dissociate this spent cofactor, as it remains bound in this structure (Fig. IV.10, D). This
lack of a clear departure signal allows us to propose that for NADP' to leave, the flavin
conformational change from the FADox(in) to FADox(out) state that we observe for KtzI, is
used to eject the oxidized nicotinamide cofactor in the last step of catalysis (Fig. IV.10, 3
'ejection'). During an interpolation between these two conformations, NADP' and FAD would
clash directly, and thus, FAD could act as a steric battering ram to eject NADP' from the active
site. This would not be the first time that a flavin has been proposed to act in a steric capacity, as
we recently proposed from structural and biochemical studies on the FAD-dependent
hydroxylase StaC, that this Class A enzyme uses the 'in' movement of its flavin moiety to
sterically-induce decarboxylation of its substrate molecule (44). As mentioned above, NADP'
ejection has been linked to the last step in the catalytic cycle of both PvdA (30) and SidA (15,
16) by pre-steady state kinetic studies, and the difficulty in deconvoluting the spectra for this
oxidative process in SidA (15), hints that something more complex than mere dissociation of
NADP' may occur. Therefore, with all the evidence presented for KtzI, a case has been made for
the 'flapping' of the flavin moiety to be an integral part of the catalytic cycle. Further evidence
for the relevance of the Tyr-loop and flavin conformational changes is provided by in crystallo
studies of the KtzI-FADox-NADP'-L-orn state.
The flavin moiety in the FADox(out)-NADP' bound conformation (Fig. IV.10, E) was
unable to be reduced by sodium dithionite or NADPH, indicating that the binding of NADP*
behind the FADox(out) conformation inhibits this reduction step (Fig. IV.10, E+C). Indeed,
reduction of the flavin, without subsequent movement inside the protein, would only be transient
in nature as it is completely solvent exposed (Fig. IV.5g). This observation allows us to propose
that NADP' is inhibiting reduction by blocking the conformational change from FADox(out) to
the 'in' position of FADred (Fig. IV.10, E+C). Further, insertion of the Tyr-loop, as seen for
the KtzI-FADox-Br (Fig. IV.10, A (left)), could function to disfavor re-binding of NADP',
guarding against this dead-end, inhibited state. As mentioned above, we were able to take
105
crystals of KtzI-FADox-Br back to the pre-turnover state by anaerobic reduction, signaling the
chemical competence of this state (Fig. IV.10, A4B), thus making it a good option to disfavor
inhibition by NADP'. The dead-end, FADox(out)-NADP' bound state (Fig. IV.10, E), was
completely recapitulated by exposure of the pre-turnover, anaerobic state to oxygen (Fig. IV.10,
C+E). This oxygen-dependent conformational change of the flavin from its 'in' to 'out'
position in crystallo, provides good evidence for the catalytic relevance of this movement, and
confirms that the motions necessary to carry out our proposal for the physical ejection of NADP'
by FAD are highly plausible.
We have observed drastic differences in the active site of a flavin-dependent
N-hydroxylase through a series of crystallographic snapshots. Many of these states, which were
observed for the first time in our study, were shown to be interchangeable in crystallo, signaling
both their chemical competence, and relevance towards understanding the catalytic mechanism
of N-hydroxylases. The conformational dynamics during catalysis proposed herein, represent a
new way of thinking about flavin-dependent enzymes, and considering the similarities between
N-hydroxylases and the other members of the Class B monooxygenases, some of these dynamic
motions could be more broadly applicable across the class. This applicability is especially true
when considering the 'ejection' of the spent NADP' cofactor, as a hallmark of Class B enzymes
is both their ability to stabilize intermediate states using this cofactor, and the necessity for its
dissociation before another round of catalysis can occur. We hope that our structural
observations will provide a new lense for further biochemical examination of these interesting
flavoenzymes.
IV.V
MATERIALS AND METHODS
Protein expression andpurification
The ktzl gene was cloned into a pET28a vector (Novagen) and the resulting N-terminal
hexahistidine construct was transformed into BL21(DE3) cells (Invitrogen) as described
previously (7). For protein overproduction, Luria-Bertani medium (LB) (100 mL supplemented
with 50 Rg/mL kanamycin) was inoculated using a glycerol stock of the expression strain and
grown overnight at 30*C and 200 RPM. Four l-L cultures of LB containing 50 [tg/mL
kanamycin were each inoculated with 20 mL of the overnight culture and incubated at 25*C and
200 RPM until the OD,00=0.5-0.6, at which point protein expression was induced by the addition
106
of 100 gM IPTG. The cells were grown for 16 hr post-induction at 15*C and 200 RPM and
harvested by centrifugation (6000 x g for 10 min). Cells were resuspended in lysis buffer (25
mM Tris-HCl pH 8.0, 500 mM NaCl) and lysed by sonication. The lysate was clarified by
centrifugation (75,000 x g for 35 min) and incubated with 5 mL of Ni-NTA slurry that had been
pre-equilibrated in lysis buffer. This mixture was agitated at 4*C for 1 hr before being loaded
into a column for purification. The resin was washed with lysis buffer supplemented with 0 mM
(60 mL), 5 mM (60 mL), and 25 mM (60 mL) imidazole. KtzI was eluted with lysis buffer
containing 200 mM (10 mL) and 500 mM (5 mL) imidazole and the resulting eluate was
concentrated to -1 mL in a 10K MWCO filter (Millipore) by centrifugation (4500 x g for 10 min
intervals). The concentrated sample was dialyzed against protein storage buffer (20 mM TrisHCl pH 8.0, 80 mM NaCl, 10% (v/v) glycerol) in a 10K MWCO dialysis cassette (Thermo
Scientific) over a period of 16 hr with three buffer exchanges. The dialyzed protein solution (-20
mg/mL by UV absorption at 280 nm;
E-cai
= 48485 M-'cm-') was flash frozen in liquid nitrogen in
15 pL aliquots and stored at -80*C. The final KtzI protein construct has 21 non-native amino
acids (including the hexahistidine tag) at its N-terminus, followed by the wild-type sequence
beginning with a valine residue at position 3 (Val3) (Fig. IV.2).
Reconstitution of KtzI for crystallization
KtzI (10 mg/mL or -212 VM) was reconstituted with FAD (212 pM) and NADPH
(4.24 mM) for all crystallization trials. To obtain substrate-bound structures, L-ornithine
(31.8 mM) was added to this mixture. FAD, NADPH, and L-ornithine (all from Sigma-Aldrich)
were diluted such that the final buffer composition of the reconstituted sample prior to
crystallization was 20 mM Tris-HCl pH 8.0, 80 mM NaCl, and 5% (v/v) glycerol. For anaerobic
reconstitution, frozen tubes of KtzI protein and aliquots of FAD, NADPH, and L-ornithine
powder were degassed and brought into an anaerobic chamber (95% Argon, 5% Hydrogen; COY
Laboratory Products, Inc.) prior to dilution with anaerobic buffer. All solutions for anaerobic
manipulations had argon bubbled through them to remove oxygen. The final reconstituted
samples were incubated in a cold block (-4*C) for 1 hr before crystallization. Under anaerobic
conditions, the characteristic color change from oxidized (FADox, yellow) to reduced (FADred,
colorless) flavin could be observed in the sample, indicating hydride transfer from NADPH to
FAD had occurred.
107
Crystallizationof KtzI
Initial crystallization conditions were found using the Phoenix Liquid Handling System
(Art Robbins Instruments) to mix 150 nL of reconstituted protein with 150 nL of precipitant in a
96-well, sitting-drop INTELLI-PLATE@ (Art Robbins Instruments) format. Crystals of
reconstituted KtzI were optimized using the hanging-drop vapor diffusion method at room
temperature by mixing 1 pL of reconstituted protein (containing 212 PM N-His6 tagged protein,
212 pM FAD, 4.24 mM NADPH, (31.8 mM L-ornithine for substrate-bound structures), 20 mM
Tris-HCl pH 8.0, 80 mM NaCl, and 5% (v/v) glycerol) with 1 pL of precipitant over a reservoir
of 0.5 mL of precipitant. KtzI crystallized with two precipitants: one of which afforded substrate
binding (0.4-0.7 M (aerobic) or 0.9-1.15 M (anaerobic) KSCN, 22-25% PEG 3350, 0.1 M
Bis-tris propane pH 8.5), while the other precluded substrate-binding due to a competing
bromide ion (0.9-1.2 M (aerobic) 1.2-1.4 M (anaerobic) NaBr, 22-25% PEG 3350, 0.1 M Bis-tris
propane pH 7.5). Data-quality crystals grew as colorless (anaerobic, Fig. IV.4a) or yellow
(aerobic, Fig. IV.4b and IV.4c) rods after 4-7 d with approximate dimensions of 70 x 70 x 200
pm. Crystals were cryoprotected with crystallization precipitant supplemented with 10-20% (v/v)
glycerol
(and
with
31.8
mM
L-orn
to
assure
full
substrate
occupancy
in
the
KtzI-FADred-NADP*-L-orn structure) before being flash frozen in liquid nitrogen prior to data
collection.
Re-reduction of oxidized KtzI crystals
To prepare re-reduced crystals, KtzI was first reconstituted with FAD and NADPH and
crystallized aerobically under the NaBr-containing conditions as described above. After the
formation of data-quality crystals, the crystallization tray was degassed and left under light
vacuum overnight. The next day, the tray was degassed again and brought into the anaerobic
chamber. Aliquots of NADPH and sodium dithionite powder were degassed, brought into the
anaerobic chamber, and mixed with anaerobic crystallization precipitant to a final concentration
of 50 mM and 25 mM respectively.
The reduction of oxidized crystals was achieved by the addition of 4 gL of either the
NADPH or sodium dithionite solution directly to the crystallization drop. Reduction was
monitored visually as a color change from yellow to colorless. Both reductants were capable of
eliciting this oxidation state change, but on very different timescales: 30 min for 50 mM NADPH
108
and <10 sec for 25 mM sodium dithionite. After reduction was complete, 4 p1L of precipitant
supplemented with 10% (v/v) glycerol was added to the drop for cryoprotection and crystals
were looped and flash frozen in liquid nitrogen. Soaking the crystals for 30 min in the 50 mM
NADPH solution deteriorated them such that usable data could not be collected. Attempts to
perform this same re-reduction protocol on crystals derived from KSCN-containing conditions
proved unsuccessful, as evidenced by a complete lack in crystal-color change even during long
incubation periods (5-10 min) with either reductant, culminating in the crystals dissolving into
solution.
Re-oxidation of reduced KtzI crystals
To prepare re-oxidized crystals, KtzI was reconstituted with FAD, NADPH, and
L-ornithine, and crystallized anaerobically under the KSCN-containing conditions as described
above. Once data-quality crystals formed, the crystallization tray was removed from the
anaerobic chamber, and the change in oxidation state could be observed visually as crystals
turned from colorless to yellow over a period of -1 hr. The crystals were allowed to further
equilibrate for ~1 d before cryoprotecting and freezing in liquid nitrogen. Various attempts to
partially oxidize these reduced crystals, in an effort to capture the C4a-hydroperoxy (or some
other) intermediate state, were unsuccessful and resulted in either the reduced or oxidized
equilibrium states described herein.
Data collection, structure determination,and structuralanalysis
X-ray diffraction data were collected at the Advanced Photon Source (Argonne, IL) on
beamline 24 ID-C and processed in the space group P2,2 1 2, using HKL2000 (45) (Tables
IV.2-IV.7). The initial structure of KtzI (reconstituted with FAD and NADPH anaerobically and
crystallized under NaBr-containing conditions) was determined by molecular replacement (MR)
in PHASER (46) with a CHAINSAW (47) constructed search model using the protein
coordinates of the N-hydroxylase from Pseudomonas aeruginosa,PvdA (PDB ID 3S5W (30)),
aligned to the sequence of KtzI. This hybrid model was created such that any conserved residues
between PvdA and KtzI were retained, while all other residues were truncated to their last
common atom according to the protein sequence of KtzI. Solvent content analysis suggested that
four molecules of KtzI would occupy the asymmetric unit, and after using MR to search for 4
109
copies of the hybrid model, a homotetrameric assembly with extensive protein-protein interfaces
was identified. All subsequent structures of KtzI, which adopted the same crystal packing, were
solved by either MR or rigid body refinement in Refmac (48) using the initial refined model's
protein coordinates. For rigid body refinements, the selection of reflections for the calculation of
Rfree was made identical in each model. Each KtzI structure has been abbreviated to signify what
is bound to the protein, and the predicted oxidation states of the cofactors. The abbreviations
used for these bound entities are as follows: FADred (reduced FAD), FADox (oxidized FAD),
FADox-red (re-reduced FAD), FADred-ox (re-oxidized FAD), NADP* (oxidized NADP), L-orn
(L-ornithine), Br (bromide ion that occupies L-orn binding pocket).
Files to describe ligand geometries were obtained in COOT (49), which uses the
REFMAC5 monomer library (50), and included FDA (FADred), FAD (FADox), NAP (NADP),
and ORN (L-ornithine). The restraints for these ligands were constructed using eLBOW, and
those for FAD/FDA were further modified in REEL (51) from the PHENIX suite (52) to allow
the flavin isoalloxazine ring to adopt its bent or "butterfly" conformation. This change was made
such that the ring could fit the non-planar electron density observed in our structures (Fig. IV.7).
The modification of the flavin restraints involved dividing the isoalloxazine into two planes, or
"wings", such that the dimethyl benzene and pyrimidine portions were restrained on separate
planes, with each of these planes including the N5 and NlO positions of the central pyrazine ring.
The pyrazine ring, however, was not restrained to be planar, which allows bending of the outer
"wings" of the isoalloxazine about the central N5-NlO axis (Fig. IV.7).
Ligand-fitting
and
model-building were done in COOT with subsequent refinement in PHENIX, which included
rounds of positional, real-space, B-factor, and simulated annealing refinements. The use of
non-crystallographic symmetry restraints between protomers, and the optimization of target
weights for geometry and B-factor restraints greatly improved model-quality and refinement
statistics. Model-building and refinement were continued iteratively until satisfactory statistics
were achieved (Tables IV.2-IV.7). Each KtzI protein structure begins at Prol0 of the wild-type
sequence (Fig. IV.2) and ends at residue 423 or 424 (of 424), as residues N- or C-terminal to
these positions, respectively, lack electron density and are therefore not included in the final
model (Table IV.8). Residues that lacked clear electron density for their protein side chain are
modeled as alanines (Table IV.8). Composite omit maps were used to verify all final models.
110
For structures crystallized under NaBr-containing conditions, all ion-like densities were
filled exclusively with bromide ions due to: 1. The high concentration of bromide used in
crystallization (>1.0 M NaBr) compared to other likely ions (e.g. ~0.08 M for CL-) and 2. The
fact that all of these ion-like densities disappeared when KSCN was substituted for NaBr in the
crystallization precipitant. It is common for halide ions, like bromide, to compete with water
molecules for hydrogen bonding sites, as well as to occupy the solvent shell in and around the
protein, and these ions are frequently not at full occupancy (53). To determine the most
representative B-factor/occupancy combination for the bromide ions, the B-factor of each was
set to the average B-factor of the late-stage refined model, while the occupancies were set at 0.8,
0.5, and 0.3 in three separate trials. These models were then subjected to 20 rounds of iterative
B-factor and occupancy refinement in PHENIX and found to all converge to the same values,
which are those denoted in the final PDB files.
All software installation support was provided by SBGrid (54). Sequence alignments
were completed using Clustal Omega (55). Structural figures were prepared using PyMOL (56)
and Chimera (57) (Chimera is developed by the Resource for Biocomputing, Visualization, and
Informatics
at the
University
of California,
San Francisco
(supported
by NIGMS
P41-GM103311). Protein surface area calculations were done using the "Protein interfaces,
surfaces
and
assemblies'
service
PISA
at
the
European
Bioinformatics
Institute.
(http://www.ebi.ac.uk/pdbe/prot-int/pistart.html) (58). All root-mean-square deviation (rmsd)
calculations were done for Cu carbons only using the Protein structure comparison service Fold
at European Bioinformatics Institute (http://www.ebi.ac.uk/msd-srv/ssm) (59).
111
IV.VI ACKNOWLEDGEMENTS
This work was supported, in part, by the US National Institutes of Health Grants
GM083464 (to J.R.H.), and GM020011 (to C.T.W.). C.L.D. is an investigator of The Howard
Hughes Medical Institute. This work is based upon research conducted at the Advanced Photon
Source on the Northeastern Collaborative Access Team beamlines, which are supported by
Award RR-15301 from the National Center for Research Resources at NIH. Use of the
Advanced Photon Source, an Office of Science User Facility operated for the US Department of
Energy (DOE) Office of Science by Argonne National Laboratory, was supported by the US
DOE under Contract DE-AC02-06CH11357. Financial support comes principally from the
Offices of Biological and Environmental Research and of Basic Energy Sciences of the US DOE,
the National Center for Research Resources (P41RRO12408), and the National Institute of
General Medical Sciences (P41GM103473) of NIH. Author contributions: J.W.S. and C.L.D.
designed the study. J.R.H. developed the ktzI plasmid construct and provided an initial protein
purification protocol. J.W.S optimized protein purification, and carried out crystallographic
studies and structural analysis. J.W.S and C.L.D wrote the manuscript.
112
TABLES & FIGURES
Table IV.1. Structures of KtzI and their respective resolution.
Structure
Resolution (A)
KtzI-FADred-NADP'-L-orn
2.2
2.4
KtzI-FADred-NADP'-Br
2.1
KtzI-FADox-Br
2.4
KtzI-FADox-NADP*-L-orn
2.6
KtzI-FADox-red-NADP'-Br
KtzI-FADred-ox-NADP*-L-orn
2.7
113
Table IV.2. Data collection and refinement statistics for KtzI-FADred-NADP*-L-orn.
Space group
P212121
Cell constants (A)
(0)
Beamline
Wavelength (A)
Resolution (A)
No. total observations
Redundancy
Completeness (%) C
<I/C(I)>
Rsym (%) a, c
Model refinement
a = 84.2, b = 156.9, c = 163.6
a = = Y = 90
APS 241D-C
0.9792
66 - 2.23 (2.27 - 2.23)
446488
4.2 (4.3)
99.0 (98.8)
20.5 (2.7)
6.5 (57.2)
Rwork(%) b
18.9
Rfee (%)
21.9
b
B-factors (A2 )
Protein
38.6
FAD
30.8
NADP
30.3
L-orn
Ions
Water
rmsd bonds (A) d
rmsd angles (0) d
Number of atoms
Protein
FAD
NADP
33.8
55.7
36.4
0.010
1.3
(4 molecules/asu)
12844
212
192
L-orn
48
Ions
8
Water
549
Ramachandran plot (%)
most favored
90.3
additionally allowed
9.3
generously allowed
0.4
disallowed
0 residues
a: Rsym= IlIhkl - <Ihkl>I/X 'hk, where I is the intensity of a reflection hkl and <I> is the average over
symmetry-related reflections of hkl.
b: Rwork = 2IFO - FeI/2IFOI in which F0 and Fc are the observed and calculated structure factor amplitudes,
respectively. Rfree is calculated from 5% of the reflections not used in the model refinement.
C: Values in parenthesis correspond
to the highest resolution shell.
d rmsd, root-mean-square deviation.
114
Table IV.3. Data collection and refinement statistics for KtzI-FADox-NADP'-L-orn.
P212121
Space group
Cell constants
(A)
(0)
Beamline
Wavelength (A)
Resolution (A)
No. total observations
Redundancy
Completeness (%) c
(%)
c =
= Y = 90
APS 241D-C
0.9795
100 - 2.41 (2.45 - 2.41)
350197
4.1(4.2)
99.0 (99.5)
21.3 (2.4)
5.9(56.8)
<I/I(()>C
Rsym
a = 84.6, b = 157.7, c = 165.3
a, c
Model refinement
Rwor (%) b
18.1
Rfre,
21.8
(%) b
B-factors (A2)
Protein
FAD
NADP
L-orn
Ions
Water
rmsd bonds (A) d
rmsd angles (0) d
Number of atoms
Protein
FAD
NADP
L-orn
Ions
Water
Ramachandran plot (%)
most favored
additionally allowed
generously allowed
disallowed
:K'SYM=11 1 hkI -
25.7
22.3
22.6
24.8
47.9
24.5
0.008
1.3
(4 molecules/asu)
12868
212
192
48
7
562
91.0
8.7
0.3
0 residues
4
' k1Z
/ik,,whereI is the intensiy 0f a reflection uI anu <>1is Me average over
symmetry-related reflections of hkl.
b: Rwork = EIFO - FCI/EIFOI in which F0 and Fe are the observed and calculated structure factor amplitudes,
respectively. Rfee is calculated from 5% of the reflections not used in the model refinement.
Values in parenthesis correspond to the highest resolution shell.
d: rmsd, root-mean-square deviation.
C:
115
Table IV.4. Data collection and refinement statistics for KtzI-FADred-ox-NADP'-L-orn.
Space group
P212121
Cell constants (A)
(0)
Beamline
Wavelength (A)
Resolution (A)
No. total observations
Redundancy
Completeness (%) c
<I/(C)>C
Rsym
(%)
a = 85.0, b = 156.4, c = 164.5
C = = Y = 90
APS 241D-C
0.9795
100 - 2.74 (2.79 - 2.74)
260313
4.5 (4.2)
97.9 (97.6)
21.3 (2.4)
6.3 (56.7)
a, c
Model refinement
18.1
Rwork (%) b
Rfee (%)b
22.1
B-factors (A )
Protein
2
32.9
FAD
29.4
NADP
L-orn
Ions
Water
rmsd bonds (A) d
rmsd angles (0) d
Number of atoms
Protein
FAD
37.9
42.3
53.7
27.0
0.012
1.5
(4 molecules/asu)
12771
212
NADP
192
L-orn
Ions
Water
Ramachandran plot (%)
most favored
additionally allowed
generously allowed
disallowed
a:Rsym= 1hkl
-
<hk1
48
8
237
91.0
8.7
0.3
0 residues
hk,
where I is the intensity of a reflection hkl and <I> is the average over
symmetry-related reflections of hkl.
b:Rork = XIFO - Fc1/XIF.I in which F. and Fc are the observed and calculated structure factor amplitudes,
respectively. Rfree is calculated from 5% of the reflections not used in the model refinement.
C: Values in parenthesis correspond
to the highest resolution shell.
d
116
Table IV.5. Data collection and refinement statistics for KtzI-FADred-NADP'-Br.
Space group
P21212 1
Cell constants
(A)
(0)
Beamline
Wavelength (A)
Resolution (A)
No. total observations
Redundancy
Completeness (%) c
<I/C(I)>
Rsym (%) a,c
Model refinement
a = 81.9, b = 151.7, c = 162.4
c = = y = 90
APS 241D-C
0.9795
50 - 2.39 (2.43 - 2.39)
429887
5.5 (5.6)
96.8 (98.1)
14.7 (2.4)
10.7 (56.9)
Rwork(%) b
20.4
Rfr,,ee(%)
23.1
b
(A2
B-factors
)
Protein
32.6
FAD
23.6
NADP
22.9
L-orn
37.5*
Ions
Water
26.6
0.006
rmsd bonds (A) d
rmsd angles (0) d
1.4
Number of atoms
(4 molecules/asu)
Protein
12781
FAD
212
192
NADP
L-orn
Ions
34
Water
369
Ramachandran plot (%)
90.5
most favored
additionally allowed
9.1
generously allowed
0.5
disallowed
0 residues
a: Rsym= Ihkl - <Ihkl>/X Ihk, where I is the intensity of a reflection hkl and <I> is the average
over
symmetry-related reflections of hkl.
b: Rwork = 2IF 0 - FcI/XIFOI in which F0 and Fe are the observed and calculated structure factor amplitudes,
respectively. Rfr,,ee is calculated from 5% of the reflections not used in the model refinement.
C: Values in parenthesis correspond
to the highest resolution shell.
d: rmsd, root-mean-square
deviation.
e: Average occupancy =
0.65.
117
Table IV.6. Data collection and refinement statistics for KtzI-FADox-Br.
Space group
P212 121
Cell constants (A)
(0)
Beamline
Wavelength (A)
Resolution (A)
No. total observations
Redundancy
Completeness (%) c
<I/__(I)> _
Rsym
(%)
a, c
a = 79.8, b= 151.9, c = 161.9
a = = Y = 90
APS 241D-C
0.9795
50 - 2.09 (2.13 - 2.09)
608641
5.2 (5.3)
99.5 (99.9)
16.5 (2.6)
8.9 (53.7)
Model refinement
Rwork (%) b
22.7
25.6
Rfree (%) b
B-factors (A )
Protein
2
FAD
NADP
L-orn
38.4
31.5
-
Ions
Water
rmsd bonds (A) d
rmsd angles (0) d
31.8e
35.4
0.003
Number of atoms
(4 molecules/asu)
Protein
FAD
NADP
L-orn
Ions
Water
Ramachandran plot (%)
most favored
additionally allowed
generously allowed
disallowed
1.0
12715
212
46
412
89.5
9.9
0.5
0 residues
7 11
-: .SYM:- -' [hkl - <'hkPZ 'ukW, wnere I is tfe intensity or a reflection nKi and <1> is the average over
symmetry-related reflections of hkl.
b: Ror = 1JF0 - FeI/XIFOl in which F and Fc are the observed and calculated structure
factor amplitudes,
0
respectively. Rfree is calculated from 5% of the reflections not used in the model refinement.
C: Values in parenthesis correspond
to the highest resolution shell.
d: rmsd, root-mean-square deviation.
e: Average occupancy
= 0.67.
118
Table IV.7. Data collection and refinement statistics for Ktzl-FADox-red-NADP'-Br.
P212121
Space group
Cell constants (A)
(0)
Beamline
Wavelength (A)
Resolution (A)
No. total observations
Redundancy
Completeness (%) C
<I/J(I)>'C14.8
Rsym (%) ac
a = 82.4, b = 152.0, c = 163.6
a =1=
= 90
APS 241D-C
0.9795
50 - 2.63 (2.68 - 2.63)
300534
4.9 (4.6)
99.2 (99.3)
(2.4)
9.7 (55.3)
Model refinement
Rwork (%) b
19.1
Rfr,,ee (%) b
B-factors (A2)
Protein
FAD
NADP
L-orn
23.4
Ions
Water
rmsd bonds (A) d
30.6
23.4
21.9
31.8e
23.7
0.010
rmsd angles (0) d
1.5
Number of atoms
(4 molecules/asu)
Protein
FAD
NADP
12727
212
192
L-orn-
Ions
Water
28
222
Ramachandran plot (%)
90.7
most favored
additionally allowed
8.9
generously allowed
0.5
disallowed
0 residues
a: Rsym= II/IkI - <Ikl>I/X Ik1, where I is the intensity of a reflection hkl and <I> is the average over
symmetry-related reflections of hkl.
b: Rwork = 1IFO - Fcl/XIFOI in which F0 and F, are the observed and calculated structure factor amplitudes,
respectively. Rfree is calculated from 5% of the reflections not used in the model refinement.
C: Values in parenthesis correspond to the highest resolution
shell.
d: rmsd, root-mean-square
deviation.
: Average occupancy = 0.65.
119
Table IV.8. List of ordered residues and those modeled as alanine in KtzI structures.
Ordered Residues
Residues modeled as alanine
(of residues 1-424)
Structure
Chain A: H120, R137, E150, R196, S197,
Chain A: 10-424
R199, R221, K261, E282, R289, K314, E420
KtzI-FADred-NADP'-L-orn
Chain B: 10-424
Chain B: H120, R137, E150, R196, R199,
R289, E295, K314, D318, E357, E367, E420
Chain C: 10-423
Chain C: R137, E150, R158, Q175, E178,
D192, R196, R199, R221, K261, E282, R289,
K314, R329, E357, E360, E367, R385, E420
Chain D: 10-424
Chain D: E150, R196, R199, R221, K261,
E282, R289, K314, D318, E357, E367, E420,
S424
Chain A: 10-424
Chain A: E150, R196, R199, K261, R272,
E282, R289, K314, R329, E360, E367
Chain B: 10-424
Chain B: H120, E150, Q175, R196, K261,
E282, E295, D353, E357, E420, R422
Chain C: 10-424
Chain C: R137, E150, R153, R158, R196,
K261, E282, R289, K314, E357, E360, E367,
R385, E420, R421
KtzI-FADox-NADP'-L-orn
Chain D: 10-423
Chain A: 10-424
Chain A: R137, E150, R153, D194, R196,
R199, R221, K261, K272, E282, K314, E357,
E360, E367, E420
Chain B: 10-423
Chain B: E150, R196, R199, K272, E282,
E295, D353,E357, E360, E367, E420, R422
Chain C: 10-423
Chain C: R137, E150, R153, R158, Q175,
E178, R191, R196, R221, K261, K272, E282,
R289, K314, R321, D353, E357, E360, H361,
E367, R385, E420
KtzI-FADred-ox-NADP'-L-orn
Chain D: 10-424
120
Chain D: R105, E150, R196, R199, R221,
K261, E282, R289, K314, E357, E420
Chain D: R105, E150, R158, Q175, R191,
R196, R221, K261, K272, E282, R289, K314,
R321,E331,E357,E367,E420
Table IV.8 (continued). List of ordered residues and those modeled as alanine in KtzI structures.
Ordered Residues
Residues modeled as alanine
(of residues 1-424)
Structure
Chain A: H120, R137, R196, S197, R199,
Chain A: 10-423
R221, E282, K314, D318, R329, E360, E367,
E420
Chain B: 10-423
Chain B: R78, H120, R191, D192, R196,
R199, R221, K261, E282, R289, K314, D318,
E357, E360, E367, E420, R421, R422
Chain C: 10-423
Chain C: H120, R137, E150, R158, Q175,
D192, D194, R196, R221, K261, E282, D318,
R329,E357,E360,E367,R385,E420,R421,
R422
KtzI-FADred-NADP'-Br
Chain D: 10-423
Chain D: E150, R191, D194, R196, R199,
R221, K261, Q262, K314, D353, E357, E360,
E367, H396,E420
Chain A: 10-423
Chain A: E121, R137, E150, R196,S197,
R199, R221, R272, E282, K314, D318, R329,
E360, E367, R385 E420, R422, K423
Chain B: 10-423
Chain B: H120, E121, D177, R196, R199,
K261, Q262, R269, R272, E282, K314, D318,
E357,E360,E367,E420
Chain C: 10-423
Chain C: H120, E121, E127, E150, R153,
E156, R158, Q175, R191, D192, R196, S197,
R199, K261, Q262, R269, N273, K314, D318,
E357, E367, R385, E420, R422
10-423
Chain D: H120, E121, E150, Q175, R191,
R193, D194, R196, R199, R221, R272, N273,
K314, E331, E357, E360, H361, E367, E420
Chain A: 10-423
Chain A: R105, E121, R137, E150, R158,
R191, R196, R199,R221, K261, E282, K314,
D318, E360, E367, D383, R385, E420
Chain B: 10-424
Chain B: H120, D192, R193, D194, R196,
R199, R221, K261, E282, K314, D318, R321,
D353, E357, E360, E367, S424
Chain C: 10-423
Chain C: H120, R137, L143, E150, R153,
R158, Q175, R191, D192, D194, R196, S197,
R199, R221, K261, E282, K314, D318, D353,
E357, L358, E360, H361, E362, R385, E414,
E420, R421, R422
Chain D: 10-423
Chain D: H120, E150, E156, R191, R193,
R196, R199, K261, K314, D353, E357, E360,
E367,H396, E420
KtzI-FADox-Br
Chain D:
KtzI-FADox-red-NADP'-Br
121
Figure IV.1. Kutzneride scaffold cryptically incorporates the product of KtzI. (a) The kutzneride
hexadepsipetides are highly decorated natural products produced by nonribosomal peptide synthesis
(NRPS). The piperazic acid moiety (red) of this scaffold is derived from the L-Nahydroxy-ornithine
produced by KtzI. (b) KtzI uses FAD, NADPH, and 02 to catalyze the production of its hydroxy-product
from L-ornithine, which is further processed and inserted into the kutzneride scaffold by uncharacterized
downstream enzymes. (c) Many of the kutzneride biosynthetic enzymes are contained in a contiguous
gene cluster in Kutzneria sp. 744 (adapted from (5)), including the stand-alone enzyme KtzI (red).
a)
OH
HNI"ine--
/
C
cl
N
N
H
0
0
\ 0
0
R, =
R2 =
R3 =
R4 =
R5 =
R5
0 0
N'R
N
HN
(R)-OH, S-(OH)
H, OH, CI
H, n-bond
H, n-bond
H, CH 3
R3
HO
NH
R?
H 3CO
NRPS
kutzneride scaffold
b)
R4
0
-o
NH 2
KtzI-FAD
'o
NADPH NADP+
NH 3
02
L-ornithine
H20
N
OH
HN-
NRPS
HO 2
NH3
N
C
R3
R2
piperazic acid derivative
'""pa
L-N 5 hydroxy-ornithine
c)
0
10
orf1,2
122
orf3 ktzA-C ktzD
ktzE
30
20
ktzF ktzG
ktzH
40
50 kb
ktzl ktzJ ktzK-M ktzN ktzO ktzP ktzQ ktzR ktzS orf4-6 orf7 orf8 orf9
orf 10
Figure IV.2. Sequence alignment of L-orn N-hydroxylases. A sequence alignment of the three
structurally characterized FAD-dependent L-orn N-hydroxylases (KtzI, PvdA, and SidA) displays a high
degree of conservation between these enzymes (33% identity between KtzI and PvdA/SidA). The
sequence shown for KtzI is that of the protein construct used in these studies, where the first 21 amino
acids (red) represent a non-native purification tag. The start methionine and a threonine at position 2 of
the wild-type sequence are absent in our construct, but the numbering has been kept consistent and begins
with the first wild-type residue, a valine at position 3 (Val3).
KtzI
PvdA
SidA
--------- MGSSHHHHHHSSGLVPRGSHMVAHAGESPTHDVVGVGFGPANLSLAVALEE
-----------------------------MTQATATAVVHDLIGIGFGPSNIALAIALEE
MESVERKSESSYLGMRNMQPEQRLSLDPPRLRSTPQDELHDLLCVGFGPASLAIAIALHD
KtzI
PvdA
SidA
SP------------AALTSAFFERRASISWHQGMLLPAAKMQVSFLKDLATFRNPASRFS
RAR---------TQGELQVLFLDKQADYRWHGNTLVSQSELQISFLKDLVSLRNPTSPYS
ALDPRLNKSASNIHAQPKICFLERQKQFAWHSGMLVPGSKMQISFIKDLATLRDPRSSFT
.
KtzI
PvdA
SidA
*
*
*
*
*
.
*
*
.**
:
.:*::
*
. ::*
.
.*
:
*
.
*.
*.
*:
.
.*
:.
.
.:
:
******
::*
.:*
*
*
::
::
.*:**:.
EIVSSIERRKS------------------EISGSLYQHLKPGTAARALHEHALAS-EMVQSIFGEQLERAAVQ---GHQLRAML
*:
. :**
:
249
258
297
:*::.**.*.::
.
309
318
357
:
****:
:****.:***
191
198
237
*::.
356
365
417
**
ELAEHCVQDAEGRWQVDRDYRMVTTPD---LRCGIYLQGGTEHTHGLSSSLLSNLATRSG
PMA-DY----LGDFEVDRNYRLITD---QRCQASIYMQGFCQASHGLSDTLLSVLPVRAE
KVQ-HLRPTGQDQWKPHRDYRVEMDPSKVSSEAGIWLQGCNERTHGLSDSLLSVLAVRGG
.
140
141
180
*
VVGAKR------IADDTRVTV-------YSMAREESYDLDVDVLVCATGYDPMDPGDLLG
VEKATA--GAEG---- IELAL-------RNTANGELSVQRYDLVILATGYERQLHRQLLE
PERKITRVEHHGPQSRMRIHLKSSKPESEGAANDVKETLEVDALMVATGYNRNAHERLLS
*
80
82
120
:
PAAVDDYFDGSKQAKDAFWRYHRNTNYSVVDDEVIRDLYRRGYDDEVAGAPRLNFVNLAH
PEFTDLIYNQEGAERERLVREYHNTUYSVVDIDLIERIYGIFYRQKVSGVPRHAFRSLRS
PERVDKFYSQSAAERQRSLLADKATYSVVRLELIEEIYNDMYLQRVKNPDETQWQHRIL
:
KtzI
PvdA
SidA
.::*:*
:::.
.::*.
KtzI
PvdA
SidA
::*:**:***
DR--DPRSLRRVAVAGGGQSAAEIVRFLHDNRPDTVVHAIMPSYGYVVADNTPFANQIFD
KQPCVNGQPMKVAIIGGGQSAAEAFIDLNDSYPSVQVDLILRGSALKPADDSPFVNEVFA
ALLKDKSKPYIAVLGSGQSAAEIFHDLQKRYPNSRTTLIMRDSAMRPSDDSPFVNEIFN
*
KtzI
PvdA
SidA
**.**:.*
.:*
.::*.******
KtzI
PvdA
SidA
*:
SVLVDVSTP------EATRTVEARNIVISTGLVPRMPAGV---QSDEFVWHSSRFLDHFR
--VEALRVTSRNAE-GEELVRTTRSVVVSAGGTPRIPLAFRHLKDDGRVFHHSQYLERML
SVVDFFTVRSRNVETGEISARRTRKVVIAIGGTAKMPSGL---PQDPRIIHSSKYCTTLP
:.
KtzI
PvdA
SidA
**
.
FVSFLHERGRLVRFANNHDFFPTRREFHDYLEWAESKLAHEVSYDSEVTAIRPGPGRPVD
FVNYLHKHGRLVDFINLGTFYPCRMEFNDYLRWVAGHFAEQSRYGEEVLSVEPMLVERKFLNYLHQKGRLIHFTNLSTFLPARLEFEDYMRWCAQQFSDVVAYGEEVVEVIPGKSDPSS
*:.:**::***:
KtzI
PvdA
SidA
*:::
32
31
60
*
413
417
476
.*.
424
443
501
*:.
123
Figure IV.3. Proposed kinetic mechanism for L-orn N-hydroxylases. The flavin and nicotinamide
cofactors are truncated to their reactive portions and schematic abbreviations are as follows: E, Enzyme;
FADox, oxidized FAD; FADred, reduced FAD; FAD-00-(H), (hydro)peroxyflavin; FAD-OH,
hydroxyflavin; L-orn, L-ornithine; L-N 5OH-orn, L-Nahydroxy-ornithine. There is biochemical evidence
to suggest that the L-orn substrate binds with a protonated side-chain amine (24) (4a), but the neutral form
is eventually necessary for catalysis to occur, as it is used to attack the flavin-hydroperoxide to generate
the hydroxy-ornithine product (5- 6).
E-FAD 'recI-NADP+-L-orn
R
N
\6/
R
N
N
0
NH
N_
L
HH
N
NH
N
E-FADox
H
HR
NH2
N
+
00H3N
E-F ADred-NADP-
HH
0
P
~
NNH
2
N
NH 2
*~~-
N
NADPH
N'I
0
3 NR
NH
N0
R/
rng
N
E-FADox
N
0
o
NH
XN
HO0
1 0 0
E-FAD-OO-(H)-NADP+ 0-(H)
H202
NADP+
~-
R
NH 2
NN
N
NY
H20
NADP+
0H
L-NOH-orn
H
OH
N
H
0
HN
HN
N
0
I,
H2N
O
OH
NH 2
*-'
NH 2
N
*H3 N
'
0
-O
E-FAD-OH-NADP+-L-N 5 OH-orn
124
+
N
I
H3N
0
'
E-FAD-OOH-NADP+-L-orn
Figure IV.4. Crystals of reconstituted KtzI. Crystals of the (a) KtzI-FADred-NADP'-L-orn, (b)
KtzI-FADox-Br, and (c) KtzI-FADox-NADP'-L-orn states are displayed. The intensity of the color
change between (a) and (b) is readily apparent, signifying the reduced (colorless) and oxidized (yellow)
flavin species respectively. In (c), the more subdued, off-yellow color is likely to be a result of the
interaction between the yellow FADox species with the bound NADP' moiety, where these cofactors
stack on one another. The crystals in (a) and (c) can also be used to represent those of the other reduced
(KtzI-FAD(ox-)red-NADP'-Br) and oxidized (KtzI-FADred-ox-NADP-L-orn) states, respectively.
a
C
125
Figure IV.5. Structure of KtzI in reduced and oxidized states. (a) Overall structure of KtzI tetramer
colored by protomer (pale cyan, pale green, wheat, and pale blue) and represented in cartoon overlayed
with a semi-transparent surface representation. (b) The active site of KtzI-FADred-NADP'-L-orn with
FADred, NADP', and L-orn bound in the active site and relevant protein residues displayed (all in sticks).
Carbons are colored gray, marine blue, green, and cyan for these entities, respectively. All other atoms are
colored as follows: oxygen (red), nitrogen (blue), and phosphorus (orange). Hydrogen bonds are
represented as black dashed lines and a red dashed line shows the distance between the reactive portions
of FAD (C4a) and L-orn (N5) during hydroxylation (all lengths denoted in A). The rest of the enzyme is
represented in cartoon form and colored as in (a). (c) The active site of KtzI-FADox-Br (aligned by the
pale cyan protomer in (b)) shows a large change in conformation from its reduced counterpart. FADox
and the same active site residues as in (b) are shown in sticks with carbons colored yellow and salmon
respectively. All other representations and colors are as in (b). (d) The active site of
KtzI-FADox-NADP'-L-orn is an amalgamation of (b) and (c), with NADP*, L-orn, and active site
residues similar to (b), and the FAD conformation similar to that in (c). Active site protein residues are
colored in dark pink with all other colors and representations as in (b) and (c). (e) One side of the active
site of KtzI-FADred-NADP+-L-orn (shown in surface representation) is capped by NADP* (colored as in
(b)) and a protein loop containing Asn275, Tyr276, and Ser277 (the 'Tyr-loop', magenta). All other
representations and colors are as in (b). (f) The surface of the active site shown in (e) changes drastically
due to conformational changes and the exit of NADP'. All colors and representations are as in (e) except
the carbons of FADox are in yellow. (g) The surface represenation of (d) is once again a combination of
(e) and (f), following the same color scheme. Aligning and superimposing the FADred (gray carbons) and
FADox (yellow carbons) equilibrium states for both KtzI (h) and para-hydroxybenzoate hydroxylase
(PHBH) (i), displays the difference between these two proposed conformational changes.
Semi-transparent sticks interpolate the movements necessary to link these two states, with the changes in
both the angle (0) between the isoalloxazine and ribityl tail, and the distances (A) from N5 to N5 of the
isoalloxazine as denoted.
126
4
g) 2)
g
45
C
s,
b
c
His5
Ile
d
His5
FADred Ser406
NADP+
12
Glu
Gu 12'
8
.
rg104
2
2,7'
H i s5 l
Arg1lN
Ser406 Lys67
FADo x
FADox
'~3
.5 S)
.2.7
. 2 .
Lys67
2.7x
Tr27
2~ Asn275 2.8~Asn245
Asn275
sn245
S277
Ser277*4
2.5
e
1
GIu
3.
Glu212
0
.
S [406
y6
AP66
NADP+
2.
2.9
28
L-OrM
Asn275 2
A
yr?76 <A.n245
-2.6
f
5-An
9
!MMI275
FADox
FADred
Tyr-loop'
7FADox
LzrTy76
Asn275
rrI4pI
L-Of~j
h
5*
5.8 A
1 37
6.5A
FADox
FADox
FADred
FADred
127
Figure IV.6. Tetrameric assembly of L-orn N-hydroxylases. (a) The structurally characterized L-orn
N-hydroxylases (KtzI, PvdA (PDB ID 3S5W (30)), and SidA (PDB ID 4B63 (29))) all adopt the same
tetrameric assembly with protomers shown in cartoon and colored pale cyan, pale green, wheat, and pale
blue (left). Aligning and superimposing the pale cyan protomers of these tetramers, while coloring the
other three chains by protein: KtzI (magenta), PvdA (dark green), and SidA (purplish blue), show these
enzymes have high structural similarity (right), but differ significantly at one interface (b). (b) The
tighter interface of KtzI allows a cross-protomer hydrogen bond (dashed line in A) from Tyr270 (magenta
carbons) to NADP' (marine blue carbons). FADred and L-orn (green carbons) are also shown in sticks
and a yellow arrow is used to show the 'clamping' at this interface in KtzI. All other atoms are colored as
in Figure IV.5. (c) Neighboring active sites in KtzI-FADred-NADP'-L-orn are shown with cofactors and
substrate colored as in (b). The wheat colored protomer (in cartoon and semi-transparent surface
representation) interacts with both the pale cyan and pale green protomers (in semi-transparent surface)
and caps one side of the pale cyan protomer's active site. (d) Extracting the pale cyan protomer as a
monomeric unit shows that without the interactions provided by its neighboring protomer, the active site
would be more solvent exposed, with the L-orn substrate (green) at the protein surface. All other atom
colors and depictions are as in (c).
128
a
OFKtzl
11Z
PvdA
SidA
NADP+
NADP'
4
4
Tyr270
r
I
FADr
L-orn
d
C
ik~om
NADP
4FADred
tmL-fn
,v-LO
NAOP
FA-orn
FADred
FADred
NADP*l
129
Figure IV.7. Wall-eyed stereoview of the electron density for bent isoalloxazine ring. Electron
density (2mF.-DFc (1.0a, blue) and mF0 -DF, (-3.0y (red), +3.0a (green)) is displayed for a representative
flavin moiety from KtzI-FADred-NADP*-L-oM with FADred shown in sticks and colored as follows:
carbon (gray), oxygen (red), nitrogen (blue) and phosphorus (orange). Refining this cofactor with its
isoalloxazine ring restricted to be planar (a) or allowed to bend along the N5-NlO axis (b), clearly shows
that a bent ring fits the 2mF0 -DFc density best, while concurrently removing all difference density (b).
The bent isoalloxazine ring is a commonality to all KtzI structures and may result from reduction by
either NADPH or X-ray photoelectrons, as described in the text.
a
b
130
131
Figure IV.8. Wall-eyed stereoviews of the active sites of reduced L-orn N-hydroxylases. The active
sites of (a) KtzI-FADred-NADP'-Br (teal carbons), (b) reduced PvdA (PDB ID 3S61 (30)), light pink
carbons), (c) reduced SidA (PDB ID 4B65 (29)), light orange carbons), and (d) KtzI-FADox-red-NADP'Br (light purple carbons) (all aligned with the pale cyan protomer of Fig. IV.5b) show these structures to
be relatively identical. Bromide ions are displayed as dark red spheres. All protomers, ligands, and other
atoms are represented as in Figure IV.5. In (c), asterisks and semi-transparent sticks represent the L-orn
and Asn293 conformations from an oxidized SidA structure (PDB ID 4B63 (29)), as the reduced structure
(PDB ID 4B65 (29)) does not have substrate bound.
132
af
His5l
His5
04 F reft04
1~
FADre
Ser406O
~ Se46
y
4~~~
As25 -Asn245
y6
A2~~ sn275 "'fAsn245
Tyr276
8b
J.
~
Ar 0
F~e Ser41 LOA
NA~~
Ari6 FADred Ser4l 04
NADW
j7 Lys~g
) Lys69
SGu2
G~u221Asn284
'sn284
~
Tyr245
Asn254
ArI4 F~rdSer;469 L
NAP,
Glu26Q.
<7
S 6 Tyr285
Y
10
FAra Ser
NADP
"
;Asn323
4As54
n5
4
3.~~5~Ty324Asn293*
Gu260~
~2 5
<~
i
Lysl07
orn
Asn323
Tyr324As23
His5lA
Arl4 ADred Ser4O6
Lys67
FArr d Ser406
NADI
'AD
~ys67,#
luGlu2
jj 1 Asn275
6 7 'yr276
An4
4
fsn4
Tyr276
133
Figure IV.9. Wall-eyed stereoviews of the active sites of oxidized KtzI. The active sites of (a)
KtzI-FADox-Br (salmon carbons), and (b) KtzI-FADred-ox-NADP'-L-orn (dark purple carbons) were
aligned with the pale cyan protomer in Fig. IV.5b and are displayed with FADox in yellow carbons and
all other protomers, ligands, and atoms represented as in Fig. IV.8.
a
FADox
Ys
His51
*
Arg104
S( r406
Tyr2
Asn275
Tyr276
Lys6 7
Arg 104
FAox
s6
.sn275
Asn2 45
er277*-'
Se 406
Lys67
.4
Tyr276
Asn245
Ser277
b
His51
His5
Sr406
Lys67
FADox
S046
FADox
NADP+
NADP+
GIu21
GIu21
L-orn
GAsn275
yr276
134
Lys67
sn245
L-orn
sn275
Tyr276
n245
135
Figure IV.10. Structurally-based proposal for L-orn N-hydroxylase mechanism. (A) L-orn
N-hydroxylases with an oxidized flavin could exist in a dynamic equilibrium between the FADox(out)
conformation observed in KtzI-FADox-Br (with or without the 'Tyr-loop' insertion) and the FADox(in)
of SidA-FADox-L-orn (protein carbons in gold, PDB ID 4B69 (29)). (B) Anaerobic reduction of
KtzI-FADox-Br
crystals
recapitulated
its
anaerobically
reconstituted
counterpart,
KtzI-FADred-NADP'-Br (protein carbons in teal). (C) Binding of L-orn, as displayed for
KtzI-FADred-NADP'-L-orn, causes minimal deviations within the active site of the reduced enzyme. A
close-up of the active site (inset) with Van der Waals radii displayed for substrate L-orn (gray spheres)
and NADP' (atom-colored spheres) show the fit of these two molecules. After oxygen addition and
hydroperoxy-flavin generation, hydroxylation to the L-N 5OH-orn product would occur (1). If this newly
hydroxylated product (orange carbons) maintains the same positioning as the substrate, a steric clash
between the Van der Waals radii of the newly added N5-OH group (gray spheres) and the bound NADP
cofactor would occur, which could lead to a destabilization of NADP' binding (2). (D) This
destabilization could explain the poorly resolved nicotinamide portion of NADP' found in the
L-N 5OH-orn-bound structure of PvdA (PDB ID 3S5W (30)). After catalysis, the resting state enzyme
would need to be reproduced through dissociation of the hydroxyornithine product, spent NADP'
cofactor, and water (3). We have shown that a conformational change takes place in crystallo after
oxygen addition to the fully-liganded reduced state in (C), in a background of excess NADP(H), which
yields the KtzI-FADox-NADP*-L-orn state shown in (E). Crystals of this oxidized state are unable to be
reduced by NADPH or dithionite, and thus it appears to be a dead-end, inhibited conformation. NADP'
binding to the FADox(out) state in (A) could cause a state similar to what we observe in (E), which could
be blocked by insertion of the Tyr-loop. Dotted-lined arrows are shown for steps without direct
experimental evidence. Colors for protein, cofactors, and ligands not described above are as in Figure
IV.5. Bromide ions were removed from the KtzI structures for clarity.
136
Ax
FADox(out)
LyS67
02 +NADP+
NADP+,,'
NADPH
KtzI-FADox-NADP+-L-Orn
©l
__
Argl04
Lys67
FADxot
Arg14
Ly)
NADPH
-G
KtzI-FADox-Br
rg
u212
P
A~re Lys
31 21
~FAr
A~1
FD~d
l Lys67
y
NADP
FA
Asnn323 Asn2
ArglArg#
Ktu212
L-or
LyS6
NAF
o-B
G1SidA-FA
6
2x-L-Orn
Asn25 As
Asn275 Asn2
324
Tyr276
Tyr276
KtzI-FAD
KtzFADred-NADP+-L-OM
red-NADP+-Br
02
.. 'ejection'
H20
NADP+
Lys6s
L-NOH-om
VF33As284
PvdA-FADox-NADP-L-NH-orn
. 2.'destabilization'
1.'hydroxylation'
''...(02)
137
II.VII REFERENCES
1.
2.
3.
4.
Newman, D. J., and Cragg, G. M. (2012) Natural products as sources of new drugs over the 30
years from 1981 to 2010., JNat Prod 75, 311-335.
Walsh, C. T., and Wencewicz, T. A. (2013) Prospects for new antibiotics: a molecule-centered
perspective., J Antibiot 67, 7-22.
Pohanka, A., Menkis, A., Levenfors, J., and Broberg, A. (2006) Low-abundance kutznerides from
Kutzneria sp. 744., J Nat Prod 69, 1776-1781.
Broberg, A., Menkis, A., and Vasiliauskas, R. (2006) Kutznerides 1-4, depsipeptides from the
actinomycete Kutzneria sp. 744 inhabiting mycorrhizal roots of Picea abies seedlings, J Nat Prod
69, 97-102.
5.
Fujimori, D. G., Hrvatin, S., Neumann, C. S., Strieker, M., Marahiel, M. A., and Walsh, C. T.
(2007) Cloning and characterization of the biosynthetic gene cluster for kutznerides., Proc Natl
Acad Sci USA 104, 16498-16503.
6.
7.
Zolova, 0. E., and Garneau-Tsodikova, S. (2013) KtzJ-dependent serine activation and
0-methylation by KtzH for kutznerides biosynthesis., J Antibiot 67, 59-64.
Neumann, C. S., Jiang, W., Heemstra, J. R., Gontang, E. A., Kolter, R., and Walsh, C. T. (2012)
Biosynthesis of Piperazic Acid via N(5) -Hydroxy-Ornithine in Kutzneria spp. 744.,
Chembiochem 13, 972-976.
8.
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142
Chapter 5. Outstanding Questions for AAG and KtzI
V.1
SUMMARY
In this dissertation, we have examined the human DNA repair protein alkyladenine DNA
glycosylase (AAG) (Chapters 2 and 3), and a flavin-dependent enzyme involved in antibiotic
biosynthesis called KtzI (Chapter 4). These dissimilar systems are linked by their reliance on
conformational flexibility for catalysis, and substantial progress was made on understanding both
systems through collaborative efforts. As science is a perpetually evolving endeavor, many
unresolved questions remain for these enzymes. In this chapter, we will bring forward what we feel
are the most interesting outstanding questions in each system, in hopes of inspiring future
generations of researchers to take the lead in answering them.
V.11
Repairing DNA with AAG
In Chapters 2 and 3, we analyzed the human DNA repair protein alkyladenine DNA
glycosylase (AAG). As mentioned in Chapter 1, the most common manifestations of DNA
damage are known as DNA lesion bases (1),
where a canonical DNA base is altered by
endogenous or exogenous factors to create a new chemical entity. These lesion bases occur at a
rate of 10,000 per cell per day (1,
2), and AAG is involved in the removal of a structurally
diverse group of these lesion bases (3, 4) to initiate a DNA repair pathway. Our lab has
collaborated with the Samson Group to solve crystal structures of an N-terminally truncated form
of this protein (A79AAG) bound to DNA containing 3,N 4-ethenocytosine (EC) (5, 6). This lesion,
caused by lipid peroxidation (7), is rather prevalent and is of specific interest because AAG has
been shown, by us (6) and others (8), to bind cC with high affinity, while being unable to excise
it. We have snapshots of the enzyme both recognizing the PC inhibitor lesion (Chapter 2 (6)),
and interacting with DNA in a lower-affinity, nonspecific manner (Chapter 3 (5)). These studies
have provided great insight, but as mentioned above, contain the caveat of using a truncated
protein construct. We feel that characterization of the full-length AAG (FL-AAG) enzyme is the
most important outstanding structural question for this system.
All crystallographic work to date has been with an N-terminally truncated AAG protein,
which has been able to give substantial insight into the recognition of DNA (5, 6, 9, 10). This
truncated protein, although shown to have wild type catalytic activity (3, 6), appears to be
hindered in its overall processivity, especially at conditions mimicking a physiological
143
environment (11).
In separate studies, AAG's N-terminus has been implicated in overcoming
product (abasic site) inhibition (12, 13) and in recognizing substrate lesions (12). Our truncated
A79AAG construct removes this N-terminus (Fig. V.1), such that the wild-type AAG sequence
begins with Thr84, and residues left from cleavage of the purification tag add Gly80, Pro8l,
His82 and Met83 prior to this residue. Our construct is unperturbed as far as catalysis is
concerned (6), so understanding the specific interactions of AAG with lesion bases should be
unaffected. In light of the functions attributed to the N-terminus of AAG, however, especially
considering the 13 Arg or Lys residues in this region (Fig. V.1) that could be used to bind DNA,
a structure showing FL-AAG would greatly increase our understanding.
We have collaborated with the laboratory of Patrick O'Brien at the University of
Michigan Medical School to attempt structural characterization of the FL-AAG enzyme, but
have yet to produce viable crystals. The full-length protein is very active and pure, but can only
be concentrated after purification to approximately 5 mg/mL (~150 uM) as determined by active
site titration (A2 60/A2 80 = 0.58 by absorbance). This low concentration has made crystallization
difficult, but some success has been had in concentrating the protein further after incubating with
DNA (unpublished results). A 13-mer DNA duplex containing a pyrrolidine abasic site mimic
opposite a thymine (pyr:T) added to FL-AAG prior to spin filter concentration, has allowed for
soluble protein up to concentrations of approximately 30 mg/mL by absorbance measurements
(A 2 60/A 2 80
=
1.57;
E28O
=
28150 M-cm' and molecular weight = 32868 (protein-DNA complex)).
Complex formation was not confirmed before crystallization efforts, and no viable crystals were
produced through these experiments. An optimization of this method, including confirming a
protein-DNA complex through another technique like native polyacrylamide gel electophoresis,
with subsequent complex purification by size exclusion chromatography, could be used to attain
crystal structures of FL-AAG. Another way to obtain a FL-AAG complex structure would be to
use the disulfide crosslinking method pioneered by the laboratory of Gregory Verdine at the
Harvard University (14). This technique allows physical linking of protein and DNA through a
disulfide bond, followed by subsequent purification steps, thus creating a purified covalent
complex for crystallization. Indeed, this method has been employed successfully for the
homologue of AAG from E. coli, AlkA (15, 16). This disulfide crosslinking is further useful as it
can be performed with both lesion-containing, and undamaged DNA, providing insight into both
specific and nonspecific protein-DNA interactions. In any case, a structure of FL-AAG in
144
complex with DNA would finally provide a depiction of its functionally important N-terminal
region, providing new perspective on this enzyme, such that some current questions could be
answered while initiating further research.
V.111
Antibiotic biosynthesis with KtzI
In
Chapter
4,
we
investigated
the
flavin-dependent,
L-orn
N-hydroxylase
(monooxygenase) from Kutzneria sp. 744, called KtzI. This enzyme takes L-ornithine to
L-N 5hydroxy-ornithine using FAD, NADPH, and 02, and this hydroxylamine product is further
incorporated into antifungal antimicrobials called kutznerides. We have been able to structurally
characterize KtzI in several states along its catalytic trajectory by using various chemical and
environmental manipulations. These states are suggestive of never-before-seen conformational
changes of the flavin, which we have been able to recapitulate in crystallo, signaling their
relevance to the catalytic cycle. The flavin-dependent hydroxylases have been studied for over
50 years, and although their overarching reaction mechanism is conserved in the majority of
enzymes (see Chapter 1 and Fig. V.2; reviewed in (17-20)), there are intriguing differences
observed in the N-hydroxylase class. As described in Chapter 4, N-monooxygenases, although
formally grouped with Class B enzymes (18), borrow qualities from both major classes of flavin
monooxygenases, such that they have the substrate specificity of the so-called "cautious" (or
Class A) enzymes (Fig V.2b), but have an oxidative half reaction wherein NADP* remains bound
throughout, mimicking that of the "bold" (or Class B) enzymes (Fig. V.2a), which tend to be
more promiscuous hydroxylaters overall (reviewed in (17, 18)). Class A enzymes couple
substrate binding to NADPH reduction using dynamic movements of the FAD isoalloxazine ring
(Fig. V.3b). FAD moves from the protein interior ('in') to the protein exterior ('out') upon
substrate binding, allowing reduction by NADPH. Once reduction occurs, the reduced flavin
moves from this 'out' position back 'in'. This control point helps guard against unproductive
uncoupling of the reaction as the reactive C4a-hydroperoxy-flavin intermediate can only be
generated in the presence of substrate. KtzI, and all other Class B flavin monooxygenases, are
just as easily reduced by NADPH in the absence of substrate, such that they do not share the
same control point as Class A enzymes. Although the precedence for flavin conformational
changes controlling catalysis exists for "cautious" enzymes, KtzI is the first Class B
monooxygenase for which flavin movement has been suggested (Fig. V.3a), and we propose that
145
these conformational changes could be correlated in both Class A and Class B systems in an
effort to control the "flaws" in their respective paths to a hydroxylated product (Fig. V.2).
As mentioned above, the Class A monooxygenases couple substrate-binding to NADPH
reduction using movements of the bound flavin (Fig. V.3b; (17, 18)). This protective step is
necessitated by the "flaw" that Class A enzymes are unable to stabilize the reactive flavin
C4a-hydroperoxide to any degree, such that addition of oxygen to the reduced flavin must occur
only if and when substrate is present to be hydroxylated. Class B monooxygenases are capable of
stabilizing the C4a-hydroperoxide on the order of minutes to hours (the quality from which the
"bold" moniker is derived), and they are able to do this due to protection afforded by a bound
NADP* molecule (removing the "flaw" from Class A systems) (17, 18). Indeed, 02 addition to a
chemically reduced flavin in the absence of this bound NADP', results in immediate uncoupling
for "bold" enzymes. These observations display the pivotal role NADP(H) plays throughout the
reaction cycle, as both a reductant and a protector of intermediate states. NADP' remaining
bound throughout catalysis bring its own "flaw", however, as this cofactor must eventually be
dissociated before another round of catalysis begins using a new NADPH reducing equivalent.
Therefore, what signals the spent nicotinamide cofactor to leave precisely after the very last step
in catalysis? Our proposal for L-orn hydroxylases, as outlined in Chapter 4, is that hydroxylation
acts as an intermediate signal through a steric clash between the hydroxy-product and NADP',
resulting in a destabilized cofactor. This intermediate signal is currently limited to the L-orn
N-hydroxylase active site architecture, but all Class B monooxygenases have the problem of
getting rid of a cofactor that has been bound throughout the catalytic cycle. As discussed in
Chapter 4, hydroxylation alone was not enough to eject NADP' in L-om N-hydroxylases,
therefore, we have proposed a conformational change-induced ejection of NADP' by a flavin
movement from an 'in' to 'out' position (Fig. V.3a), which is distinct from the 'in' and 'out'
states observed for Class A enzymes (Fig V.3b). This proposal need not be limited to
N-hydroxylases, and we believe that similar structural characterization of chemically and
environmentally manipulated Class B monooxygenases could tease out similar conformational
changes. Further, there is very little biochemical data that support flavin conformational changes
in N-hydroxylases, and as mentioned in Chapter 4, this may be due to similar spectroscopic
signals coming from structurally dissimilar states. We believe it is worthwhile to re-examine
existing pre-steady state kinetic data in light of our observations of varied flavin conformations,
146
but also to design new studies to look specifically for these changes. It will likely be useful to
examine Class B monooxygenases using derivatives of flavin, and accompanying active site
mutants, in an effort to slow down, or enhance the signal of, what may be very transient flavin
movements. In any case, our structural data suggest that movements of the flavin cofactor in both
Class A and Class B hydroxylase enzymes could be correlated to address respective "flaws" in
their kinetic mechanism, linking these two distinct classes by their reliance on conformational
changes for controlling catalysis.
147
148
FIGURES
Figure V.1. Sequence alignment of our N-terminal truncation mutant (A79) and full-length (FL)
alkyladenine DNA glycosylase (AAG). Positively charged residues in the N-terminus of FL-AAG are
colored blue, and the residues left behind after tag cleavage in our A79AAG construct are colored pink.
FL-AAG
A79AAG
MVTPALQMKKPKQFCRRMGQKKQRPARAGQPHSSSDAAQAPAEQPHSSSDAAQAPCPRER
------------------------------------------------------------
FL-AAG
A79AAG
CLGPPTTPGPYRSIYFSSPKGHLTRLGLEFFDQPAVPLARAFLGQVLVRRLPNGTELRGR
GPHMTRLGLEFFDQPAVPLARAFLGQVLVRRLPNGTELRGR
-------------------************
*F**************************
FL-AAG
IVETEAYLGPEDEAAHSRGGRQTPRNRGMFMKPGTLYVYIIYGMYFCMNISSQGDGACVL
A7 9AAG
IVETEAYLGPEDEAAHSRGGRQTPRNRGMFMKPGTLYVYI IYGMYFCMNI SSQGDGACVL
* *******************
* ************************
***************
A79AAG
LRALEPLEGLETMRQLRSTLRKGTASRVLKDRELCSGPSKLCQALAINKSFDQRDLAQDE
LRALEPLEGLETMRQLRSTLRKGTASRVLKDRELCSGPSKLCQALAINKSFDQRDLAQDE
FL-AAG
AVWLERGPLEPSEPAVVAAARVGVGHAGEWARKPLRFYVRGSPWVSVVDRVAEQDTQA
A7 9AAG
AVWLERGPLEPSEPAVVAAARVGVGHAGEWARKPLRFYVRGSPWVSVVDRVAEQDTQA
FL-AAG
* *******************
*************************
*************
149
Figure V.2. Kinetic mechanism for hydroxylation in (a) bold and (b) cautious flavin-dependent
monooxygenases. Abbreviations are as follows: E, Enzyme; FAD, oxidized FAD; FADre, reduced
FAD; FADOO-(H), (hydro)peroxyflavin; FADOH, hydroxyflavin; S, substrate; SOH, hydroxylated product.
Light gray states are those that are transient in nature. Green stars indicate proposed control points where
flavin conformational changes are vital to the catalytic cycle for each system.
a)
NA
E-FA D
- E-FADe-NADP+ -0-2-'- E-FAD
APH
-NADP+
E-FAD
E-FADoH-NADP+-SOH
S.DE-FAD
-NADP+-S ->E-FAD
-NADP+-SH
1.NADPH; 2.S
b)
""at E
E-FAD-S
H20
NADP.
NADPH
S
F
:-S
NADP+
150
-TsAE-FAD,-S
EF
-rpuuX
S
02_1E--AV
-
E-FAD O-SOH
SHOH
x
Figure V.3. Diverse flavin conformations are found in (a) KtzI and (b) para-hydroxybenzoate
hydroxylase (PHBH). The 'in' (grey carbons) and 'out' (yellow carbons) conformations in KtzI (a) vs.
PHBH (b) suggest very different flavin movements. All other atoms are colored as follows: oxygen (red),
nitrogen (blue), and phosphorus (orange).
FAD 'out'
90
FADF'in'
FAD 'in'
FAD
Fout'
b
FAD 'out'
90
FA D ' i n'
FAD 'out'
FAD 'in'
151
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Lingaraju, G. M., Davis, C. A., Setser, J. W., Samson, L. D., and Drennan, C. L. (2011)
Structural basis for the inhibition of human alkyladenine DNA glycosylase (AAG) by 3,N4ethenocytosine containing DNA, J Biol Chem 286, 13205-13213.
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basis for discriminating between normal and damaged bases by the human alkyladenine
glycosylase, AAG, Proc Natl Acad Sci USA 97, 13573-13578.
Lau, A., Scharer, D., Samson, L., Verdine, G., and Ellenberger, T. (1998) Crystal Structure of a
Human Alkylbase-DNA Repair Enzyme Complexed to DNA: Mechanisms for Nucleotide
Flipping and Base Excision, Cell 95, 249-258.
Hedglin, M., and O'Brien, P. J. (2008) Human Alkyladenine DNA Glycosylase Employs a
Processive Search for DNA Damage, Biochemistry 47, 11434-11445.
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glycosylase is required for turnover in hypoxanthine excision reaction, J Biol Chem 30078-30084.
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repair enzyme N-methylpurine-DNA glycosylase. Interaction, conformational change, and role in
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Jeremy W. Setser
77 Massachusetts Ave., BLDG 68-688D
Cambridge, MA 02139
Tel.: (216) 375-5606
Fax: (617) 258-7847
Email: jsetser@mit.edu
EDUCATION
June 2014
Massachusetts Institute of Technology (MIT), Cambridge, MA
Ph.D., Biological Chemistry
May 2008
The University of Akron (UA), Akron, OH
B.S., Chemistry, summa cum laude
HONORS AND AWARDS
Koch Institute for Integrative Cancer Research Fellowship (2009), MIT Chemistry Department Award for
Outstanding Teaching (2009), UA Chemistry Department Outstanding Senior (2008), National SMART Grant
(2007-2008), Emmanuel and Rose Gurin Scholarship in Chemistry (2007-2008), Oelschlager Pin Oak
Scholarship (2004-2008)
RESEARCH EXPERIENCE
2008-Present
MIT Department of Chemistry and HHMI, Cambridge, MA
Position: Graduate Research Assistant, Laboratory of Prof. Catherine L. Drennan
Investigating enzyme mechanism through structural snapshots using protein X-ray
crystallography.
Provided mechanistic insight for a required enzymatic step in the biosynthesis of an
antimicrobial and antifungal class of natural products known as kutznerides
(collaboration with Prof. Christopher T. Walsh, Harvard Medical School). The enzyme in
question, KtzI, uses a bound flavin cofactor, reducing equivalents from NADPH, and
molecular oxygen to install a hydroxyl group on the side-chain nitrogen of the amino acid
L-ornithine, which is subsequently incorporated into the kutzneride scaffold. KtzI was
structurally characterized after being subjected to various chemical and environmental
factors, capturing the enzyme in several states along its catalytic trajectory. These states
suggest that novel conformational changes of both the protein backbone and flavin
cofactor occur during the enzymatic cycle of KtzI, and this drastic rearrangement was
recapitulated in the protein crystal.
-
Collaborated with Prof. Leona D. Samson (MIT) to elucidate the molecular interactions
of a human DNA repair protein, alkyladenine DNA glycosylase (AAG), both in complex
with inhibitor DNA, and in a low-affinity 'searching state' on DNA. When taken
together, these studies allow us to compare how AAG interacts with substrates compared
to inhibitors, and also provide evidence for the searching mechanism of this enzyme.
SKILLS
Technical: bacterial cell culture, recombinant protein expression and purification including selenomethionine
derivatization, molecular biology, high-throughput (Formulatrix,TTP Labtech, Art Robbins Instruments) and
anaerobic (Coy Lab Products, MBraun) protein crystallization and imaging, single-crystal X-ray data
collection and processing using in-house (Rigaku) and synchrotron (APS, ALS, CHESS) facilities, small-angle
X-ray scattering (SAXS) sample preparation and data collection (CHESS), solving and refinement of protein
co-crystal structures, enzyme assays, UV-Vis spectroscopy.
Computer: Linux-based OS, Windows, Mac OS, Microsoft Office, Adobe Illustrator, experienced with most
major crystallography software packages including but not limited to: HKL/HKL2000, SCALEPACK, CCP4,
PHENIX, CNS, SHELX, SHARP, Coot, PyMOL, UCSF Chimera.
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PUBLICATIONS
3. J.W. Setser, J.R. Heemstra, C.T. Walsh, C.L. Drennan. Crystallographic evidence for drastic conformational
changes in the active site of a flavin-dependent N-hydroxylase. In Preparation.
2. J.W. Setser, G.M. Lingaraju, C.A. Davis, L.D. Samson, C.L. Drennan (2012) Searching for DNA lesions:
Structural evidence for lower- and higher-affinity DNA binding conformations of human alkyladenine DNA
glycosylase. Biochemistry 51, 382-390.
1. G.M. Lingaraju, C.A. Davis, J.W. Setser, L.D. Samson, C.L. Drennan (2011) Structural basis for the
inhibition of alkyladenine DNA glycosylase (AAG) by 3,N 4-ethenocytosine containing DNA. J Biol Chem
286, 13205-13213.
ORAL PRESENTATIONS
Apr. 2014
Apr. 2013
Feb. 2013
June 2012
Mar. 2010
Interview Seminar at Constellation Pharmaceuticals
"Mechanistic crystallography of a flavoenzyme involved in antibiotic biosynthesis"
MIT Chemistry Student Seminar Series
"Structural investigation of a flavin-dependent N-monooxygenase"
Protein Structure Function Supergroup Meeting
"Structural investigation of a flavin-dependent N-monooxygenase"
MIT Department of Biology Retreat
"Structural insight into the searching mechanism of human alkyladenine DNA glycosylase
(AAG)"
Boston DNA Repair and Mutagenesis Seminar Series
"Structural insight into the catalytic and searching mechanism of human alkyladenine DNA
glycosylase (AAG)"
POSTER PRESENTATIONS
July 2012
Aug. 2010
Jan. 2010
June 2009
May 2009
Gordon Research Conference - Enzymes, Coenzymes, and Metabolic Pathways
"Structural investigation of a flavin-dependent N-hydroxylase"
240th Meeting of the American Chemical Society
"Crystallographic study of a human DNA repair protein: alkyladenine DNA glycosylase
(AAG)"
MIT Biology Department Research Symposium and Poster Session
"Crystallographic study of a human DNA repair protein: alkyladenine DNA glycosylase
(AAG)"
MIT Biology Retreat Poster Session
"Crystal structure of human 3-methyladenine DNA glycosylase (AAG) complexed with
3,N4-ethenocytosine containing DNA"
MIT Center for Environmental Health Sciences Poster Session
"Crystal structure of human 3-methyladenine DNA glycosylase (AAG) complexed with
3,N4-ethenocytosine containing DNA"
TEACHING AND MENTORING EXPERIENCE
Dec. 2012
2010-2012
Summer 2011
154
Laboratory Mentor, Drennan Laboratory, MIT
- Trained and mentored a rotation graduate student from the Biology Department resulting
in the co-crystallization and structural determination of an enzyme-ligand complex.
Laboratory Mentor, Drennan Laboratory, MIT
" Trained and mentored an undergraduate conducting research in the Drennan Lab.
Participant, HHMI-MIT Summer Mentoring Seminar, MIT
* Enrolled in an HHMI-affiliated training course at MIT to learn and refine effective
undergraduate mentoring strategies in laboratory research.
TEACHING AND MENTORING EXPERIENCE (continued)
Fall 2010
Spring 2009
Fall 2008
Laboratory Teaching Assistant, MIT
Advanced Biochemistry
" Supervised and instructed graduate students (10) in biochemical techniques.
" Observed and graded overall performance.
Laboratory Teaching Assistant, MIT
Biochemistry & Organic Lab
" Supervised and instructed undergraduate students (14) in standard biochemical
techniques.
" Observed and graded presentations and lab reports.
HHMI Teaching Assistant, MIT
Principles of Chemical Science
- Taught a twice-weekly recitation class of 23 undergraduates for an HHMI-funded
chemistry course focused on bringing medical and biochemical examples into freshman
chemistry.
- Addressed students' questions and needs on an individual basis and graded assignments
and tests.
LEADERSHIP AND DEVELOPMENT
2008-Present
2012-Present
2012-2014
2010-2013
2013
2010-2012
2012
MIT Chemistry Graduate Student Committee, President, Treasurer, Member
" Advocated for chemistry graduate students' interests by working with the department
head and senior department officials to amend departmental procedures (e.g. Ph.D.
candidacy exams).
" Improved the quality of life for chemistry graduate students (230 students) by organizing
and executing various events including forum discussions and socials (e.g. boat cruise,
BBQ).
" Presided over 15 executive committee members and a $15,000 annual budget.
MIT Chemistry Career Panel, Founding Member
- Created and secured departmental funding ($1450) for an annual series of 10 career panel
discussions.
- Recruited and prepared individual panel organizers (10) and panelists (38), receiving a
panel-by-panel approval rating of 92%.
- Enlisted incoming panel leadership and provided knowledge transfer and guidance.
MIT Chemistry Graduate Student Survey Task Force, Founding Member
- Formulated survey questions and analyzed responses from chemistry graduate students.
- Communicated results through written and oral presentations to the MIT chemistry
faculty and students.
MIT Graduate Student Council, Representative from the Department of Chemistry
" Expressed the sentiments and opinions of the chemistry graduate student body during
meetings with 60 graduate student Representatives from across the Institute.
- Voted on agenda items on behalf of all chemistry graduate students.
Strategic Decision Making in the Biomedical Business, MIT Sloan Course
* Explored the product development life cycle for four biomedical industries: smallmolecule drugs, biological therapeutics, medical devices, and diagnostics/personalized
medicine.
MIT Chemistry Outreach Program
" Teamed up with fellow graduate students to perform chemistry demonstrations at local
schools.
NERCE Medicinal Chemistry and Preclinical Development Workshop, Participant
- National Institutes of Health workshop describing resources available for translational
research.
155
ADDITIONAL RESEARCH EXPERIENCE
2006-2008
UA Department of Chemistry, Akron, OH
Position: Undergraduate Researcher, Laboratory of Prof. Daniel J. Smith
Created a polymeric bandage imbued with a functionalized dextran sugar for the release
of nitric oxide to aid slow-healing wounds.
REFERENCES
Ph.D. Advisor
Catherine L. Drennan
Professor of Chemistry and Biology
HHMI Investigator and Professor
Massachusetts Institute of Technology
77 Massachusetts Ave, BLDG 68-680
Cambridge, MA 02139
Tel.: 617-253-5622
Email: cdrennan@mit.edu
Thesis Committee Chair
Elizabeth M. Nolan
Pfizer-Laubach Career Development Assistant Professor of Chemistry
Massachusetts Institute of Technology
77 Massachusetts Ave, BLDG 16-573B
Cambridge, MA 02139
Tel.: 617-452-2495
Email: Inolan@mit.edu
Faculty Advisor, MIT Chemistry Career Panel
Bradley L. Pentelute
Assistant Professor of Chemistry, MIT
Associate Member, Broad Institute of Harvard and MIT
77 Massachusetts Ave., BLDG 18-596
Cambridge, MA 02139
Tel.: 617-324-0180
Email: blp@mit.edu
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