INHIBITION AND NUCLEIC ACID BINDING STUDIES
OF THE CARBOXYLTRANSFERASE COMPONENT OF
BACTERIAL ACETYL-COA CARBOXYLASE
A Dissertation
Submitted to the Graduate Faculty
of the Louisiana State University and
Agricultural and Mechanical College
in partial fulfillment of the
requirements for the degree of
Doctor of Philosophy
in
the Department of Biological Sciences
by
Brian K. Benson
B.S., Louisiana State University, May 2002
December 2011
ACKNOWLEDGEMENTS
I would like to thank Dr. Grover Waldrop, my major professor, for accepting me into his
laboratory and providing technical expertise throughout this work and for fostering my belief
that our efforts someday will help sick people get out of hospital beds. I am also appreciative of
my graduate committee members, Professor Paul Russo, Professor Roger Laine, Professor
Jeffrey Gimble, and Associate Professor Anne Grove, for their valuable advice. Thanks to
Patrick Bilder and Michael Oldham for their suggestions and to Carol Dieckmann, Cindy
Fleming-Wood and Angela Hopp for their careful consideration of this dissertation. I wish also
to acknowledge my wife; without her communicable faith and indispensable support, I certainly
would not have achieved this task. Finally, to my children: Thanks for helping me realize how
precious life is and for inspiring me to always work smarter and harder.
ii
TABLE OF CONTENTS
ACKNOWLEDGEMENTS ............................................................................................................ ii ABSTRACT.................................................................................................................................... v CHAPTER 1 INTRODUCTION .................................................................................................... 1 SECTION 1..................................................................................................................................... 1 Overview of Fatty-Acid Biosynthesis......................................................................................... 1 The Biotin Molecule ................................................................................................................... 2 The History of Biotin .............................................................................................................. 2 Biotin in Nutrition ................................................................................................................... 3 Structure and Chemistry of Biotin .......................................................................................... 3 Biotin-Dependent Enzymes ........................................................................................................ 5 Acetyl-CoA Carboxylase, a Biotin-Dependent Carboxylase ..................................................... 6 Enzyme Biotinylation ............................................................................................................. 8 Structure of Biotin Carboxyl Carrier Protein .......................................................................... 9 Catalytic Mechanism of Biotin Carboxylase ........................................................................ 10 Structure of Biotin Carboxylase............................................................................................ 14 Catalytic Mechanism of Carboxyltransferase ....................................................................... 19 Structure of Carboxyltransferase .......................................................................................... 20 Regulation of Acetyl-CoA Carboxylase ............................................................................... 23 Acetyl-CoA Carboxylase in Agriculture and Medicine ........................................................... 26 Acetyl-CoA Carboxylase as a Herbicide Target ................................................................... 26 The United States’ Obesity Epidemic ................................................................................... 28 Acetyl-CoA Carboxylase in Chronic Diseases ..................................................................... 28 Need For New Antibiotics — Acetyl-CoA Carboxylase a Valid Target.............................. 30 SECTION 2................................................................................................................................... 35 Some Metabolic Enzymes Bind Nucleic Acids ........................................................................ 35 The Types and Functions of the Zinc Fingers .......................................................................... 40 The Tetracyclines, Additional Inhibitors of Carboxyltransferase ............................................ 45 Objectives and Rationale for Study .......................................................................................... 47 References ................................................................................................................................. 48 CHAPTER 2 LINKAGE BETWEEN NUCLEIC ACID BINDING AND CATALYSIS IN THE
CARBOXYLTRANSFERASE SUBUNIT OF ACETYL-COA CARBOXYLASE ................... 65 Introduction ............................................................................................................................... 65 Materials and Methods.............................................................................................................. 68 Purification and Enzymatic Assay of Carboxyltransferase .................................................. 68 Substrate Nucleic Acids ........................................................................................................ 69 Eletrophoretic Mobility Shift Assay (EMSA) ...................................................................... 70 Data Analysis ........................................................................................................................ 71 Results ....................................................................................................................................... 73 DNA Binding by Carboxyltransferase .................................................................................. 73 RNA Binding by Carboxyltransferase .................................................................................. 81 iii
Inhibition of Enzyme Activity .............................................................................................. 82 Discussion ................................................................................................................................. 88 DNA Binding Enzymes ........................................................................................................ 88 Zinc Domains Associated with Proteins Involved in DNA Metabolism .............................. 91 Does Carboxyltransferase Bind DNA in Vivo? .................................................................... 92 The Mode of DNA Binding Suggests Communication Between the Dual Active Sites ...... 94 Pharmaceutical Relevance .................................................................................................... 95 References ................................................................................................................................. 96 CHAPTER 3 INHIBITION OF THE CARBOXYLTRANSFERASE SUBUNIT OF ACETYLCOA CARBOXYLASE FROM ESCHERICHIA COLI AND STAPHYLOCOCCUS AUREUS
BY TETRACYCLINES.............................................................................................................. 100 Introduction ............................................................................................................................. 100 Materials and Methods............................................................................................................ 102 Purification and Enzymatic Assay of Carboxyltransferase ................................................ 102 Electrophoretic Mobility Shift Assays ................................................................................ 104 Data Analysis ...................................................................................................................... 104 Results and Discussion ........................................................................................................... 105 Tetracyclines Inhibit Carboxyltransferase Activity ............................................................ 105 Effect of Tetracycline on DNA Binding ............................................................................. 111 Does Tetracycline Bind Carboxyltransferase in Vivo? ...................................................... 113 Pharmaceutical Relevance .................................................................................................. 119 References ............................................................................................................................... 119 CHAPTER 4 CONCLUSION .................................................................................................... 122 Subsequent Studies ................................................................................................................. 123 Future Directions .................................................................................................................... 125 References ............................................................................................................................... 127 APPENDIX A LETTER OF PERMISSION .............................................................................. 129 APPENDIX B ABBREVIATIONS ............................................................................................ 130 APPENDIX C ANALYTICAL TECHNIQUES APPLIED TO NUCLEIC-ACID BINDING
PROTEINS ................................................................................................................................. 132 References ............................................................................................................................... 134 VITA ........................................................................................................................................... 135 iv
ABSTRACT
Acetyl-CoA carboxylase is an essential enzyme, as it catalyzes the first committed and
regulated step in fatty-acid biosynthesis in all organisms excepting few Archaea and Eubacteria.
Acetyl-CoA carboxylase from gram-negative and gram-positive bacteria is a multifunctional
enzyme composed of three separate proteins. The carboxyltransferase subunit catalyzes the
transfer of a carboxyl group from carboxybiotin to acetyl-CoA, forming malonyl-CoA. The
crystal structure of the Escherichia coli (E. coli) carboxyltransferase component of acetyl-CoA
carboxylase revealed a unique Zn-domain, presumed to mediate nucleic acid binding, that is
absent in the eukaryotic enzyme. Notably, the Zn-domain, adjacent to the active site of
carboxyltransferase, makes for a unique target in the development of novel antibiotics capable of
highly specific binding. Utilizing an Electrophoretic Mobility Shift Assay as part of this study,
we investigated the nonspecific nucleic-acid binding and substrate (malonyl-CoA and biocytin)
inhibition of DNA:carboxyltransferase complex formation. Inhibition of carboxyltransferase
activity by single-stranded DNA, double-stranded DNA, RNA, and heparin was measured in the
reverse direction with a spectrophotometric assay in which the production of acetyl-CoA was
coupled with the combined citrate synthase-malate dehydrogenase reaction requiring NAD+
reduction (Blanchard and Waldrop, 1998). NADH formation was followed
spectrophotometrically at 340 nm. We then determined and characterized the mechanism of
inhibition by tetracycline (and derivatives) on carboxyltransferase from E. coli and
Staphylococcus aureus. The tetracyclines are broad-spectrum antibiotics that inhibit translation
by binding to the 30S ribosomal subunit and preventing the binding of the acylated-tRNA to the
A-site. Tetracycline exhibited competitive inhibition with respect to both malonyl-CoA and
v
biocytin. Multiple inhibition analyses with a bisubstrate analog showed that tetracycline and the
substrates can bind to the enzyme simultaneously. Surprisingly, tetracycline did not interfere
with the DNA-binding properties of carboxyltransferase. This introduction begins with a
historical perspective of carboxylation reactions. Next biotin and the structure, function and
practical applications of acetyl-CoA carboxylase are described. Subsequently a review of
moonlighting enzymes, or those capable of catalyzing reactions in basic metabolism while acting
as regulators of gene expression, is provided, as are the functions and structures of several types
of zinc finger.
vi
CHAPTER 1
INTRODUCTION
SECTION 1
Overview of Fatty-Acid Biosynthesis
Most organisms utilize fatty acids in the construction of cell membranes, and
higher organisms also use them for energy storage, and, the biosynthesis occurs via one
of two potential routes. Large, multifunctional enzyme complexes, called Type I fattyacid synthase or FAS-I (Wakil, 1962), are responsible for synthesizing fatty acids in
animal cells, in the cytosol of plants and in yeasts (Chang and Hammes, 1989; Singh et
al., 1985). However, in prokaryotes and plant chloroplasts, separated enzymes
collectively referred to as FAS-II (Cronan and Waldrop, 2002) catalyze the same
reactions. Mycobacterium tuberculosis utilizes both pathways, FAS-I and FAS-II (Bhatt
et al., 2007; Kurth et al., 2009). Excepting the thermoacidophilic archaeon
Metallosphaera sedula and autotrophic Archaea Sulfolobus metallicus and Acidianus
infernus, Archaea do not appear to synthesize fatty acids, as most organisms in that
domain have membranes made of prenylated ether lipids (Boucher et al., 2004; Hayes
2000) and altogether lack small biotin-carrying proteins (Menendez et al., 1999)
necessary for biosynthesis of fatty acids. Parasitic bacteria of the genus Mycoplasma do
not possess a fatty-acid biosynthetic pathway and do not synthesize fatty acids de novo.
Instead, their source of fatty acids is derived exclusively from their immediate
environment and must be imported (Romono et al., 1976; McAllister et al., 2006).
An essential substrate in both FAS-I and -II pathways is malonyl-CoA. The
enzyme that synthesizes malonyl-CoA, acetyl-CoA carboxylase, is the focus of this
1
study. The chemistry surrounding the acetyl-CoA carboxylase catalyzed reaction centers
on the cofactor biotin; thus, this molecule is described before the enzyme is discussed.
The Biotin Molecule
Biotin (also called vitamin H or B7), an essential vitamin, acts as a cofactor for a
group of enzymes catalyzing the transfer of carboxyl groups. Biotin-dependent enzymes
participate in metabolic reactions, including sodium transport, gluconeogenesis, urea
degradation, amino-acid catabolism, and fatty-acid synthesis and degradation (Samols et
al., 1988).
The History of Biotin
The biotin molecule was discovered in the 1920s when M.A. Boas found that rats
fed a diet rich in raw egg whites developed skin rashes and lost their fur, prior to
becoming paralyzed (Boas 1927; Leitner 1948). It is now known that egg whites contain
avidin, a protein that irreversibly binds biotin with a Kd of 10-15 mol/L (Bonjour, 1977;
Green 1975; Roth, 1985). In 1936, biotin was isolated from egg yolks and shown to be a
growth factor for yeast (Kögl and Tönnis, 1936). However, not until 1949 did its function
in carbon dioxide metabolism begin to be understood. That year, Lardy and co-workers
observed that Lactobacillus arabinosum incorporated 14CO2 into aspartate only when
biotin was present in the growth media (Lardy et al., 1949).
The function of biotin was further supported after observing biotin-deficient rats
fix lower levels of 14CO2 into cellular metabolites (i.e., aspartate and citrate) (MacLeod
and Lardy, 1949). Subsequently, in 1950, researchers observed the biotin-dependent
incorporation of 13CO2 into oxaloacetate in cell-free extracts of Micrococcus
2
lysodeikticus (Wessman and Werkman, 1950). An association between biotin and an
enzyme was first discovered when purified avian liver acetyl-CoA carboxylase was found
to contain biotin (Wakil et al., 1958), suggesting an enzyme cofactor role for biotin.
Biotin in Nutrition
Biotin is synthesized by plants, most bacteria and some fungi. The prevalence of
biotin in virtually every food, in conjunction with its synthesis and secretion via normal
gastrointestinal flora, makes biotin deficiency extremely rare. Therefore, few diseases
occur as a result of biotin deficiency; however, protracted parenteral nutrition therapy or
malnutrition can lead to severe biotin deficiency. Furthermore, an autosomal recessive
disorder traced to mutations in the genes coding for biotinidase and biotin ligase, two
enzymes involved in biotin metabolism (Nyhan, 1988), can lead to profound biotin
deficiency. Sustained biotin deficiency can lead to glucose intolerance (Bender, 1999)
and is considered teratogenic in mammals (Mock et al., 2002).
Structure and Chemistry of Biotin
Biotin carries labile carboxyl groups at the 1′ nitrogen (Figure 1-1). Although Du
Vigneaud in 1940 worked out the coenzyme chemical formula -- C10H16N2O3S -- and
suspected a complicated two-ring structure, the complete structure of biotin was not
determined until 1942 (Melville et al., 1942) and later reproduced through its chemical
synthesis (Harris et al., 1943). The biotin molecule contains three functional domains: a
ureido ring attached to a tetrahydrothiophene ring (fused cis to one another) and a valeric
acid side chain (also in cis, with respect to the ureido ring) (Melville et al., 1942). Biotin
is covalently attached to biotin-dependent enzymes via an amide linkage between the
3
valeric side chain and the ε-amino group of a specific, highly conserved lysine residue.
Although biotin contains three chiral carbon atoms, only the d-(+)-biotin isomer is
biologically active.
O
HN
2'
1'
3'
4
3
5
1
NH
OH
2
S
O
Figure 1-1. Biotin Molecule
H
The carboxyl-group-carrying capacity
of biotin initially was confirmed when a
S
HN
OH
methyl ester derivative of carboxybiotin, isolated from the reaction catalyzed by βmethylcrotonyl-CoA carboxylase,
was observed to be carboxylated
at the 1′ nitrogen
O
N
O
H
H
(Lynen et al., 1961). Subsequently, Lane’sbiotin
group demonstrated that chemically
synthesized biotin carboxylated at the 1′ nitrogen position could be used as a substrate for
acetyl-CoA carboxylase (Guchhait et al., 1974a).
The cofactor biotin is deprotonated at the 1′ nitrogen and undergoes
tautomerization into an enol-like form and acts as a nucleophile (Bruice and Hegarty,
1970). Through high-resolution X-ray crystallographic studies on biotin and biotin
derivatives, researchers determined that enolization is important for carboxyl-transfer
reactions (Stallings and DeTitta, 1985). Specifically, a longer carbonyl bond on the
ureido ring and shorter C2′-N1′ and C2′-N3′ bonds suggests a polarized ureido ring that
can interact with ions and polar compounds.
4
Biotin-Dependent Enzymes
All biotin-dependent enzymes follow ping-pong kinetics (Wood and Barden,
1977; Easterbrook et al., 1978), having two discrete steps, or half-reactions. Each step is
catalyzed at a separate active site, and biotin transfers the carboxyl group between active
sites of the enzyme. The half-reactions catalyzed by biotin-dependent enzymes are as
follows: First, the carboxyl group is transferred from the donor to biotin, forming the
carboxybiotin intermediate, and, consequently, the carboxyl group from carboxybiotin is
passed to the carboxyl acceptor. The enzymes are further categorized according to
whether they catalyze the fixation of CO2 (Class I, Figure 1-2), the decarboxylation with
release of CO2 as part of bicarbonate (Class II), or the transfer of a carboxyl group from
one molecule to another (Class III) (Moss and Lane, 1971).
Carboxylases, or Class I biotin-dependent enzymes, represent the largest class of
biotin-dependent enzymes and are the only biotin-dependent enzymes in eukaryotes.
Biotin-dependent carboxylases use bicarbonate as the carbon dioxide source, and fixation
of carbon requires hydrolysis of one molecule of ATP. Six examples characterize the
class: pyruvate carboxylase in gluconeogenesis, β-methylcrotonyl-CoA carboxylase in
amino-acid catabolism, acetyl-CoA carboxylase in fatty-acid synthesis, propionyl-CoA
carboxylase in fatty-acid oxidation, geranyl-CoA carboxylase in isoprenoid catabolism,
and urea carboxylase in urea catabolism (Wood and Barden, 1977).
Decarboxylases, or Class II biotin-dependent enzymes, are limited to the
membranes of anaerobic prokaryotes. There, they couple the free energy produced from
decarboxylation with the transport of sodium ions into the periplasm. Concomitantly, the
electrochemical gradient is used to synthesize ATP (Jitrapakdee and Wallace, 2003), and
5
bicarbonate is released. These enzymes are named according to the molecule that donates
the carboxyl group, i.e.: oxaloacetate decarboxylase, methylmalonyl-CoA decarboxylase,
and glutaconyl-CoA decarboxylase.
Transcarboxylase, or Class III biotin-dependent enzyme, represent the smallest
group, as the only known enzyme of this class is transcarboxylase. Biotin-dependent
transcarboxylase is found in the prokaryote Propionibacterium shermanii, in which it
catalyzes a carboxyl transfer from methylmalonyl-CoA to pyruvate in propionic acid
synthesis (Gerwin et al., 1969).
(I)
HCO3-
+ ATP-Mg
2+
Mg2+
ADP- Mg2+ + Pi + RCO2- + H+ (carboxylation)
+ RH
(II) RCO2- + H2O + 2 Na+intracellular
RH + HCO3- + Na+extracellular (decarboxylation)
(III) RCO2- + R’H
RH + R’CO2- (carboxyl transfer)
Figure 1-2. Net Reactions Catalyzed by Biotin-Dependent Enzymes
Acetyl-CoA Carboxylase, a Biotin-Dependent Carboxylase
Acetyl-CoA carboxylase catalyzes the first and committed step of the FAS-I and
FAS-II pathways, which makes it essential for de novo synthesis of long-chain fatty
acids, and is present in all plants, animals and nearly all bacteria. In a two-step reaction,
shown in Figure 1-3, the enzyme produces malonyl-CoA from acetyl-CoA in the
presence of bicarbonate and ATP (Polakis et al., 1974).
(1) Enzyme-biotin + MgATP + HCO3
-
Mg2+
(2) Enzyme-biotin-CO2- + Acetyl-CoA
Enzyme-biotin-CO2- + MgADP + Pi
Malonyl-CoA + Enzyme-biotin
Figure 1-3. Acetyl-CoA Carboxylase-Catalyzed Reaction.
6
The activation of bicarbonate by ATP produces a reactive acyl phosphate
intermediate called carboxyphosphate. In the presence of biotin, carboxyphosphate
decomposes and carboxybiotin is formed; this first half-reaction is catalyzed by the biotin
carboxylase component. The second half-reaction whereby the carboxyl group is
transferred from carboxybiotin to acetyl-CoA is catalyzed by the carboxyltransferase
component. The third component of the enzyme, biotin carboxyl carrier protein, contains
conserved lysine residue to which biotin is covalently attached. In eukaryotes the enzyme
exists as a single, multifunctional polypeptide chain (Lane et al., 1974). However,
studying eukaryotic acetyl-CoA carboxylase and establishing the catalytic mechanism
has proved difficult, as this form of the enzyme does not bind free biotin and purification
of the multi-enzymatic complex is arduous.
Though some bacteria, e.g., M. tuberculosis (Norman et al., 1994) and
Myxococcus xanthus (Kimura et al., 2000), have the biotin carboxyl carrier protein and
biotin carboxylase components on one polypeptide chain, prokaryotic acetyl-CoA
carboxylase exists as separable subunits (Guchhait et al., 1974b). In contrast with the
eukaryotic form of the enzyme, not only do the catalytic subunits of prokaryotic acetylCoA carboxylase remain active in the absence of the other enzymatic components, the
two catalytic subunits are able to recognize free biotin as a substrate. All subunits of
bacterial acetyl-CoA carboxylase can be easily studied, because the genes for biotin
carboxylase (Li and Cronan, 1992a, b) and carboxyltransferase (Blanchard and Waldrop,
1998) have been cloned, they can be overexpressed and their gene products purified.
Furthermore, the molecular mass, oligomerization state and Kd (in the biotin-ligating
reaction) have been dertermined for biotin carboxyl carrier protein (Nenortas and
7
Beckett, 1996), and an assay exists for both biotin carboxylase (Blanchard et al., 1999)
and carboxyltransferase (Blanchard and Waldrop, 1998). Moreover, crystal structures for
bacterial biotin carboxylase (Waldrop et al., 1994; Thoden et al., 2000a),
carboxyltransferase (Bilder et al., 2006) and biotin carboxyl carrier protein (Athappilly et
al., 1995) have been solved to 2.0 Å resolution or less. These breakthroughs have
improved our understanding of the relationship between the structure and function of
acetyl-CoA carboxylase in E. coli and, as a result, have offered insight into the catalytic
mechanism of other biotin-dependent enzymes.
Enzyme Biotinylation
Biotin-dependent enzymes are post-translationally modified via the covalent
attachment of biotin to the carrier subunit. This is mediated by the enzyme biotin protein
ligase, also called BirA or biotin holoenzyme synthetase (Chapman-Smith and Cronan,
1999). The linkage was elucidated after biocytin was isolated from yeast extract as ε-Nbiotinyl-L-lysine; thus, a lysine residue was presumed to be the site of attachment
(Wright et al., 1952). The site of biotin attachment in the target enzyme occurs in the
conserved peptide sequence containing Ala-Met-Lys-Met (Samols et al., 1988). Zhao and
co-workers (Zhao et al., 2009) proposed that, in a two-step mechanism, biotin ligase
binds biotin and ATP to catalyze synthesis of bio-5′-AMP, while pyrophosphate is
released. Secondly, biotin is activated via adenylation then transferred to the ε-amino
group of the lysine residue located 35 residues from the carboxyl terminus of biotin
carboxyl carrier protein (Choi-Rhee & Cronan, 2003; Lane et al., 1964).
8
Structure of Biotin Carboxyl Carrier Protein
Biotin carboxyl carrier protein from E. coli, shown in Figure 1-4, is a 16.7 kDa
protein and in bacteria is generally a separate subunit; yet it is always essential for the in
vivo reaction catalyzed by acetyl-CoA carboxylase (Choi-Rhee & Cronan, 2003). The
crystal structure of the C-terminus of biotin carboxyl carrier protein originally was solved
to 1.8 Å resolution, and the overall fold was described as a capped β-sandwich with
quasi-dyad symmetry, with each half containing a characteristic hammerhead motif
(Athappilly et al., 1995). A subsequent multidimensional NMR analysis of the same
region was used to validate the protein crystal structure (Yao et al., 1997). Aside from the
biotin-carrying domain, the remainder of the protein consists of a proline-/alanine-rich
sequence acting as a mobile linker (Cronan, 2002), with no other known function, and an
N-terminal domain that is involved in dimerization and for interaction with biotin ligase,
biotin carboxylase and carboxyltransferase (Yao et al., 1997). The stoichiometry of the
biotin carboxylase:biotin carboxyl carrier protein complex was determined to be a dimer
of biotin carboxylase plus four biotin carboxyl carrier protein molecules (Choi-Rhee and
Cronan, 2003).
Catalytic Mechanism of Biotin Carboxylase
The reaction catalyzed by biotin carboxylase requires bicarbonate, ATP, biotin,
and a large free-energy hurdle because a) bicarbonate is a poor electrophile and b) the 1′
nitrogen on biotin is weakly nucleophilic. In the course of elucidating the mechanism for
biotin carboxylase, the first consideration became the fate of ATP. It was postulated that
one of two possible scenarios existed -- the first involving ATP interacting directly with
9
biotin, forming an O-phosphobiotin, or, alternatively, ATP interacting with bicarbonate,
forming carboxyphosphate (Figure 1-5). Though, Calvin, Pon, and Lynen suggested the
formation of an O-phosphobiotin intermediate that could interact with bicarbonate to
form carboxybiotin (Calvin and Pon, 1959; Lynen et al., 1961), the currently accepted
model involves a carboxyphosphate intermediate.
Figure 1-4. Ribbon Rendering of the Carboxy-Terminal Fragment of the Biotin Carboxyl
Carrier Protein subunit of acetyl-CoA carboxylase from E. coli, with the lysine residue at
position 122 shown in color. (Image courtesy of Tyler Broussard)
O
O
-O
+ ATP
HO
O-
+ ADP
P
HO
O-
bicarbonate
O
O
carboxyphosphate
Figure 1-5. Bicarbonate and ATP Form Carboxyphosphate.
10
Carboxyphosphate is said to have a half-life near 70 msec (Sauers et al., 1975);
thus, trapping it seemed difficult. However, two groups have evidence of the existence of
this intermediate. While working with carbamyl phosphate synthetase (an enzyme that
catalyzes the formation of carbamyl phosphate using ATP, bicarbonate, and ammonia),
Wimmer and co-workers used a method suitable for following a reaction whereby ATP
reversibly phosphorylates an acceptor molecule. The method was called positional
isotope exchange (PIX) and allowed for the observation of the bridge-oxygen to
nonbridge-oxygen interactions during the rotation of the β-phosphate (Wimmer et al.,
1979). As bicarbonate and ATP are used in the reaction, the initial chemistry is thought to
be nearly identical to the mechanism for biotin carboxylase and by inference is applied to
biotin carboxylase. In a similar example, also while studying carbamyl phosphate
synthetase, Powers and Meister trapped carboxyphosphate using a diazomethane
quencher (Powers and Meister, 1976). This reaction was conducted via the incubation of
carbamyl phosphate synthetase, H13CO3- and ATP, and the reaction was methylated with
diazomethane; the resulting 13C product co-chromatographed with trimethyl
carboxyphosphate. Moreover, while repeating the assay in the presence of labeled ATP,
molar ratios of the two isotopes were nearly 1-1. Together, these experiments provide
evidence for the phosphorylation of bicarbonate by ATP.
In another experiment, researchers—aware that acetyl-CoA carboxylase was
capable of catalyzing the phosphorylation of ADP if the reaction was run opposite of the
physiological direction—used an analog of carboxyphosphate, carbamyl phosphate, and
observed the formation of ATP (Polakis et al., 1972; Polakis et al., 1974; Ashman and
11
Keech, 1975). Furthermore, the enzyme is capable of slow bicarbonate-mediated ATPhydrolysis activity in the absence of biotin (Levert et al., 2000).
The next and possibly strongest piece of evidence supporting the formation of
carboxyphosphate as an intermediate in the formation of carboxybiotin came from an
experiment using HC18O 3 while studying propionyl-CoA carboxylase and biotin
carboxylase. Investigators noted an exchange of 18O from bicarbonate to Pi, suggesting
direct contact between bicarbonate and the γ-phosphate of ATP (Kaziro et al., 1962;
Ogita and Knowles, 1988).
Prior to the subsequent step in the reaction mechanism, i.e. carboxylation of
biotin, researchers suspected carboxyphosphate either dissociates to carbon dioxide (and
Pi) before being bound by biotin or that biotin attacks the carboxyl group of
carboxyphosphate. In 1998, Gibson and co-workers attempted to determine the presence
of carbon dioxide as an intermediate in the reaction catalyzed by carbamyl phosphate
synthetase (Gibson et al., 1998). This enzyme catalyzes the formation of
carboxyphosphate, and, in the absence of a nitrogen source, carboxyphosphate breaks
down as bicarbonate and Pi. They measured the rate of ADP formation as an indicator of
reaction progress while recording pH. If carboxyphosphate undergoes water-mediated
hydrolysis, one proton is released for every molecule of carboxyphosphate hydrolyzed. If
the pH does not decrease as the reaction proceeds, carboxyphosphate decomposes to
carbon dioxide and Pi, with subsequent generation of bicarbonate and a proton in the
presence of water, leaving the pH unchanged. Their results suggested
carboxyphosphate—and not carbon dioxide—directly carboxylated biotin. However,
12
more recently, computational chemists Ito and co-workers have suggested the release of
CO2 from carboxyphosphate (Ito et al., 2008).
The final aspect of the biotin carboxylase reaction mechanism to address involves
the activation of biotin. For the N1′ of biotin to become nucleophilic, it must be
deprotonated. Tipton and Cleland first proposed the presence of an active-site acid base
pair in the activation of biotin (Tipton and Cleland, 1988). Specifically, an active-site
lysine residue was thought to remove the proton from the thiol group of an active-site
cysteine, while a thiolate anion subsequently removed the N1′ proton of biotin. The
protonated lysine residue was proposed to stabilize the negative charge at the ureido
oxygen. However, using site-directed mutagenesis, Levert and co-workers found that a
substitution of alanine in the place of the active-site cysteine (C230) produced an enzyme
with near-wild-type activity. In the same work, it was concluded that the lysine positions
carboxyphosphate for efficient carboxyl transfer to biotin (Levert et al., 2000). Levert
and co-workers suggested the phosphate oxygens of carboxyphosphate could act to
deprotonate the N1′ of biotin (Figure 1-6, interaction A). In agreement with this
hypothesis, Ito and co-workers state that biotin can undergo enolization in the presence of
carboxyphosphate, as this intermediate serves as a general acid-base catalyst (Ito et al.,
2008), and the carboxyl group donates a proton to the ureido carbonyl (Figure 1-6,
interaction B). Moreover, Ito and co-workers suggest that carboxyphosphate collapses
into CO2 and Pi (Figure 1-6, interaction C) with the resulting “highly labile, bent” CO2
molecule rapidly condensing (with enolic biotin) into carboxybiotin (Figure 1-6,
interaction D).
13
B
O
O-
H
D
O
C
O
NH
N
O
H
P
HO
BCCP
O-
A
carboxyphosphate
S
biotinyl-BCCP
O
Figure 1-6. Activation of Biotinyl-BCCP via Carboxyphosphate.
Structure of Biotin Carboxylase
The crystal structure of E. coli biotin carboxylase originally solved to 2.4 Å
resolution (Waldrop et al., 1994) confirmed a homodimer of 50 kilodalton polypeptide
chains, which was first suggested by size-exclusion chromatography (Dimroth et al.,
1970). Waldrop et al. described the enzyme as having an overall compressed structure
divided into three structural domains, which are shown in Figure 1-7 and detailed as
follows:
1. The N-terminal domain (also called the A-domain), containing a dinucleotide
binding motif, has five strands of parallel β-pleated sheets with four α-helices
on two sides. The A-domain extends to Ile-103. The A-domain has a minor
contribution to subunit dimerization. Dividing the A- and B-domains is a
helix-turn-helix motif, extending from Ala-107 to Ala-126. Both α-helices in
this region are connected by Asp-115 and are oriented perpendicular to one
another.
14
2. The C-terminal (or C-domain) is composed of an eight-stranded antiparallel βpleated sheet with a three-stranded antiparallel β-pleated sheet and seven αhelices. This domain starts with Arg-208, is the largest domain and also
contains the residues that are most involved in subunit dimerization. The
initial structure revealed several crystal contacts within surface loops starting
at alanine 341 and ending at proline 351.
3. The central (or ATP-grasp) domain consists of two α-helices and three
antiparallel β-pleated sheets. This domain, also called the B-domain,
beginning at Val-131 to Tyr-203, projected outward from the rest of the
protein in the initial crystal structure of biotin carboxylase, while possessing
some disordered surface loops. Additionally, the absence of the X-ray
diffraction data in the glycine-rich region of the polypeptide between the Bdomain and the main body of the protein support the likely presence of a
highly flexible linker region. A second crystal structure of biotin carboxylase,
with an active-site mutation and in complex with ATP, was solved to 1.9 Å
resolution (Thoden et al., 2000a) and showed the B-domain clamped down on
the body of the enzyme.
The authors of the second crystal structure described the B-domain as follows:
“The major conformational change that occurs upon nucleotide binding is a rotation of
approximately 45° of one domain relative to the other domains thereby closing off the
active site pocket” (Thoden et al., 2000a); thus, the B-domain was proposed to act as a lid
that closes down on the active site after the substrates are bound and positioned for
catalysis. Another group, having solved a crystal structure of wild-type biotin
15
carboxylase in complex with biotin, bicarbonate and Mg-ADP, suggested the biotin
molecule remains solvent exposed after closure and that biotin carboxyl carrier protein is
needed to completely close the active site before catalysis (Chou et al., 2009). Results
from computational chemistry studies suggest that during this closure motion a small
change in twist angle also occurs (Novak et al., 2009). This large rotation about the hinge
region was supported by the co-crystal structure (with nucleotide analog 5′adenylylimidodiphosphate) of a structurally related enzyme also capable of binding
nucleotides, carbamyl phosphate synthetase, which revealed a similar closure (Thoden et
al., 1999; Thoden et al., 2000a).
Figure 1-7. Ribbon Rendering of the Biotin Carboxylase subunit of acetyl-CoA
carboxylase from E. coli, with one subunit in white and the other in color. (Image
courtesy of Tyler Broussard)
16
Yet researchers, having solved a crystal structure of biotin carboxylase in the
presence of ATP analogs, proposed that closing of the hinged region in one subunit
occurs in concert with the opening of the hinged region in the other subunit, as this
structure revealed only one ATP analog bound to a single active site at a time (Mochalkin
et al., 2008). Thus, these same authors suggest the catalytic cycle “flip-flops” between
active sites.
The closed conformation of the ATP-bound structure of biotin carboxylase
exposed that a number of residues converged on the active-site pocket: e.g., Lys-116,
Lys-159 and Met-169 (Thoden et al., 2000a). According to sequence alignments for
enzymes having ATP-grasp domains, these three residues appear highly conserved across
prokaryotic and eukaryotic genomes (Galperin and Koonin, 1997). The results of the
crystal structure solved by Mochalkin et al. depict Lys-116 and Lys-159 interacting with
ATP; these interactions are reinforced by the 50-fold (K116Q) and 90-fold (K159Q)
greater Km reported previously (Sloane et. al. 2001). Bordelon and co-workers suggest
two neighboring glycine residues (165 and 166) are involved in the binding and
positioning of ATP, while mutations of these residues appear to result in the
“misalignment of the reactants for optimal catalysis”; recent crystallographic data also
support this interaction (Bordelon et al., 2009; Mochalkin et al., 2008). Moreover, Chou
and co-workers suggest that two oxygens within the bicarbonate molecule participate in
ion-pair interactions with the guanidinium of arginine 292. They further explain, one of
these two bicarbonate oxygens hydrogen bond with the main chain of valine 295 and
glutamic acid at position 296 likely functions as a general base, extracting the proton
from the third oxygen of bicarbonate. Subsequently, valine 295 and a tyrosine at position
17
82 are said to be very near the biotin molecule, while an arginine at position 338 may be
involved in the bonding of hydrogen and stabilizing the carbonyl oxygen in the ureido
ring of biotin. However, this same arginine most importantly may function to stabilize the
enolate biotin transition state, as mutation to alanine at this position resulted in an
inactive enzyme (270-fold decrease in kcat) (Chou et al., 2009).
Nilsson and co-workers suggested two Mg2+ ions were critical to catalysis by
alleviating electrostatic repulsion (Nilsson Lill et al., 2008); this, too, has been supported
by recent crystallographic data (Mochalkin et al., 2008). The Mg2+ ions are coordinated
to the phosphates of ATP. Specifically, one Mg2+ atom bridges between the α- and γ–
phosphoryl groups of ATP, while the other Mg2+ atom coordinates one β- and one γphosphoryl oxygen of ATP, and glutamic acid residues at positions 274 and 288 play key
roles in bridging both metal ions, respectively (Mochalkin et al., 2008; Nilsson Lill et al.,
2008).
Subsequent to the discovery of the three-dimensional structure of biotin
carboxylase, researchers initially noticed analogous folds between this enzyme and two
peptide ligases, glutathione synthetase and a peptidoglycan biosynthesis enzyme Dalanine:D-alanine ligase (Artymiuk et al., 1996). The high similarity in the structure of
these enzymes is noteworthy, as they have only 11% primary sequence identity, yet they
catalyze similar reactions. The catalytic mechanism of these three enzymes couples the
hydrolysis of ATP to the formation of a carbon-nitrogen bond, and each of these
enzymatic mechanisms includes the same intermediate, an acyl phosphate (Artymiuk et
al., 1996; Galperin and Koonin, 1997).
18
Not long after these similarities were realized, the structure of carbamyl
phosphate synthetase was determined (Thoden et al., 1997). This enzyme seems to have
structural and mechanistic similarities to glutathione synthetase and D-alanine:D-alanine
ligase; however, it appears most similar to biotin carboxylase, as this enzyme also binds
ATP and bicarbonate as substrates, and the mechanism evolves through a
carboxyphosphate intermediate. Collectively, these enzymes represent the first members
of the ATP-grasp family of enzymes (Galperin and Koonin, 1997). To date, several
additional enzymes have been added to the ATP-grasp family, including all biotindependent carboxylases; biotin-dependent carboxylase domains of pyruvate carboxylase;
propionyl-CoA carboxylase; urea amidolyase; tubulin-tyrosine ligase; and three enzymes
of purine biosynthesis, including glycinamide ribonucleotide synthetase (Wang et al.,
1998), N5-carboxyaminoimidazole transformylase (Thoden et al., 1999), and
glycinamide ribonucleotide transformylase (Thoden et al., 2000b).
Catalytic Mechanism of Carboxyltransferase
The exact mechanism of the transfer of the carboxyl group from carboxybiotin to
acetyl-CoA in the reaction catalyzed by carboxyltransferase has yet to be elucidated, yet
the reaction involves an initial abstraction of a proton from the methyl group of acetylCoA with subsequent carboxyl-group transfer from carboxybiotin to acetyl-CoA. Bilder
et al. describe two glycine residues (206 and 207) that form an oxyanion hole that
stabilizes the negatively charged ureido enolate that develops within the biotin molecule
following decarboxylation. Likewise, upon removal of a proton from the methyl group of
acetyl-CoA, an enolate is created (Waldrop, 2011), thus stabilization of both enolate
19
anions likely contributes to the lowering of the activation energy for the enzymecatalyzed reaction. Unfortunately, to date, very few mutagenic studies on
carboxyltransferase have been done; however, these studies likely will provide
information toward elucidating the enzymatic mechanism.
Structure of Carboxyltransferase
In 2006, the long-awaited initial crystal structure of carboxyltransferase from E.
coli and S. aureus, shown in Figure 1-8A, was solved and revealed a surprising Zndomain not seen in the eukaryotic homolog (Bilder et al., 2006).
Figure 1-8A. Ribbon Rendering of the Carboxyltransferase subunit of Acetyl-CoA
Carboxylase from E. coli, the beta subunit in white and the alpha subunit in color, with a
Close-up of Zinc Domain (1-8B). (Images courtesy of Tyler Broussard and Pat Bilder,
respectively)
20
(Figure 1-8 continued)
The overall structure of the α2β2 tetramer, or dimeric dimer, of the functional
enzyme has been referred to as a truncated rectangular pyramid. The structurally
homologous α- and β-subunits also assume wedge shapes, and a tapering core, initially 13
Å extending to 23 Å, is presumed to bind the substrates. The quaternary structure
validated the α2β2 subunit composition suggested by Lane and colleagues (Guchhait et
al., 1974b), while structural homology to enoyl-CoA hydratase and the presence of an
oxyanion hole in the β-subunit for CoA-thioenolate stabilization showed that the enzymes
belong to the crotonase superfamily (Bilder et al., 2006; Gerlt and Babbitt, 2001; Murzin,
1998).
The active site of carboxyltransferase most closely resembles the “ridges in
groves” stacking motif, whereby a six-helix bundle stabilizes the interconnection of
subunits and forms a catalytic platform. Additionally, two helical domains from the αsubunit and a Zn-domain from the β-subunit combine to form a porous catalytic lid over
the catalytic platform. The holes in the catalytic lid have been suggested to act as entry
21
points for substrates. By homology to propionyl-CoA carboxylase from Streptomyces
coelicolor (Diacovich et al., 2004), it has been suggested that residues G206 and G207
(near the N-terminus of the eighth alpha helix) and G204 – G205 (in the β-subunit)
contribute to the formation of a conserved oxyanion holes. These oxyanion holes are
thought to stabilize the ureido enolate that forms upon decarboxylation of carboxybiotin
at the 1′ nitrogen by acting as H-bond donors to the carbonyl and enolate intermediate in
propionyl-CoA carboxylase (Diacovich et al., 2004); a similar function is expected of the
conserved residues in carboxyltransferase. The overall fold, not surprisingly, is similar to
that of carboxyltransferase from yeast (Zhang et al., 2003) and S. coelicolor (Diacovich
et al., 2004). However, when the gene for the β-subunit of E. coli carboxyltransferase
was cloned and sequenced more than 20 years ago (Bognar et al., 1987), the authors
noted the tandem C-X-X-C sequences separated by 15 residues near the amino terminus
and hypothesized that the protein had metal-binding domains homologous to nucleic
acid-binding proteins.
The crystal structures of carboxyltransferase from S. aureus and E. coli, along
with X-ray fluorescence studies, have confirmed this prediction. The metal atom is zinc,
and it forms part of a zinc motif that is unique to bacterial carboxyltransferase (Figure 18, B). A space-filling representation reveals that the Zn-domain, specifically classified as
an atypical zinc ribbon, forms part of a saddlelike structure, and an electrostatic surfacepotential rendering shows that the inner face of the zinc-finger domain has an
electropositive surface potential, while most of the protein has an electronegative surface
potential. Sequence alignment of the primary sequence of this zinc finger (versus
orthologous enzymes from Eubacteria, algae, plants, and animals) suggests the existence
22
in prokaryotes alone. This zinc ribbon is thought to most closely resemble those of
ribosomal proteins (Bilder et al., 2006); a brief explanation of zinc fingers in the context
of carboxyl transferases is provided in the second section of this introduction.
Regulation of Acetyl-CoA Carboxylase
Because acetyl-CoA carboxylase acts as the committed step in fatty-acid
biosynthesis in most kingdoms (Cronan and Waldrop, 2002), excepting the Archaea and
some mycoplasmas (McAllister et al., 2006), it is tightly regulated. The product of the
reaction catalyzed by acetyl-CoA carboxylase (malonyl-CoA) becomes the substrate for
the second enzyme in the pathway (fatty-acid synthase) in an energetically costly reaction
and should be tightly regulated. As such, different mechanisms of regulation exist in
prokaryotes versus eukaryotes. Here, the two processes are briefly explained.
In E. coli, the genes for biotin carboxylase and biotin carboxyl carrier protein are
co-transcribed as the accBC operon (Li and Cronan, 1992a). However, this gene
arrangement is not universal among prokaryotes, as several gram-positive bacteria
possess the acc genes as clustered with the fatty-acid synthetic genes (Cronan and
Waldrop, 2002). Researchers first noticed that bacteria harboring high copy number
plasmids (10 – 50 plasmids per cell) containing the accBC operon did not overproduce
proteins commensurate with the plasmid copies (Li and Cronan, 1993). They also
proposed that the replacement of the native promoter with a heterologous promoter
produced a “straightforward dependence on copy number,” as seen in protein production.
The results suggest the requirement for a transcriptional activator or that these proteins
repress their own synthesis (Cronan and Waldrop, 2002). Interestingly, biotin carboxyl
23
carrier protein was reported to be capable of suppressing transcription of the accBC
operon, suggesting an autoregulatory feedback mechanism of transcriptional control
(James and Cronan, 2004).
However, Li and Cronan suggest an alternative regulatory mechanism exists in
the expression of carboxyltransferase in E. coli, and this process remains poorly
understood (Li and Cronan, 1993). They agree that both genes for carboxyltransferase
reside in complex gene clusters that are part of vital biosynthetic processes (5'-ORF17lpx4-lpxB-ORF23-polC-accA-3' and 5'-ubxI-hisTdedA- accD-folC-dedD-dedE-purFdedF-argT-3', respectively) and are likely cotranscribed. Interestingly, I reported the
specific binding of carboxyltransferase to mRNA encoding the α and β subunits (Meades
et al., 2010). This information may help to explain the vastly different expression levels
observed between accBC and accA-accD-specific mRNA reported (Li and Cronan,
1993). These same authors reported a high copy number of accA-accD-specific mRNA
was detected in bacterial strains harboring the high copy number plasmid. This may be
suggestive of a translational regulatory mechanism; thus, it remains possible for the two
operons containing the genes for accA-accD to be expressed at a high level, while high
concentrations of carboxyltransferase shift equilibrium toward enzyme:mRNA complex
and translation suppression. Aside from regulation for acetyl-CoA carboxylase synthesis,
other mechanisms for regulation exist, i.e. the regulation of enzymatic activity.
Though acetyl-CoA carboxylase was observed to be inhibited in vitro and thought
to be regulated by high levels of (p)ppGpp, a molecule said to accumulate under
conditions of amino-acid starvation, it since has been realized that this process is
physiologically irrelevant (Cronan and Waldrop, 2002). However, acyl carrier protein
24
serves as the hub for the pathway, as it tethers the nascent fatty acyl chain until it
dissociates as palmitic acid. In 2001, Davis and co-workers, while studying E. coli,
suggested acylated derivatives of this protein partially inhibit acetyl-CoA carboxylase.
Though the inhibition never reached 100%, E. coli acyl carrier protein was specifically
required, as acylated derivatives of the homologous protein from spinach had no effect.
Interestingly, acyl chain lengths between C6 and C20 had equal inhibitory effect, and
acyl carrier protein was reported to inhibit the overall reaction but not either partial
reaction. It is hypothesized that this feedback mechanism acts to conserve energy and as a
key metabolic intermediate (acetyl-CoA) in the presence of sufficient levels of fatty acid
(Davis and Cronan, 2001).
In mammals, two isoforms of acetyl-CoA carboxylase exist: ACC1 and ACC2
(Abu-Elheiga et al., 1995; Widmer J et al., 1996); the former is primarily expressed in
lipogenic tissues, i.e. kidney and liver, and the latter is expressed in muscle. Abu-Elheiga
and co-workers have shown that ACC1 is involved in fatty-acid biosynthesis and that
ACC2 is involved in inhibiting fatty-acid oxidation (Abu-Elheiga et al., 1997; Saddik et
al., 1993). Specifically, ACC2 increases levels of malonyl-CoA that inhibit carnitine
palmitoyl transferase-1 and terminate the delivery of fatty acids for oxidation (AbuElheiga et al., 1997; Saggerson, 2008).
Mammalian acetyl-CoA carboxylase can be regulated by several mechanisms. In
fasting organisms, acetyl-CoA carboxylase expression is low, yet it increases upon
feeding (Iritani, 1992). Mabrouk and co-workers reported acetyl-CoA carboxylase
isolated from rat livers after an insulin bolus had higher enzymatic activity; whereas, the
enzyme had a lower activity after administration of glucagon, or epinephrine (Mabrouk et
25
al., 1990). A high degree of polymerization of acetyl-CoA carboxylase has been found to
correspond to an increase in enzyme activity, and dephosphorylation by [acetyl-CoA
carboxylase]-phosphatase 2 leads to protomer polymerization (Thampy and Wakil,
1988). Conversely, one mechanism involved in the short-term reduction of the catalytic
activity of acetyl-CoA carboxylase is phosphorylation and subsequent depolymerization;
the enzyme AMP-activated protein kinase (SNF1 gene product in yeast) phosphorylates
acetyl-CoA carboxylase on multiple serine residues (Brownsey et al., 2006; Mitchelhill et
al., 1994; Woods et al., 1994). Another form of regulation in the human ACC1 gene
occurs via the expression of alternative exons. Three promoters (PI, II and III) for the
enzyme have been described, suggesting the possibility of three isoforms (Kim, 1997;
Mao et al., 2003). ACC1 is constitutively and highly expressed, PII is regulated by the
thyroid hormone, Triiodothyronine (T3), and PIII plays a critical role during lactation
needed for baby development (Mao et al., 2003).
Acetyl-CoA Carboxylase in Agriculture and Medicine
Acetyl-CoA Carboxylase as a Herbicide Target
Molecules from the chemical classes aryloxyphenoxypropionates (AOPPs),
cyclohexanediones (CHDs) and phenylpyrazolin inhibit acetyl-CoA carboxylase and are
used as selective herbicides on some rice, wheat and grass weeds (Burton et al., 1987;
Walker et al., 1988). Though variations in uptake, translocation, detoxification, and
activation contribute to herbicide selectivity in plants, the greatest determinant of
susceptibility lies in the type of the plant’s acetyl-CoA carboxylase (Harwood, 1988).
Most plants contain two forms of acetyl-CoA carboxylase, namely prokaryotic and
26
eukaryotic forms. The eukaryotic form of acetyl-CoA carboxylase is inhibited by these
herbicides, while the prokaryotic form is not. Plants lacking the prokaryotic form of the
enzyme are susceptible to these inhibitors (Konishi and Sasaki, 1994).
Inhibition studies of alloxydim, sethoxydim and clethodim, examples of the CHD
chemical class, and eukaryotic acetyl-CoA carboxylase showed half-maximal values
ranging from 0.12 to 5.0 µM (Rendina and Felts, 1988); whereas, the R isomer of
fluazifop has a half-maximal value of 4 µM (Walker et al., 1988). These same authors
noted a decrease in the chain length of fatty acids in the presence of fluazifop. These
investigators also showed specifically that the carboxyltransferase half-reaction was the
target of these herbicides (Rendina et al., 1989; Rendina and Felts, 1988; Walker et al.,
1988). More recently co-crystal structures were reported for yeast acetyl-CoA
carboxylase and haloxyfop (Zhang et al., 2004), tepraloxydim (Xiang et al., 2009), and
pinoxaden (a phenylpyrazolin) (Yu et al., 2010) and proved herbicide binding at the
carboxyltransferase active site.
Some plants that were once sensitive to these herbicides have become resistant.
Plant insensitivity is conferred by the following mechanism: (1) enhanced herbicide
detoxification (De Prado et al., 2004) and (2) point mutations in isoforms of acetyl-CoA
carboxylase (De Prado et al., 2004; Alarcon-Reverte and Moss, 2008; Parker et al.,
1990), which were described by one author as being controlled by a single dominant or
semi-dominant nuclear gene (Price et al., 2003). Researchers studying acetyl-CoA
carboxylase that was either inherently resistant or that developed insensitivity to these
herbicides described a critical isoleucine-to-leucine substitution (Zagnitko et al., 2001)
and report that some insensitive forms of the enzyme bind the herbicide in a co-operative
27
manner (Price et al., 2003). The utility of these herbicides, as well as the resistance
mechanisms, warrant future investigation and inhibitor design.
The United States’ Obesity Epidemic
The U.S. Centers for Disease Control and Prevention reported (Behavioral Risk
Factor Surveillance System and The Morbidity and Mortality Weekly Report, Centers for
Disease Control, 2006), “During the past 20 years there has been a dramatic increase in
obesity in the United States,” and, “In 2007, only one state (Colorado) had a prevalence
of obesity less than 20%. Thirty states had prevalence equal to or greater than 25%; three
of these states (Alabama, Mississippi, and Tennessee) had a prevalence of obesity equal
to or greater than 30%.” Having surpassed the threshold for epidemic status, finding ways
to treat obesity to avoid associated sequela, e.g. metabolic syndrome, hypertension,
hyperlipidemia, type II diabetes, cardiovascular disease and cancer, is paramount. The
vital role of acetyl-CoA carboxylase in long-chain fatty-acid synthesis makes it an
obvious target for therapeutic intervention in the context of a variety of diseases.
Consequently, it has been implicated as having a role in the treatment of obesity
(Magnard et al., 2002), and inhibiting acetyl-CoA carboxylase in humans may provide
for medically induced weight loss.
Acetyl-CoA Carboxylase in Chronic Diseases
Acetyl-CoA carboxylase has been considered a therapeutic target in the treatment
of cardiovascular risk factors (Harwood et al., 2003), cancer (Magnard et al., 2002) and
obesity (Levert et al., 2002). Proof of concept for this theory came indirectly as one study
demonstrated a reduction in body weight in mice treated with fatty-acid synthase
28
inhibitors (Loftus et al., 2000). Though one should anticipate fat loss when fatty-acid
synthase is inhibited, an interesting twist came from the proposed cause of the weight
loss. Researchers observed that these mice had reduced appetite and an increase in
malonyl-CoA, and the researchers suggested that excess malonyl-CoA is an appetite
suppressant (Lane and Cha, 2009). In a second study, mice lacking an isoform of acetylCoA carboxylase (ACC), specifically ACC2 (previously referred to as ((H))ACC-β), as a
result of genetic engineering, had lower levels of malonyl-CoA and were observed to lose
weight despite greater food intake than control mice, and these mice were reported to
have normal life spans (Abu-Elheiga et al., 2001). In the absence of malonyl-CoA, fattyacid oxidation outpaces fatty-acid synthesis, and fats are mobilized from hepatic and
adipose stores to muscle cells. Taken together, there appears to be a connection in
abnormal malonyl-CoA levels and weight loss. An unequivocal connection between
acetyl-CoA carboxylase and obesity exists, and it is reasonable to expect, in the near
future, anti-obesity therapy to target acetyl-CoA carboxylase.
The potential for improving the health of patients with cardiometabolic risk
factors, because of diabetes and metabolic syndrome, by inhibiting ACC2 shows promise.
In another knockout murine model, the results of mice treated with isozyme-specific
antisense oligonucleotides or with isozyme-nonselective acetyl-CoA carboxylase
inhibitors have demonstrated the potential for treating metabolic syndrome (Harwood,
2003). Interestingly, isozyme-nonselective acetyl-CoA carboxylase inhibitors seem to
show the most therapeutic potential (Corbett et al., 2007). This work has underscored the
valid role of acetyl-CoA carboxylase in the treatment of noninfectious diseases.
29
Need For New Antibiotics — Acetyl-CoA Carboxylase a Valid Target
The incidence of multidrug-resistant bacterial pathogens is rising at an alarming
rate (Chen et al., 2009). Previously limited to hospitals, common infections from
antibiotic-resistant, gram-positive (S. aureus and Streptococcus pneumoniae) and gramnegative (E. coli and Pseudomonas aeruginosa) bacteria have become common in
community settings as well (Menichetti, 2005). Clinicians remain concerned about the
emergence of the “ESKAPE” pathogens (Enterococcus faecium, S. aureus, Klebsiella
pneumoniae, Acinetobacter baumannii, P. aeruginosa, and Enterobacter species). The
acronym reflects the fact that these bacteria represent the majority of nosocomial
infections and effectively elude the antibacterial drug’s mechanism of action (Boucher et
al., 2009). Moreover, strains of gram-positive pathogenic bacteria are proving resistant to
our “last line of defense,” namely vancomycin. Here again, strains of S. aureus have
emerged resistant (Menichetti, 2005). Over the latter third of the last century, there was a
steep decline in the rate at which new antibiotics became available, and the effectiveness
of subsequent-generation antibiotics is shorter lived than parent, presently ineffective
antibiotics; thus, novel classes of both antibiotics and enzyme targets should be
considered.
Prior to the year 2000, the last novel class of antibiotics was trimethoprim,
released in 1968, and, though new antibiotics continued to be released, most were
permutations of natural products or chemical modifications of previously discovered
classes of drugs (Powers, 2004). Fortunately, in 2003 the United States Food and Drug
Administration approved the use of daptomycin for treating complicated skin and softtissue infections from gram-positive bacteria (Kern 2006). This molecule represents the
30
first in a new class of antibiotics called the cyclic lipopeptides and is described as having
a cationic antimicrobial peptide-like mechanism (Ernst et al., 2009), yet the mechanism
of action for this antibiotic is not completely understood. It is proposed to bind to the
cytoplasmic membrane via a calcium-dependent insertion of the hydrophobic tail and
oligomerization. Subsequently, the membrane is disrupted, resulting in efflux of
potassium, arrest of nucleic acid and protein synthesis and cell death (Canepari et al.,
1990; Silverman et al., 2003; Steenbergen et al., 2005; Shah, 2005). However, Ernst et al.
describe MprF, a bacterial defensin resistance protein, as having a critical function in
microbe virulence and being involved in resistance to daptomycin. Unfortunately, to date
drug-development strategies have included limited cellular targets; thus, it has become
imperative to design pharmaceuticals against additional metabolic processes.
To expand the cellular targets of antibiotics, bacterial enzymes involved in fattyacid synthesis (Type II) have materialized as important targets for drug design (Campbell
and Cronan, 2001; Heath et al., 1998; Marrakchi et al., 2002; Zhang et al., 2006). Two
such examples of inhibitors designed to target a bacterial FAS-II enzyme are the
antitubercular drugs isoniazid and ethionamide, which inhibit enoyl-ACP reductase.
Enoyl-ACP reductase reduces the trans double bond between positions C2 and C3 of a
fatty acyl chain linked to the acyl carrier protein. This reaction is essential in mycolic
acid (virulence factor) biosynthesis (Banerjee et al., 1994; Heath et al., 1998). Other
antibiotics, such as cerulenin and thiolactomycin (also classified as an antifungal and
antimalarial, respectively), inhibit beta ketoacyl-ACP synthase, which is involved in
FAS-II chain condensation (D’Agnolo et al., 1973; Hayashi et al., 1983). Unfortunately,
the structures of enoyl-ACP reductase and ketoacyl-ACP synthase are highly variable
31
across species, and high variability is thought to result in weaker binding and reduced
effectiveness in some species. Thus, these antibiotics are considered narrow spectrum
with limited advantages.
In contrast with the high sequence and structure variability seen in the fatty-acid
biosynthesis gene products, acetyl-CoA carboxylase is widely conserved among bacterial
species and may be a better antibiotic target. Interestingly, an inhibitor of one of the
many isoforms of acetyl-CoA carboxylase, namely AccD6, in Mycobacterium has been
described (Kurth et al., 2009). Additionally, andrimid and moiramide, a class of natural
products with a pseudopeptide pyrrolidinedione backbone, (Freiberg et al., 2004;
Pohlmann et al., 2005) inhibit the carboxyltransferase component, while a
pyridopyrimidine inhibitor targets the biotin carboxylase component (Miller et al., 2009);
thus, theoretically these antibiotics would have a synergistic therapeutic effect.
Andrimid is an antibiotic produced by Pantoea agglomerans (Liu et al., 2008),
and it directly targets the accD gene product (β-subunit of carboxyltransferase) in acetylCoA carboxylase. Resistance to andrimid in P. agglomerans (and likely Vibrio cholera)
is conferred via horizontal gene transfer or gene duplication of AccD. The duplicated
AccD, called AdmT, is actually part of the andrimid biosynthesis gene cluster and has
nearly 90% sequence similarity when compared to AccD. The AdmT gene product, when
mixed with the alpha subunit of carboxyltransferase, is capable of catalyzing the same
reaction as wild-type carboxyltransferase, and this hybrid enzyme has similar kinetic
parameters as the wild-type enzyme. Researchers have determined a critical active-site
residue (position 203) that contributes to increased resistance. Specifically, a methionineto-leucine change at this position in the wild-type beta subunit of carboxyltransferase
32
increases resistance to andrimid; while, in AdmT, a leucine-to-methionine change
increases sensitivity to the antibiotic. Methionine or leucine in this position are thought to
be important in the interaction with andrimid or the substrate acetyl-CoA, respectively
(Liu et al., 2008).
Miller and colleagues, while searching for new antibiotics, used a library
containing ~1.6 million compounds of mostly ATP-competitive ligands -- initially
intended to discover inhibitors for eukaryotic protein kinases (Miller et al., 2009). A
whole bacterial cell screening assay was deployed and identified a series of
pyridopyrimidines that proved to be bactericidal against E. coli, Haemophilus influenzae,
and Moraxella catarrhalis. A biological macromolecular biosynthesis assay revealed
fatty-acid synthesis was targeted by this inhibitor, and the interaction with biotin
carboxylase was confirmed using isothermal titration calorimetry and surface-plasmon
resonance (Miller et al., 2009). A co-crystal structure provided evidence that the inhibitor
binds to the ATP-binding site in biotin carboxylase, and a mutation (I437T) in the gene
for biotin carboxylase from E. coli conferred resistance to the inhibitor. Interestingly,
when this inhibitor was originally screened as a potential eukaryotic protein kinase
inhibitor, it was found to be only weakly active toward Src eukaryotic protein kinase.
This inhibitor has at least a 70-fold lower affinity for Src protein kinase and eukaryotic
acetyl-CoA carboxylase. Moreover, this inhibitor was found to work synergistically when
administered with another inhibitor of fatty-acid biosynthesis, the antiseptic triclosan
(Miller et al., 2009). This work may inspire researchers to consider other bacterial ATP/GTP-utilizing enzymes as potential antibiotic targets and identify hit molecules similarly
(Walsh and Fischbach, 2009).
33
Based on three-dimensional shape and electrostatic complementarity, researchers
virtually screened 2.2 million compounds for inhibition of biotin carboxylase and
selected 525 for a high-through screening assay. A second experiment more carefully
probed the active site of the enzyme with 5,200 structurally diverse, small molecules via
saturation transfer difference NMR in addition to the acetyl-CoA carboxylase assay. This
assay revealed weak, yet ligand-efficient, binding fragments, and six molecules with an
IC50 less than 95 µM were found. X-ray crystallography provided visualization of binding
modes for the higher-affinity compounds and fragments, while overlays of several cocrystal structures exposed binding trends and more effective inhibitors were synthesized.
Iterative cycles of structure-based drug design involved fragment growing, merging and
morphing on successive fragment costructures. This process improved the potency of the
inhibitors up to 3000-fold, with the lead compounds being a series of amino-oxazoles
(Mochalkin et al., 2009).
Researchers from Schering-Plough used Affinity-Selection Mass Spectrometry to
identify a series of low-molecular-weight benzimidazoles that bind biotin carboxylase.
This series of molecules also inhibit the enzyme with an IC50 in the low µM range, and
the X-ray co-crystal structure with this inhibitor was solved and revealed important
interactions between the inhibitor and the biotin carboxylase ATP-binding site. Through
the use of Computer Aided Structure Based Drug Design and parallel synthetic
approaches, these workers were able to optimize inhibitor binding and generate a potent
and selective inhibitor of biotin carboxylase with antibacterial activity, having a 360-fold
improvement in enzyme inhibition (IC50 of 20 nM) compared with the initial compounds
(Cheng et al., 2009).
34
CHAPTER 1
INTRODUCTION
SECTION 2
Some Metabolic Enzymes Bind Nucleic Acids
Recently, it has been realized that metabolic enzymes, which are not traditionally
considered components of transcription or translation, can control gene expression
(Commichau and Stulke, 2008). This secondary function for these enzymes was coined
by Jeffery in 1999 as “moonlighting” and is thought of as a fossil from the RNA world
(Jeffery, 1999; Kyrpides and Ouzounis, 1995; Ciesla, 2006). One author suggested the
logic in this mechanism of regulation lies in the fact that proteins are the “most informed
molecules in the cell,” regarding concentration of metabolites, making these enzymes
ideal candidates for either controlling other gene regulators or regulating gene expression
directly (Commichau and Stulke, 2008). There are many examples of eukaryotic
metabolic moonlighting enzymes (Hall et al., 2004); however, there are far fewer in
prokaryotes. In 1995, Kyrpides and Ouzounis noted 16 examples from various organisms
(Kyrpides and Ouzounis, 1995). Since then, the list has grown, and a non-exhaustive
compilation, including 25 examples, is provided (Table 1-1), with more detail on a few
representative examples.
Though several moonlighting enzymes were listed, here a few well-understood
prokaryotic and eukaryotic examples from differing metabolic pathways are briefly
highlighted.
•
In prokaryotes:
35
o BirA is capable of synthesizing biotinyl-5′-AMP, transferring biotin to a lysine
residue of the biotin carboxyl carrier protein and repressing biotin biosynthesis
via site-specific DNA binding (Xu et al., 1995). The central domain of BirA is
catalytic, and the N-terminal domain binds DNA via a helix-turn-helix structural
module (Rodionov et al., 2002).
o The DNA-binding domain PutA can be expressed and purified without the
enzymatic domain and still bind DNA (Ostrovsky de Spicer and Maloy, 1993).
In the wild-type enzyme, the inactive form binds the promoter regions of related
genes and acts as a repressor; yet after a proline-dependent reduction of FAD
and a concomitant conformational change in the protein, the enzyme
translocates to the membrane for catalytic function (Zhang et al., 2004b).
o Iron regulatory protein 1 (also known as citrate dehydrogenase) catalyzes the
conversion of citrate to isocitrate in the tricarboxylic acid cycle and serves as a
lone example of a trigger enzyme with RNA-binding ability (Beinert and Kiley,
1996). In the presence of iron, this enzyme adopts a compact conformation; yet,
in the absence of iron, the enzyme assumes an open conformation capable of
binding mRNA.
•
In eukaryotes:
o The transcription factor from Drosophila, eyes absent, is known to have Tyr
phosphatase activity and the ability to autocatalytically self-dephosphorylate
and belongs to the haloacid dehalogenases family of enzymes (Tootle et al.,
2003).
36
o An enzyme known to bind DNA, a cysteine protease called LeCp, also has dual
functions. In the cytoplasm, this enzyme functions as a protease; however, upon
the binding of a small ubiquitin-related modifier protein, LeCp enters the
nucleus and acts as a transcription factor. Specifically, in tomato plants this
enzyme’s secondary function is to induce LeAcs2 (1-aminocyclopropane-1caboxylic acid synthase) expression.
o The mitochondrial enzyme Arg5,6 is involved in catalyzing two key steps in the
biosynthesis of ornithine, which is a precursor to arginine (Hall et al., 2004).
Ilv5p binds DNA and catalyzes a reaction in branched-chain amino-acid
biosynthesis (Bateman et al., 2002a).
o Interestingly, the binding of DNA and the enzymatic activity of Ilv5p are
independent of one another, as either the DNA-binding domain or the enzymatic
domain can be inactivated without affecting the activity of the other domain
(Bateman et al., 2002b).
o This thesis describes the dual functions of carboxyltransferase, one of which is
involved in binding nucleic acids, via a zinc finger. In the next section, the
catalytic and structural contribution of protein zinc fingers are considered as is
their interaction with nucleic acids.
Table 1-1. Examples of Nucleic-Acid Binding Metabolic Enzymes
Name
Organism
Function
Nucleic
Acid
Substrate
Reference
Human
Fermentation
DNA
Perucho, et
Eukaryotic DNA binding
LDH
37
al., 1980
hRoDH-E2
Human
Retinol
dehydrogenase
DNA
Markova et
al., 2006
Arg 5, 6
Yeast
Amino acid
synthesis
ds-DNA
Hall et al.,
2004
Eyes absent
Fly
Protein tyrosine
phosphatase
ds-DNA
Tootle et al.,
2003
HXK
Yeast
Glycolysis,
reaction 1
ds-DNA
Prior et al.,
1993
LeCp
Plants
Protease
ds-DNA
Matarasso et
al., 2005
NDP kinase
Human
NTP synthesis
ds-DNA
Postel et al.,
1993
Fatty Acid Synthase
Yeast
Fatty acid
synthesis
ss- and dsDNA
Kaslan and
Heyer, 1994
C1-THFS
Yeast
Amino acid
synthesis
ss-DNA
Wahls et al.,
1993
G3PD
Human
Glycolysis,
reaction 6
DNA,
(t)RNA
Perucho, et
al., 1980
Ilv5p
Yeast
Amino acid
synthesis
mt-DNA
Bateman et
al., 2002a
Aco1p
Yeast
Citric acid cycle mt-DNA
Chen et al.,
2005
1, 4-α-gBE
Rabbit
Glycogen
biosynthesis
RNA
Korneeva et
al., 1979
Catalase
Rat
H2 O2
breakdown
Cognate
mRNA
Clerch et al.,
1991
DHFR Human
Nucleotide
Cognate
Chu et al.,
Eukaryotic RNA binding
38
TS
Human
Thymine
synthesis
Cognate
mRNA
GDH
Cow
Amino acid
synthesis
Noncognate Preiss et al.,
RNA
1993
IDH
Yeast
Citric acid
cycle, reaction
3
Noncognate Elzinga et
RNA
al., 1993
Thiolase
Fly
Fatty acid
metabolism
Noncognate Nanbu et al.,
RNA
1993
Enolase
Yeast
Glycolysis
tRNA
Entelis et
al., 2006
BirA
Bacteria
Biotin synthesis
ds-DNA
Cronan,
1989
PutA
Bacteria
Amino acid
catabolism
ds-DNA
Ostrovsky
de Spicer
and Maloy,
1993
NadR
Bacteria
NAD
biosynthesis
ds-DNA
Raffaelli et
al., 1999
PepA
Bacteria
Aminopeptidase ds-DNA
Charlier et
al., 2000
Bacteria (and
humans)
Citric acid
cycle, reaction
2
Beinert and
Kiley, 1996
Chu et al.,
1993
Prokaryotic DNA binding
Prokaryotic RNA binding
Aconitase (IRP1)
39
IRE
(mRNA)
The Types and Functions of the Zinc Fingers
In 1869, while working with Aspergillus niger, Raulin first demonstrated zinc to
be essential for growth (Raulin, 1869). More recently, the metal has been shown
necessary for growth, development and differentiation (Vallee et al., 1986), and it is
present in all domains of life. Zinc is ubiquitous in all tissue types of higher organisms,
and the element typically exerts a catalytic, cocatalytic or structural function while bound
to proteins (McCall, et al., 2000). Though zinc atoms more often have a structural
function, there are more than 300 zinc-containing enzymes known, with examples
coming from the six classes of enzymes (Vallee and Auld, 1990).
The catalytic role of zinc has been described in the first known zinc
metalloenzyme, carbonic anhydrase II, discovered by Keilin and Mann in bovine blood
(Keilin and Mann, 1940). In most cases, the zinc ion in metalloenzymes contains a filled
d orbital and acts as a Lewis acid, accepting a pair of electrons (Williams, 1987). The
binding geometry most often observed is a distorted tetrahedron, and the zinc ion
coordinates three or four amino-acid residues (Grishin, 2001).
The structural role of the zinc atom is to bind amino-acid side chains, specifically
the nitrogen of histidine, the oxygen of aspartate or glutamate and the sulfur of cysteine
(Gregory et al., 1993), leading to the formation of discrete structural domains that are
self-stabilizing and that fold autonomously relative to the rest of the protein (Klug and
Schwabe, 1995). These structural domains are called motifs, as they have been observed
in different polypeptides, and they often confer a related function. The motif refers only
to the small region of the protein and is considered a supersecondary structure, as
opposed to a global conformation or tertiary structure.
40
The zinc finger, as a supersecondary structure, is capable of interacting with an
assortment of ligands that includes proteins, lipids (Grishin, 2001) and small molecules,
while functioning in a variety of biological processes, encompassing replication and
repair, transcription and translation, metabolism and signaling, cell proliferation and
apoptosis (Krishna et al., 2003). Over the next few paragraphs, zinc domains are
described with a focus on their main function of nucleic-acid binding; special attention is
paid to the structure most analogous to the zinc finger of carboxyltransferase.
Zinc fingers often are lumped with other DNA-binding motifs, i.e. helix-turnhelix and leucine-zipper, as having the capacity to recognize specific DNA sequences and
facilitate DNA binding by regulatory proteins. Zinc fingers were so named because the
individual modules “grip” the DNA (Klug and Schwabe, 1995) like fingers, while the rest
of the protein serves as the analogous “hand.”
The first zinc finger discovered was part of “factor A,” or TFIIIA, a transcription
factor from Xenopus laevis, and contained pairs of cysteines and histidines bound by a
single zinc atom. The TFIIIA zinc finger is a Cys2His2 zinc finger binding one zinc atom.
The residues that ligate zinc are spaced as follows: Cys-X(2 or 4)-Cys-X12-His-X3-His
(Miller et al., 1985). Since then, the discovery of other types of zinc fingers has followed,
but past methods of classification have focused on the types of residues that bind zinc or
the geometry of the interactions. The most recent method of classification focuses on the
protein backbone surrounding the zinc finger; specifically, Krishna and co-workers have
emphasized fold groups and described the conformation of the protein backbone within
the domain as a newer method for grouping zinc fingers (Krishna et al., 2003). A
41
reproduction of their table listing zinc finger classes and some zinc finger proteins is
provided (Table 1-2).
Table 1-2. List of Structural Classes of Zinc Fingers Highlighting Metal-Binding
Residues
42
It is important to note that the most common function of the zinc atom in the zinc
fingers is to provide regional secondary structure instead of directly binding nucleic
acids. Zinc fingers often interact with the major groove of DNA while wrapping around
both strands. Specific sequence recognition is determined via the side chains of several
amino acids, which are typically part of an α–helix adjacent to the zinc atom (Iuchi,
2005). Zinc-finger domains interact with DNA by forming hydrogen bonds and
hydrophobic interactions with nucleotide bases and often recognize specific triplets of
DNA sequences (Krishna et al., 2003). Moreover, most sequence-specific DNA-binding
proteins contain two or three tandem zinc fingers, if no other alternative DNA-binding
domains are present. In the case of the single zinc finger, low-affinity or nonspecific
binding is expected, yet the possibility remains for the necessity of an adaptor protein(s)
to facilitate sequence specific high-affinity DNA binding (Iuchi, 2005).
Krishna and co-authors provided copious information on the eight fold groups of
zinc-finger proteins; however, this dissertation will provide detail on the zinc ribbon fold
group that Bilder and co-workers suggest best resembles the zinc domain of
carboxyltransferase subunit of acetyl-CoA carboxylase from E. coli and S. aureus (Bilder
et al., 2006). Interestingly, Krishna and co-workers describe this group as containing the
most numerous examples of zinc fingers; however, aside from the metal-binding
residues, this group has minimal sequence conservation and is structurally variable.
The four residues binding zinc are paired, with each pair forming a zinc-knuckle,
and each zinc atom bound by two knuckles; however, the four ligands need not be
cysteine, and the metal atom may be iron. Each zinc-knuckle is made of β-hairpins,
usually two to four residues in length, and a variable third β-strand, if present, may
43
contribute hydrogen bonding with one of the other β-strands. This fold group has been
further subdivided into left- and right-handed zinc ribbons, based on the order of the zinc
ligands as positioned in a plane of the tetrahedron after orientation with the primary
hairpin above the secondary hairpin. Krishna and co-workers have subdivided the zinc
ribbon fold group; the subdivisions are called families and include:
1. Classical zinc ribbons with the following subfamilies: transcription factors,
primases, polymerases, topoisomerases and ribosomal proteins.
2. Rubredoxin-like fold, which binds zinc via a pair of cysteines and a pair of
histidines. In this family, each amino acid pair coordinates a separate zinc atom.
3. Single-stranded DNA-binding protein ADDBP.
4. Rubredoxin-like domains in enzymes that “function as interaction modules, e.g. to
provide a ‘lid’ for the enzyme's active site.” Examples include: aminoacyl tRNA
synthetase, silent information regulator (abbreviated SIR2) and adenylate kinase.
The zinc finger in carboxyltransferase is thought to be most structurally analogous
to the zinc fingers of proteins in the classic zinc ribbon family; thus, this subgroup is
further described. The classic zinc-ribbon family is defined by having a longer secondary
hairpin and longer three-stranded β-sheet when compared with other zinc-ribbon proteins.
Within the classic zinc-ribbon family, there are several subfamilies. One of the
subfamilies is called the transcription factors, and this group includes transcription
elongation factor SII (e.g. TFIIS and 1tfi), the transcription initiation factor TFIIB (e.g.
1pft and 1dl6) and the N-terminal 12 kDa fragment of DNA primase (e.g. 1d0q). Another
subfamily of the classic zinc-ribbon subgroup is the polymerase proteins subfamily. It
44
includes Rpb1, Rpb2, Rpb9 and Rpb12; however, the zinc domains of these proteins are
more variable in length. One more subfamily of the classic zinc ribbons consists of the
ribosomal proteins, such as the 50S ribosomal proteins L44E (chain 2 of 1jj2), L37E
(chain Z of 1jj2), and L37Ae (chain Y of 1jj2) from Haloarcula marismortui. Examples
of proteins within this subfamily are thought to be the closest structural match to
carboxyltransferase (Krishna et al., 2003; Bilder et al., 2006).
The Tetracyclines, Additional Inhibitors of Carboxyltransferase
Chapter 3 covers tetracycline inhibiting carboxyltransferase, and background
information on tetracycline, including a brief history, follows. Though we consider
antibiotics to be less than a century old, mankind has used these natural products for
much longer. The presence of tetracyclines, as discovered by ultraviolet microscopy in
1,400-year-old bone found in what is present day Sudan, suggests that man has long since
coexisted with microbes and benefitted medicinally from the family of secondary
metabolites collectively known as tetracyclines (Armelagos et al., 2001). In 1947,
Benjamin Minge Duggar first isolated tetracycline from Streptomyces aureofaciens,
naming it Aureomycin (Duggar, 1948). Around the same time, Pfizer described a
molecule with similar melting point, solubility, pH range stability, and minimum
concentrations needed to inhibit growth for various bacterial species, while patenting the
fermentation and production process (Finlay and Hobby, 1950). These compounds
quickly obtained wonder-drug status, and, aside from their gram-positive and gramnegative antibiotic effect, derivatives have had therapeutic indications to include the
treatment of infectious diseases such as Chlamydiae, mycoplasmas, and rickettsiae, as
45
well as noninfectious diseases such as bone resorption, cartilage degradation, diabetes,
periodontal disease, arthritis, and cancer (Chopra and Roberts, 2001; Greenwald and
Golub, 2001).
The tetracycline family of therapeutics is founded on the ABCD naphthacene
ring-based chemical structure (Figure 1-9) and positional locants (locants are used to
specify the position of a functional group within a molecule) designated at scaffold
positions C1 (carbon number 1) through 12a (the carbon that is part of ring A and is also
bound to C12). Nitrogen- and oxygen-containing functional groups are attached to C10,
C11, C12, C1, as well as the 2N region and C3-C4 regions. Most medicinal derivatives of
tetracycline have a pattern of hydroxyl, keto-enol, and carbonyl groups, attached at the
lower peripheral region, owing to the molecule’s potency and selectivity. The functional
groups across the lower peripheral region are critical for Mg2+-binding, and the bacterial
ribosome contains a Mg2+ that guides the molecule into its binding position. Thus, some
chemical changes to the lower peripheral region result in loss of antibacterial activity.
The functional groups bound at the upper peripheral region (carbons 5 – 9) also dictate
the derivative’s pharmacological characteristics. Chemical derivatization surrounding the
D-ring aromatic region resulted in second- and third-generation tetracycline-based
antibiotics. The exocyclic carbonyl attached to carbon 2 and the keto-enolate group
bound to carbon 3 also mediates antimicrobial activity, and broad-spectrum effectiveness
is based on the orientation of the C4 dimethylamino group.
In E. coli, the mechanism of action for tetracyclines involves binding the
ribosomal 30S subunit (Last, 1969; Chopra and Roberts, 2001) and excluding the
46
incoming aminoacyl-tRNA from the acceptor (A-site) site (Semenkov et al., 1982).
However, tetracyclines also are known to chelate metal ions, as tetracyclines complexed
with proteins containing divalent metal atoms are capable of inhibiting enzymatic activity
of zinc- (or calcium-) containing enzymes, i.e., matrix metalloproteinase. Evidence has
been marshaled supporting the theory that tetracyclines inhibit enzymes by binding: (1)
catalytic and structural zinc atoms (Greenwald et al., 1998), (2) enzyme-associated zinc
atoms in positions other than active sites (Smith et al., 1999), and (3) soluble zinc atoms
before binding active sites as tetracycline — metal ion complexes (Takahashi et al.,
1991).
Figure 1-9. The Tetracycline ABCD Ring System and Designation of the Upper and
Lower Peripheral Regions. (Image courtesy of Mark Nelson)
Objectives and Rationale for Study
Most structural and mechanistic work on prokaryotic acetyl-CoA carboxylase has
focused on biotin carboxylase. After the crystal structure of carboxyltransferase was
47
solved, a zinc finger, or putative nucleic-acid binding domain, was revealed (Bilder et.
Al., 2006). Thus, our goal was to understand the function of this supersecondary
structure.
Our preliminary objective was to determine the nucleic-acid binding capacity of
the enzyme. Next we wondered, “What is the ideal nucleic-acid substrate, and how is this
related to regulation?” After we concluded that the enzyme bound DNA nonspecifically,
high-affinity and specific binding to RNA was confirmed. Subsequently, we supposed the
presence of biotin and acetyl-CoA, the reaction substrates, would have some effect on
nucleic-acid binding. The next obvious question became, “Do nucleic acids inhibit
carboxyltransferase-mediated catalysis?” After the inhibitory patterns were produced for
nucleic acids and heparin, we studied the inhibitory effects of a class of molecules that
was only recently found to inhibit the enzyme, namely the tetracyclines. At that point, we
wanted to determine (a) the inhibition patterns and (b) if tetracyclines were sufficient to
prevent nucleic-acid binding and then we wanted to propose a hypothesis as to the
biological relevance of this inhibition in the context of polyketide synthesis. Lastly, to
better understand inhibitor binding sites and modes of action we wondered how each of
these inhibitors might function in the presence of a second inhibitor.
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Yao X, Wei D, Soden C Jr, Summers MF, and Beckett D. (1997). Structure of the
carboxy-terminal fragment of the apo-biotin carboxyl carrier subunit of Escherichia coli
acetyl-CoA carboxylase. Biochem 36, 15089-15100.
Yu LP, Kim YS, Tong L. (2010). Mechanism for the inhibition of the carboxyltransferase
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22072-22077
63
Zagnitko O, Jelenska J, Tevzadze G, Haselkorn R, Gornicki P. (2001). An
isoleucine/leucine residue in the carboxyltransferase domain of acetyl-CoA carboxylase
is critical for interaction with aryloxyphenoxypropionate and cyclohexanedione
inhibitors. Proc Natl Acad Sci U S A 98, 6617-6622.
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diclofop. Proc Natl Acad Sci USA 101, 5910-5915.
Zhang H, Yang Z, Shen Y, and Tong L. (2003). Crystal structure of the
carboxyltransferase domain of acetyl-coenzyme a carboxylase. Science 299, 2064-2067.
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64
CHAPTER 2
LINKAGE BETWEEN NUCLEIC ACID BINDING AND CATALYSIS IN THE
CARBOXYLTRANSFERASE SUBUNIT OF ACETYL-COA CARBOXYLASE*
Introduction
Carboxyltransferase is one component of the multifunctional biotin-dependent
enzyme acetyl-CoA carboxylase, and the bacterial homolog of this enzyme contains a
domain with no counterpart in the eukaryotic enzyme. Acetyl-CoA carboxylase catalyzes
the first committed and regulated step in fatty acid biosynthesis in bacteria via a two-step
reaction (Scheme 1).
(1) Enzyme-biotin + MgATP + HCO3-
Mg2+
(2) Enzyme-biotin-CO2- + Acetyl CoA
Enzyme-biotin-CO2- + MgADP + Pi
Malonyl CoA + Enzyme-biotin
Scheme 1
In contrast to eukaryotes, which encode the multifunctional acetyl-CoA
carboxylase as a single polypeptide chain, bacterial acetyl-CoA carboxylase is composed
of three separate proteins: biotin carboxylase, carboxyltransferase, and biotin carboxyl
carrier protein (BCCP) (Cronan and Waldrop, 2002). Biotin is covalently attached to
BCCP and this complex is designated Enzyme-biotin in Scheme 1. Biotin carboxylase
catalyzes the first half-reaction, which is an ATP-dependent carboxylation of biotin to
form carboxybiotin. The second half-reaction is catalyzed by carboxyltransferase, which
transfers the carboxyl group from carboxybiotin to acetyl-CoA to generate malonyl-CoA.
The recently obtained crystal structures (Figure 2-1) of carboxyltransferase from
Escherichia coli and Staphylococcus aureus revealed a unique domain absent from
* Reprinted by permission of The Journal of Protein Science
65
eukaryotic homologs (Bilder et al., 2006). The structure confirmed the α2β2 subunit
composition suggested by Lane and colleagues (Guchhait et al., 1974) and showed that
the enzyme belongs to the crotonase superfamily (Gerlt and Babbitt, 2001). The overall
fold, not surprisingly, is similar to that of carboxyltransferase from yeast (Zhang et al.,
2003) and Streptomyces coelicolor (Diacovich et al., 2004). However, when the gene for
the β subunit of E. coli carboxyltransferase was cloned and sequenced 20 years ago, the
authors noted the tandem C-X-X-C sequences separated by 15 residues located at the
amino terminus and hypothesized that the protein may bind a metal ion (Bognar et al.,
1987). The crystal structures of carboxyltransferase from S. aureus and E. coli, along
with X-ray fluorescence studies, have confirmed this prediction. The metal atom is zinc,
and it forms part of a zinc motif that is unique to bacterial carboxyltransferase (Figure 21). A space-filling representation reveals that the Zn domain forms part of a saddle-like
structure (Figure 2-2), and an electrostatic surface potential rendering shows that the
inner face of the zinc finger domain has an electropositive surface potential, while most
of the protein has an electronegative surface potential.
Given that Zn domains are commonly associated with nucleic acid binding, and
noting the favorable electrostatic potential, we explored the ability of E. coli
carboxyltransferase to bind nucleic acids. Herein nucleic acid binding properties of E.
coli carboxyltransferase are characterize along with the effect of DNA binding on
enzymatic activity. Notably, DNA exhibits synergism with a bisubstrate analog inhibitor
and with heparin, suggesting communication between the dual active sites of the
functional protomers.
66
Figure 2-1. Ribbon Drawing of Carboxyltransferase from S. aureus. The α-chain is in
purple and the β-chain is in gold. The zinc atom is depicted as a blue sphere in the βchain.
Figure 2-2. Electrostatic Surface Potentials of the Heterotetramer of Carboxyltransferase.
Blue represents a net positive charge, while red represents a net negative charge. The
surface potentials were generated with the program Grasp (Nicholls et al., 1991).
67
The potential for the binding of DNA or structural analogs of DNA to affect
enzymatic activity of carboxyltransferase is particularly significant because there is
renewed interest in targeting enzymes in the bacterial fatty acid biosynthetic pathway for
antibiotic drug discovery (Campbell and Cronan, 2001; Zhang et al., 2006). Since the
zinc finger domain is only found in the bacterial form of the enzyme, it provides a unique
target to achieve specificity for pharmaceutical intervention.
Materials and Methods
Restriction enzymes, dNTPs, and T4 DNA ligase were purchased from New
England Biolabs. Pfu Turbo DNA polymerase was from Stratagene. Primers for PCR
and oligonucleotides used to generate dsDNA and ssDNA used as inhibitors were
obtained from MWG BioTech. Heparin was from Sigma/Aldrich. The bisubstrate
analog (BiSA) inhibitor of carboxyltransferase was synthesized according to the method
of Levert and Waldrop (17) [γ-32P] ATP and [α-32P] UTP were from PerkinElmer, and
MegaScript/ MegaClear/ RNase Zap was acquired from Ambion, Inc. QIAquick Gel
Extraction Kit and DNeasy Tissue Kit were from Qiagen, Inc. IHF was obtained as
described by Grove et al. (Grove et al., 1996), MfpA was kindly provided by Dr. John
Blanchard, Albert Einstein College of Medicine.
Purification and Enzymatic Assay of Carboxyltransferase
Carboxyltransferase was purified from E. coli transformed with an overexpression
plasmid containing a mini-operon with genes for the α and β subunits of the enzyme
(Blanchard and Waldrop, 1998). Carboxyltransferase activity was measured in the
reverse direction with a spectrophotometric assay in which the production of acetyl-CoA
68
was coupled to the combined citrate synthase- malate dehydrogenase reaction requiring
NAD+ reduction. Biocytin is preferred over biotin because biocytin produces a maximal
velocity three orders of magnitude greater than does biotin.45 Biocytin is a biotin
molecule with a lysine appended to the carboxyl group of the valeric acid side chain via
an amide linkage at the ε-amino group (Blanchard and Waldrop, 1998). NADH
formation was followed spectrophotometrically at 340 nm using a Uvikon 810 (Kontron
Instruments) spectrophotometer interfaced to a PC equipped with a non-commercial data
acquisition program. Since the crystal structure showed that carboxyltransferase contains
two active sites, the initial velocities were calculated per active site using a molecular
weight of 68.5 kDa for each αβ dimer (i.e., active site). Control experiments showed that
neither DNA nor heparin at the highest concentration inhibited the coupling enzymes
citrate synthase or malate dehydrogenase.
Substrate Nucleic Acids
The promoter regions for the α (292 bp and 500 bp fragments) and β (175 bp and
300 bp fragments) genes of carboxyltransferase, the promoter region for the biotin
carboxylase/ biotin carboxyl carrier protein (300 bp fragment) and a 280 bp control
fragment encompassing part of the accD coding region were PCR amplified using the
primers listed in Table 2-1. E. coli (JM109, New England Biolabs) chromosomal DNA
isolated via DNeasy Tissue kit was used as the template. The PCR products were
electrophoresed and their concentration was assessed by densitometry by comparing
sample intensity to that of similar sized molecular weight standards. The PCR products
69
were 5′-end labeled with T4 polynucleotide kinase and [γ-32P] ATP and purified via
QIAquick Gel Extraction Kit, assuming 90% sample recovery.
The mRNA encoded by the genes for α (accA) and β (accD) subunits of
carboxyltransferase, including their respective leader sequences, was obtained as follows.
The accA gene (1190 bp) was PCR amplified from the E. coli genome using primers
listed in Table 2-1. Primers introduced EcoRI and BamHI sites at either end of the PCR
product. For the β subunit (accD, 1027 bp) an XhoI site was added to the 5′ end and a
BamHI site was incorporated at the 3′ end. These PCR products were cloned into pGEM11Zf(+). The presence of both genes was confirmed via sequential digestions, agarose
gel electrophoresis, and observation of the correct insert size. These recombinant
plasmids were linearized with BamHI and 390 ng of DNA was used as templates for in
vitro transcription of accA and accD. Two identical reactions were carried out except for
the addition of 1µL of [α-32P] UTP to one reaction. In vitro transcription and mRNA
purification were carried out according to the instructions in MegaScript and MegaClear,
respectively. A 1.0% agarose gel was used to visualize the nonradioactive mRNA to
confirm the correct size of the transcript. The nonradioactive RNA was quantified by
measuring its absorbance at 260 nm. Measured parameters for the nonradioactive
samples were applied to the radioactive samples.
Eletrophoretic Mobility Shift Assay (EMSA)
The reactions (total reaction volume of 10 µL) included 50 fmol of DNA or RNA,
with carboxyltransferase titrated from 0 to 2.6 µM for assays involving DNA and 0 to 0.6
µM for assays involving RNA. The binding buffer was: 20 mM Tris-HCl (pH 8), 0.1
70
mM Na2EDTA, 0.075% BRIJ58, 50 mM NaCl, 5 mM MgCl2, 50 µg/ml BSA, and 4%
(v/v) glycerol. Reactions were incubated on ice for 0.5 h. Samples were resolved on
prerun 6.0% (w/v) native polyacrylamide gels (39:1 acrylamide:bisacrylamide) at room
temperature in TBE buffer (45 mM Tris borate (pH 8.3), 1.25 mM Na2EDTA), and
samples were loaded with the power on. After 2.0 h of electrophoresis, the gels were
dried and complexes were visualized by phosphorimaging and quantified using
ImageQuant TL (GE Healthcare). For calculating percent complex formation, the region
on the gel from the slowest migrating complex to the free nucleic acid was considered as
complex. Percent complex was calculated as complex intensity divided by the total
intensity (or total nucleic acid) in a given lane. Binding assays were carried out in
duplicate. Binding isotherms are from representative EMSAs, with less than 17.5%
variation between experiments.
Inhibitors of DNA and RNA binding by carboxyltransferase were analyzed by measuring
percent complex versus inhibitor concentration. A competition assay was carried out
similar to the binding assay for affinity determination (above), except the
carboxyltransferase was held constant and the inhibitor concentration was varied. The
order of addition for each of the regents in the competition assay is as follows: buffer,
carboxyltransferase, inhibitor molecule(s), and labeled DNA.
Data Analysis
The binding isotherms for carboxyltransferase binding to DNA were analyzed by fitting
the data to equation 1 to calculate the concentration of protein resulting in half-maximal
saturation (EC50).
71
Table 2-1. Primers Used for Amplification of Substrate DNA or for Enzyme Inhibition
Assays
Primer Amplicon locus Upstream Primer
Downstream Primer
Name
Sequence (5′ to 3′)
Sequence (5′ to 3′) or
complement for annealing
accD #1 -161 to +13
AAATAAAAAGTAACT
TCAATCCAGCTCATTAGGG
CCGCGGTTCG
ACCTTT
accD #2 -288 to +11
TTTCTTCGGTACAATC AATCCAGCTCATTAGGGAC
CCGATGGT
CTTTC
accA #1 -6,729 to -6,437 CAAAGTCTGGCGCAA
CTTCAAAATCCAGCACGCG
AACCGCTGC
ATCCA
accA #2 -495 to +4
GTTGCCGCGCGGGTTA CTCATAGTATTCCTGTATT
TGGTCACC
AGTCA
accB/C
-292 to +7
control
-28 to +251
ivt
accA
ivt
accD
4 nt
30 nt
-213 to +960
-96 to +914
N/A
N/A
TTGCTACGAAATCGTT
ATAATGTG
GCCTGGCATTTGCTGA
ATTTGACGA
GCCGGGAATTCTGACC
AGCTTTTAAACCG
GCCGGCTCGAGTGTGC
AACATTCATGGTCT
TACG
TGACCATGATTACGCC
AAGCTATTTAGGTG
ATATCCATGAGTGGGTTCC
GTACT
TCGTCAAATTCAGCAAATG
CCAGGC
CCGGCGGATCCTTACGCGT
AACCGTAGCTC
CCGGCGGATCCTCAGGCCT
CAGGTTCCTGA
N/A
CACCTAAATAGCTTGGCGT
AATCATGGTCA
Amplicon locus is given relative to the A of the start codon (ATG). Primers accD
#1 and #2 were used to amplify a 175 bp and a 300 bp region of the E. coli chromosome
predicted to contain the promoter for accD (carboxyltransferase β gene). accA #1 and #2
were used to amplify a 292 bp and a 500 bp region of the E. coli chromosome predicted
to contain the promoter for accA (carboxyltransferase α gene). accB/C were used to
amplify a 300 bp region of the E. coli chromosome reported to contain the promoter for
the accB and accC (genes for biotin carboxylase and biotin carboxyl carrier protein,
respectively). “Control” was used to amplify a 280 bp region starting at the 5′ end of the
coding sequence within accD. ivt accA was used to amplify accA from the -213 position
(Li and Cronan, 1993) to the end of the gene to generate template for in vitro
transcription. ivt accD was used to amplify accD from the -96 position (Li and Cronan,
1993) to the end of the gene and was used to generate in vitro transcription template. The
4 nt and both 30 nt were used to study the inhibitory affects of DNA on catalysis by
carboxyltransferase.
72
In equation 1, Y equals the fractional complex, while min is the minimum value
of Y and max is the maximum value of Y. “x” is the concentration of carboxyltransferase
and n is the Hill coefficient. The binding isotherms for carboxyltransferase binding to
RNA were analyzed by fitting the data to equation 2 where Kd reflects half maximized
saturation, Y is the fractional complex, X is concentration of carboxyltransferase, and
Ymax is the maximal complex or horizontal asymptote.
Competitive inhibition data were fitted to equation 3, using the programs of
Cleland (Cleland, 1979). In equation 3, v is the initial velocity, Vm is the maximal
velocity, A is the substrate concentration, I is the concentration of inhibitor, Km is the
Michaelis constant, Kis is the slope inhibition constant. Data for multiple inhibition were
fitted to equation 4 where v is the initial velocity, I and J are the concentrations of the two
inhibitors, v0 is the velocity in the absence of inhibitors, Ki and Kj are the apparent
dissociation constants for the two inhibitors and β is a measure of the degree of
interaction of the two inhibitors (Cleland, 1990).
!"!!!"#
Y = min + !!!" !"#$%&'!! ∗!
Y=
v=
v=
[1]
!!"# !
[2]
!! ! !
!! ∗!
!! !!
!
!!"
[3]
!!
!!
!
!
!"
!! ! !
!! !! !!! !!
[4]
Results
DNA Binding by Carboxyltransferase
73
To address the functional significance of the unique Zn domain, we first pursued
its potential role in DNA binding. Since the four components (i.e. biotin carboxylase,
BCCP and the α and β subunits of carboxyltransferase) of the E. coli acetyl-CoA
carboxylase are likely produced in a defined, stoichiometric ratio, and since expression of
the genes has been reported to be directly correlated to cellular growth rates (Li and
Cronan, 1993), we explored the possibility that carboxyltransferase may contribute to
regulation of any of the genes encoding the acetyl-CoA carboxylase subunits.
The gene encoding the Zn domain-containing β subunit (accD) is located between
the dedA (Nonet et al., 1987) and the folC (Bognar et al., 1987) genes. Li and Cronan (Li
and Cronan, 1993) suggested that accD is transcribed monocistronically and that the
accD mRNA starts 87 bp upstream of the coding region. The ability of
carboxyltransferase to bind a 175 bp DNA containing the 5′ noncoding region of accD
was tested using electrophoretic mobility shift assay (EMSA). As shown in Figure 2-3A,
carboxyltransferase binds DNA as evidenced by the formation of a complex that fails to
migrate from the well of the gel (designated as complex 1). The complex (designated
complex 2) that migrates much faster than complex 1 may represent a very compact
species, perhaps resulting from a single DNA molecule wrapping around the enzyme
molecule, which contains two zinc domains located on opposite lobes. Quantification of
the data shows that DNA binds to carboxyltransferase in a cooperative fashion (Figure 23B), with nH = 1.7 and a half-maximal saturation of 946 ± 32 nM (Table 2-2). Halfmaximal saturation of 300 bp DNA that included the 3′ end of the dedA gene was similar
to that measured for the 175 bp fragment (Table 2-2).
74
Table 2-2. Half-Maximal Values for Carboxyltransferase Binding to Substrate DNA
Molecules
DNA
EC50 (nM)
Hill Slope
175 bp carboxyltransferase β promoter
946 (+/- 32)
1.7 (+/- 0.2)
300 bp carboxyltransferase β promoter
673 (+/- 36)
1.8 (+/- 0.2)
292 bp carboxyltransferase α promoter
751 (+/- 25)
2.1 (+/- 0.2)
500 bp carboxyltransferase α promoter
821 (+/- 28)
2.0 (+/- 0.2)
300 bp BCCP/ BC promoter
794 (+/- 47)
1.5 (+/- 0.2)
280 bp control sequence
783 (+/- 37)
1.5 (+/- 0.2)
30 nt PCR primer
480 (+/- 18)
2.6 (+/- 0.2)
EC50 values and Hill slope for DNA substrates were determined by quantifying EMSA gels
and fitting data to equation 1.
Figure 2-3. Electrophoretic Analysis of 175 bp dsDNA from Promoter Region of accD
(carboxyltransferase β subunit) titrated with carboxyltransferase. A, reactions contain
50.0 fmol DNA. Reactions in lanes 2 - 14 contain: 0.1, 0.2, 0.4, 0.6, 0.8, 1.0, 1.2, 1.4,
1.6, 1.8, 2.0, 2.2, 2.4, and 2.6 µM carboxyltransferase. B, binding isotherm for
carboxyltransferase binding to 175 bp dsDNA.
75
(Figure 2-3 continued)
The gene coding for the α subunit (accA) of carboxyltransferase is located
downstream of the polC gene (Li and Cronan, 1992), and Li and Cronan (Li and Cronan,
1993) suggested that the promoter is located within the 3′ end of the polC coding
sequence. Carboxyltransferase bound to 292 and 500 bp DNA fragments encompassing
the promoter region of accA (data not shown). As with the promoter region for the β
subunit, binding was cooperative and the half-maximal saturation was between 0.5 and
1.0 µM (Table 2-2). Similarly, low-affinity binding of carboxyltransferase to a 280 bp
DNA representing the accD coding region and to a 300 bp DNA containing the promoter
region for the operon that includes the genes for biotin carboxylase and the biotin
carboxyl carrier protein was observed (Table 2-2), and the binding was cooperative.
76
The comparable half-maximal saturation of the DNA sequences examined
suggests that carboxyltransferase binds DNA nonspecifically. To confirm nonspecific
DNA binding, a competition assay was performed. The addition of unlabeled 175 bp β
subunit promoter DNA fails to inhibit complex formation with the labeled DNA, even at
a 10-fold molar excess of unlabeled competitor (Figure 2-4). The increase in complex
formation in the presence of higher amounts of unlabeled 175 bp β subunit promoter
DNA is likely attributable to a carboxyltransferase concentration below the Kd for single
site binding. A titration of the cloning vector pGEM5 (up to a 10-fold molar excess)
reveals inhibition of the carboxyltransferase DNA complex, further confirming
nonspecific binding (Figure 2-4). Inhibition was independent of DNA conformation
(supercoiled or linear, data not shown).
So far, the data indicate that carboxyltransferase binds duplex DNA
nonspecifically and with low affinity. While it cannot rule out that carboxyltransferase
may be recruited to specific DNA sites by association with other factors, it is noted that
addition of biotin carboxylase or the biotin carboxyl carrier protein (biotinylated and
unbiotinylated) did not increase the affinity of carboxyltransferase for duplex or singlestranded DNA, suggesting that the other components of acetyl-CoA carboxylase do not
confer high affinity DNA binding (not shown). Considering the potential for
carboxyltransferase looping DNA and the presence of AT-rich sequences weakly
resembling consensus binding sites for integration host factor (IHF) (Freundlich et al.,
1992; Hales et al., 1994; and Hales et al., 1996), we assessed the ability of IHF to
enhance carboxyltransferase-DNA complex formation. IHF did not increase the affinity
77
of carboxyltransferase for either of the promoter regions of the genes for the α and β
subunits (data not shown).
Figure 2-4. DNA Competition Assay. Reactions contain 50 fmol labeled 175 bp
carboxyltransferase β promoter, reactions in lanes 2 - 15 have 600.0 nM
carboxyltransferase. Unlabeled 175 bp carboxyltransferase β promoter DNA (lanes 3 - 8)
and pGEM5 (lanes 10 – 15) are titrated using molar ratios of unlabeled DNA/ labeled
DNA from 0.2 to 10 (3.4 – 171 molar ratio of bp for pGEM5).
Carboxyltransferase binds ssDNA in preference to duplex (Figure 2-5, A).
Although the EC50 for ssDNA was not significantly lower than the EC50 for the 30 nt
dsDNA (Figure 2-5, B, Table 2-2), the ssDNA has orders of magnitude fewer binding
sites in comparison to other dsDNA tested, suggesting a lower Kd for each single site. As
seen for binding to duplex DNA, carboxyltransferase binds ssDNA cooperatively. In
light of this result DNA mimetics were tested for their ability to bind to
carboxyltransferase. Not surprisingly, the DNA mimic heparin did inhibit formation of
the carboxyltransferase-DNA complex (data not shown). In contrast, MfpA, a protein
from Mycobacterium tuberculosis that provides resistance to fluoroquinolones by
mimicking DNA and binding to DNA gyrase (Hedge et al., 2005) did not inhibit
78
formation of the carboxyltransferase-DNA complex (not shown). Taken together, the
results indicate that carboxyltransferase binds DNA nonspecifically and with low affinity.
Figure 2-5. Electrophoretic Analysis of 30 nt ssDNA Versus 30 bp dsDNA Titrated with
Carboxyltransferase. A, Lane 1 is ssDNA (50.0 fmol). The remaining reactions
contained a 50 fmol ssDNA with one-half molar ratio complementary strand, annealed by
cooling from 95°C to 0°C over 3 hours. Reactions in lanes 3 - 14 contain increasing
amounts of carboxyltransferase (from 0.1 to 2.4 µM). B, binding isotherm for
carboxyltransferase binding to ssDNA (30 nt upstream primer sequence, Table 2-1).
79
Figure 2-6. A, Competition Assay with RNA. All reactions have 50.0 fmol 280 bp
labeled DNA (control, Table 2-1) and 1.6 µM carboxyltransferase, except the reaction in
lane 1 which has only free DNA. The transcript from the gene for the α-subunit of
carboxyltransferase is titrated in reactions in lanes 2 - 7. The transcript from the gene for
the β-subunit of carboxyltransferase is titrated between lanes 8 - 13. The reaction in lane
14 is labeled DNA and carboxyltransferase only. Total RNA is titrated over the reactions
in lanes 15 - 20. tRNA from Saccharomyces cerevisiae is titrated over the reactions in
lanes 21 - 26. The titration for each type of RNA had mass ratios (DNA/ RNA) of 0.05,
0.5, 1.0, 5.0, 50.0, and 500.0. B, graphical representation of RNA competition EMSA.
80
Figure 2-7. Electrophoretic Analysis of the Transcript from Carboxyltransferase α
Subunit Titrated with Carboxyltransferase. A, All lanes contain 50.0 fmol mRNA, while
lanes 2 - 14 contain 1.0, 5.0, 10.0, 25.0, 40.0, 55.0, 70.0, 85.0, 100.0, 200.0, 300.0, 400.0,
500.0, and 600.0 nM carboxyltransferase. B, binding isotherm for carboxyltransferase
binding to the transcript from carboxyltransferase α subunit.
RNA Binding by Carboxyltransferase
Considering that the closest structural homolog of the carboxyltransferase Zn
domain is from the 50S ribosomal protein L37Ae (Bilder et al., 2006), the ability of
81
carboxyltransferase to bind RNA was investigated. Four different types of RNA were
tested for their ability to inhibit formation of the DNA:carboxyltransferase complex. The
RNA molecules considered included mRNA coding for the α subunit, mRNA coding for
the β subunit, a total RNA extract from E. coli, and a tRNA extract from Saccharomyces
cerevisiae. All four types of RNA were found to inhibit formation of the
carboxytransferase-DNA complex (Figure 2-6A) indicating that carboxyltransferase also
binds RNA. Quantification of the data showed that the α subunit mRNA inhibited
complex formation most effectively (Figure 2-6B). Therefore, a transcript including the
α subunit mRNA starting 213 nt upstream of the AUG (Li and Cronan, 1993) through the
end of the structural gene was used in an EMSA (Figures 2-7A and B). In contrast to the
sigmoidal binding isotherm obtained for carboxyltransferase binding to DNA, fits of
carboxyltransferase-RNA complex formation yield a hyperbolic curve with Kd, of 187 ±
17 nM, which is about 3-5 fold lower than the half-maximal saturation for DNA binding.
Inhibition of Enzyme Activity
While the EMSA studies indicated that carboxyltransferase binds nucleic acids
nonspecifically and with low affinity, the important question was whether DNA binding
had any impact on the enzymatic function of carboxyltransferase. As shown in Figure
2-8, increasing concentrations of a 4 nt, 30 nt and 30 bp fragment of DNA did indeed
result in a decrease in enzymatic activity. It was not possible to test larger DNA
fragments because the increased viscosity of the assay solution became prohibitive. A
single stranded DNA substrate (30 nt upstream sequence, Table 2-1) was used to examine
the type of inhibition with respect to the substrates malonyl-CoA and biocytin†. The 30
82
nt ssDNA exhibited competitive inhibition with respect to both malonyl-CoA and
biocytin (Figure 2-9A and B). Fitting the data to equation 3 gave slope inhibition
constants (Kis) of 85.1 ± 10.3 µM, with respect to malonyl-CoA and 34.2 ± 4.0 µM, with
respect to biocytin. Heparin was also found to inhibit enzymatic activity, and like DNA,
exhibited competitive inhibition with respect to both substrates (Figure 2-9C and D). The
slope inhibition constants (Kis) for inhibition by heparin are 1.2 ± 0.1 µM with respect to
malonyl-CoA, and 2.2 ± 0.3 µM with respect to biocytin.
ss4-mer
ds30-mer
ss30-mer
Fractional Activity
1.0
0.5
0.0
10 0
10 2
10 4
10 6
10 8
10 10
[DNA] (pg)
Figure 2-8. Dose Response Curve for Carboxyltransferase with Both ssDNA and
dsDNA. Initial velocity was measured at increasing amounts of DNA (4 nt ssDNA, 30 nt
ssDNA, and 30 nt dsDNA, Table 2-1).
83
Figure 2-9. Inhibition of Carboxyltransferase by DNA (A and B) and Heparin (C and D).
When malonyl-CoA was the variable substrate, biocytin was held constant at 5.0 mM,
and when biocytin was the variable substrate, malonyl-CoA was held constant at 0.1 mM.
The points are the reciprocal of the experimental velocities and the lines are derived from
the best fit of the data to equation 3.
84
(Figure 2.9 continued)
85
catalyzes a reaction in branched-chain amino acid biosynthey either act as enzymes or as nucleic acid binding
thesis (Bateman et al. 2002a); and iron regulatory protein 1
proteins, and, most importantly, these two functions can be
(IRP1), which binds mRNA and functions as aconitase
separated. For example, the DNA-binding domain of PutA
(Walden et al. 2006). These proteins all have dual functions;
they either act as enzymes or as nucleic acid binding
proteins, and, most importantly, these two functions can be
separated. For example, the DNA-binding domain of PutA
A.
B.
Complex
DNA
C.
Figure 5. Inhibition of the formation of DNA:carboxyltransferase complex by malonyl-CoA and BiSA. All reactions contained 50 fmol of 300-bp
carboxyltransferase b promoter DNA. (A) Reactions in lanes 2–10 have
800.0 nM carboxyltransferase; malonyl-CoA is titrated in reactions from
lanes 3–10 (from 10.0 mM to 10.0 mM). (B) Reactions in lanes 2–10 have
800.0 nM
BISA is of
titrated
in reactions in lanes 3–10
Figure
5. carboxyltransferase;
Inhibition of the formation
DNA:carboxyltransferase
com(from
2.5
mM
to
2.5
mM).
plex by malonyl-CoA and BiSA. All reactions contained 50 fmol of 300-bp
Figure
Inhibition
Formation
DNA:
Carboxyltransferase
Complex by
carboxyltransferase
b 2-10.
promoter
DNA. of
(A)theReactions
inoflanes
2–10
have
Substrates and Substrate Analog. All reactions contained 50 fmol 300 bp
800.0 nM carboxyltransferase; malonyl-CoA is titrated in reactions from
carboxyltransferase β promoter DNA. A, reactions in lanes 2 - 10 have 800.0 nM
lanes 3–10 (from 10.0 mM to 10.0 mM). (B) Reactions in lanes 2–10 have
carboxyltransferase, malonyl-CoA is titrated in reactions from lanes 3 - 10 (from 10.0 µM
800.0 nM carboxyltransferase;
is titrated
in 2reactions
lanes
3–10
to 10.0 mM). B,BISA
reactions
in lanes
- 8 have in
800.0
nM
carboxyltransferase, biocytin is
(from 2.5 mM to
2.5 mM).
titrated
in reactions in lanes 3 - 8 (from 800.0 µM to 800.0 mM). C, reactions in lanes 2 -
10 have 800.0 nM carboxyltransferase, BISA is titrated in reactions in lanes 3 - 10 (from
2.5 µM to 2.5 mM).
86
These inhibition data are consistent with EMSA assays, which show that
increasing concentrations of either malonyl-CoA or biocytin inhibit formation of the
carboxyltransferase-DNA complex (Figure 2-10A and B). Moreover, a bisubstrate
analog (BiSA) inhibitor of carboxyltransferase, in which coenzyme A is covalently
attached to carboxybiotin (Levert and Waldrop, 2002) was an even more efficient
inhibitor of the enzyme-DNA complex (Figure 2-10C).
The competitive inhibition patterns observed for DNA and heparin versus the
substrates malonyl-CoA and biocytin suggest that DNA and heparin bind in the active
site of carboxyltransferase. However, this seemed curious given the significant
difference in structure between the substrates and the fact that DNA and heparin are
polymers. Therefore, to characterize the topological relationship between the DNA
binding site and the active site (e.g., do the two sites overlap) multiple inhibition studies
were performed. Multiple inhibition experiments are carried out by measuring the initial
velocity at increasing concentrations of one inhibitor while the second inhibitor is held
constant. The substrate concentrations are held constant at subsaturating levels. The
initial velocities are measured again at higher levels of the second inhibitor and then
plotted as 1/velocity versus the concentration of the first inhibitor (sometimes referred to
as a Yonetani-Theorell plot) (Yonetani and Theorell, 1964). The first inhibitor for these
studies was the bisubstrate analog (BiSA) used above for the EMSA assays. It is
important to note that the substrate concentrations are held constant at subsaturating
levels for multiple inhibition analyses, which enables BiSA to bind to the enzyme.
Double inhibition of carboxyltransferase by either DNA or heparin at different
fixed levels of BiSA (Figure 2-11A and B) yielded intersecting patterns, which indicate
87
that the two inhibitors can bind to the enzyme simultaneously. Thus, the binding site for
DNA and the active site of the enzyme are topologically distinct, which suggests DNA
binding at one αβ dimer while BiSA binds to the other αβ dimer. This scenario is
supported by fitting the data to equation 4; β values of 0.50 and 0.85 were found for the
DNA-BiSA inhibition and the heparin-BiSA inhibition, respectively. The β value is an
indication of the interaction of the two inhibitors. Values of β greater than 1 indicate that
the binding of the two inhibitors interfere with one another, while a value of 1 indicates
no interaction between the inhibitors. A β value less than 1 indicates synergism in the
binding of the inhibitors. Thus, the binding of either DNA or heparin shows very weak
synergism with the binding of BiSA and vice versa. Surprisingly, double inhibition
analysis with DNA and heparin also resulted in an intersecting pattern (Figure 2-11C),
indicating that DNA and heparin can bind to the enzyme simultaneously. However, the β
value of 0.02 reflects a strong synergistic relationship in the binding of the two inhibitors.
Given the presence of two Zn domains per protomer, the simplest explanation is that
binding at one site enhances binding at the other.
Discussion
DNA Binding Enzymes
Few enzymes that catalyze a reaction in intermediary metabolism also bind DNA
or RNA. Examples include PutA (proline utilization A) (Brown and Wood, 1992); BirA,
or biotin ligase, which attaches biotin to the biotin carboxyl carrier protein (Beckett,
2005); the plant cysteine protease LeCp (Matarasso et al., 2005); Arg 5,6, which is
involved in arginine biosynthesis (Hall et al., 2004); Ilv5p, which catalyzes a reaction in
88
Figure 2-11. (A-C) Multiple Inhibition Patterns for DNA (ssDNA 30 nt upstream
sequence, Table 2-1), Heparin and BiSA. The points are the reciprocal of the
experimental velocities and the lines are derived from the best fit of the data to equation
4. Malonyl-CoA was held constant at 0.1 mM, while biocytin was held constant at 5.0
mM.
89
(Figure 2-11 continued)
90
branched-chain amino acid biosynthesis (Bateman et al., 2002a); and iron regulatory
protein 1 (IRP1), which binds mRNA and functions as aconitase (Walden et al., 2006).
These proteins all have dual functions; they either act as enzymes or as nucleic acid
binding proteins and, most importantly, these two functions can be separated. For
example, the DNA binding domain of PutA can be expressed and purified without the
enzymatic domain, and it still binds DNA (Gu et al., 2004). For Ilv5p, either the DNA
binding domain or the enzymatic domain can be inactivated without affecting the activity
of the other domain (Bateman et al., 2002b). In contrast, the enzymatic activity and
nucleic acid binding of carboxyltransferase are inextricably linked.
Zinc Domains Associated with Proteins Involved in DNA Metabolism
The zinc domain in bacterial carboxyltransferase belongs to the zinc ribbon class
of zinc fingers (Krishna et al., 2003). Other proteins that contain this type of zinc finger
include the transcription factors TFIIS (Qian et al., 1993) TFIIB (Zhu et al., 1996) TFIIE
(Okuda et al., 2004) several subunits from RNA polymerase II (Cramer et al., 2003)
human ssDNA binding protein RPA (Cochkareva et al., 2002) bacteriophage T4 and T7
primases (Cha and Alberts, 1986; Mendelman and Richardson, 1991) and several
ribosomal proteins, most notably, L37E from the 50S subunit (PDB code 1JJ2).
Carboxyltransferase was found to bind DNA non-specifically and with low
affinity. Compared to other proteins featuring the zinc ribbon motif, non-specific DNA
binding by carboxyltransferase is not unexpected. Isolated zinc fingers like that in
carboxyltransferase do not bind DNA tightly and recognize only 3 nucleotides (Wolfe et
al., 2000). For example, T7 and T4 primases, which contain a zinc ribbon type of zinc
91
finger, recognize a 3 nt sequence (Mendelman et al., 1999). Recognition of 3 nt is
consistent with the observation that enzymatic activity was inhibited as efficiently by a 4
nt DNA as well as longer DNA molecules. That carboxyltransferase also binds ssDNA
and RNA is consistent with properties of other zinc ribbon proteins. Clearly, zinc ribbons
are associated with proteins involved in DNA metabolism, even if some, such as the
domain found in TFIIB mediates interactions with the large RNA polymerase subunit and
not nucleic acid, however, this zinc ribbon is almost entirely β-sheet and has less in
common with other characterized zinc ribbons. While we have not found evidence for a
role in regulation of gene expression, we cannot rule out the possibility that
carboxyltransferase may be recruited to specific sites. Could the non-specific DNA
binding by carboxyltransferase be an evolutionary relic? We consider this interpretation
unlikely, given the close mechanistic relationship between DNA binding and catalysis.
Does Carboxyltransferase Bind DNA in Vivo?
A prediction as to whether carboxyltransferase binds DNA in vivo can be made by
assuming a 3 nt binding site which is consistent with other zinc finger binding sites
(Wolfe et al., 2000) and with inhibition of enzymatic activity by 4 nt DNA. However,
based on the dimensions of carboxyltransferase a single molecule of enzyme would
occlude approximately 30 bp. Given that the concentration of the E. coli genome is 5 nM
(Sundararaj et al., 2004) and with 4.6 x 106 bp in the E. coli genome (Mathews et al.,
1999) there would be 1.5 x 105 or 0.75 mM potential binding sites for
carboxyltransferase. The intracellular concentration of carboxyltransferase is
approximately 500 nM (Guchhait et al., 1974). Using 0.8 µM as the binding constant of
92
DNA to carboxyltransferase, the fraction of carboxyltransferase bound to DNA
intracellularly can be calculated with the following equation (Segel, 1975):
½
[ES] = (1/2) (([Et] + [St] + Ks) - (([Et] + [St] + Ks)2 - 4[Et] [St]) )
where Et is the concentration of carboxyltransferase, St is the concentration of DNA
binding sites and Ks is the dissociation constant for DNA binding to carboxyltransferase.
Assuming that the entire E. coli genome is accessible to carboxyltransferase, the
calculation suggests that all of the enzyme would be bound to DNA in vivo. However,
most of the genomic DNA in E. coli is compacted by proteins (Minsky, 2004) and would
be inaccessible to carboxyltransferase, yet recalculation assuming only 10% of the
genomic DNA available for binding to carboxyltransferase also suggests that all of the
carboxyltransferase in E. coli would be bound to DNA. The levels of RNA in E. coli
were not included in this calculation, but if they were, it would only reinforce the
conclusion that all of the carboxyltransferase is bound to either DNA or RNA in the cell.
What might be the consequences of a significant fraction of carboxyltransferase
being bound to nucleic acids? The observation that both substrates inhibit DNA binding
is very important when considering how carboxyltransferase could function as a critical
enzyme for membrane biogenesis. It is tempting to speculate that during the stationary
phase of E. coli growth where nutrients are limited, carboxyltransferase functions as a
nucleoid-associated protein (Drlica and Rouviere, 1987) to compact and protect the
chromosome from damage. During the growth phase when nutrients are abundant, the
levels of substrates, most notably acetyl-CoA, increase dramatically and compete with
DNA for binding to carboxyltansferase and the enzyme functions to synthesize fatty acids
93
for cell membrane assembly. During stationary phase, when carboxyltransferase activity
is low, association with nucleic acids may also prevent futile activity and/or the
sequestration of cellular metabolites.
The Mode of DNA Binding Suggests Communication Between the Dual Active Sites
DNA and heparin are competitive inhibitors with respect to both substrates
malonyl-CoA and biocytin. The competitive inhibition patterns indicate that saturating
levels of either substrate prevent DNA binding. This makes intuitive sense for malonylCoA; the β subunit contains the zinc domain, the presumed DNA binding site, in close
proximity to the malonyl-CoA binding site. Biocytin, however, binds to the α subunit.
The competitive inhibition pattern could be due to a steric effect where the polymeric
DNA blocks access to the biocytin binding site. Alternatively, biocytin binding to the α
subunit could propagate a conformational change to the β subunit to inhibit DNA
binding. Inhibition of carboxyltransferase by DNA is consistent with EMSA data
showing that both malonyl-CoA and biocytin as well as a bisubstrate analog prevent
formation of the DNA-protein complex. Thus, enzymatic activity and DNA binding are
not separable functions.
The cooperative binding of DNA to carboxyltransferase is likely a possible
manifestation of two separate physical phenomena. Carboxyltransferase has two zinc
domains diametrically opposed and DNA has multiple binding sites. Accretion of
carboxyltransferase mediated by protein-protein interactions could account for the
observed cooperativity. Assuming protein-protein interaction as the driving force,
however, it is difficult to imagine why such recruitment would not occur on RNA, to
94
which carboxyltransferase does not bind cooperatively. Second, the binding of DNA to
one site on carboxyltransferase could increase the affinity for DNA binding to the second
site. Evidence for intersubunit communication is also seen in the multiple inhibition
experiments with DNA and heparin (Figure 2-11) where the β value, which reports on the
degree of synergism in the binding of the two inhibitors, is well below 1 (0.02). This
synergistic binding of DNA to carboxyltransferase could certainly contribute to the
sigmoidal curve observed in Figure 2-3B. While DNA binding to carboxyltransferase is
clearly cooperative, the binding of RNA is not. It is not clear what the reason is for this
difference. Since most RNA molecules change conformation upon protein binding
(Leulliot and Varani, 2001; Williamson, 2000), perhaps the RNA bound to
carboxyltransferase assumes a conformation that does not induce cooperative binding.
Thus, DNA binding may cause an induced fit in carboxyltransferase while RNA binding
results in an induced fit of the RNA.
Pharmaceutical Relevance
The presence of the zinc finger domain and the finding that DNA binding inhibits
enzymatic activity is very important for developing pharmaceutical agents specific for
bacterial carboxyltransferase. Acetyl-CoA carboxylase and, therefore,
carboxyltransferase is found in humans as well as in bacteria. Thus, antibiotics that target
acetyl-CoA carboxylase will have to achieve species specificity in order to minimize
toxicity. Since the eukaryotic form of carboxyltransferase does not have a zinc domain,
inhibitors binding to the zinc domain of the bacterial enzyme are likely to be species
specific. The findings in this report demonstrate that molecules that bind to the DNA
95
binding site will likely inhibit enzymatic activity since DNA binding and catalysis are
reciprocally coupled.
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99
CHAPTER 3
INHIBITION OF THE CARBOXYLTRANSFERASE SUBUNIT OF ACETYL-COA
CARBOXYLASE FROM ESCHERICHIA COLI AND STAPHYLOCOCCUS
AUREUS BY TETRACYCLINES
Introduction
The first member of the tetracycline group of antibiotics was discovered in the
late 1940s when chlortetracycline (Figure 3-1 and Table 3-1) was found to be produced
by Streptomyces aureofaciens (Duggar, 1948). The isolation of oxytetracycline and
demeclocycline (Figure 3-1 and Table 3-1) soon followed along with tetracycline (Figure
3-1 and Table 3-1) which was also produced by Streptomyces aureofaciens (Nelson,
2001).
Tetracycline could also be synthesized by catalytic hydrogenation of
chlortetracycline (Nelson, 2001). Tetracyclines exhibit a broad spectrum of activity
against both gram-positive and gram-negative bacteria as well as chlamydiae,
mycoplasmas, rickettsiae and protozoan parasites (Chopra and Roberts, 2001).
Tetracyclines mode of action involves inhibiting protein synthesis by binding to the 30S
subunit of the ribosome and preventing the binding of the incoming acylated tRNA
(Chopra and Roberts, 2001). In 1973, workers (Vanek et. al., 1973) described
oxytetracycline mediated inhibition of acetyl-CoA carboxylase (and pyruvate
dehydrogenase) in S. aureofaciens. Here we characterize the inhibition mechanism on the
carboxyltransferase subunit of acetyl-CoA carboxylase from gram-positive and gramnegative bacteria.
Acetyl-CoA carboxylase (ACC) catalyzes the first committed and regulated step
in fatty acid biosynthesis in bacteria via a two-step reaction (Scheme 1).
100
R1
R2
R3
R4
N(CH 3) 2
H
H
OH
CONH 2
OH
OH
O
OH
O
Figure 3-1. Stick Rendering of a Basic Tetracycline Molecule. R groups at positions 1 –
4 for several derivatives of tetracycline are detailed in Table 3-1.
Table 3-1. Functional Groups for Tetracycline and Derivatives. Derivative names and
functional groups at positions R1 – R4 are shown. Species of origin for each derivative is
listed; semisynthetic means the molecule is a result of synthetic modification of a
naturally occurring precursor molecule.
Derivative
R1
R2
R3
Tetracycline
H
CH3 OH H
Streptomyces rimosus/ aureofaciens
Chlortetracycline
Cl
CH3 OH H
Streptomyces aureofaciens
Demeclocycline
Cl
OH
Streptomyces aureofaciens
Oxytetracycline
H
CH3 OH OH Streptomyces rimosus
Minocycline
N(CH3)2 H
Doxycycline
H
CH3 H
OH semisynthetic
Anhydrotetracycline H
CH3 C
H
H
R4
H
H
H
Mg2+
(1) Enzyme-biotin + MgATP + HCO3(2) Enzyme-biotin-CO2- + Acetyl CoA
Origin
semisynthetic
Streptomyces rimosus/ aureofaciens
Enzyme-biotin-CO2- + MgADP + Pi
Malonyl CoA + Enzyme-biotin
Scheme 1
101
In gram-positive and gram-negative bacteria, ACC is composed of three separate
proteins: biotin carboxylase, carboxyltransferase, and biotin carboxyl carrier protein
(BCCP) (Cronan and Waldrop, 2002). Biotin is covalently attached to BCCP and is
designated Enzyme-biotin in Scheme 1. Biotin carboxylase catalyzes the first-half
reaction, which is an ATP-dependent carboxylation of biotin, to form carboxybiotin. The
second half-reaction is catalyzed by carboxyltransferase, which transfers the carboxyl
group from carboxybiotin to acetyl-CoA to generate malonyl-CoA.
Given the dramatic increase in antibiotic resistance of pathogenic bacteria there is
a renewed interest in targeting fatty acid biosynthesis enzymes for antibiotic drug
discovery (Campbell and Cronan, 2001, Zhang et al., 2006). Since the metabolic role of
acetyl-CoA carboxylase in bacteria is membrane biogenesis it is a prime target for
antibiotics. Inhibition of the carboxyltransferase subunit of acetyl-CoA carboxylase by
tetracyclines not only provides new insight into the antibacterial mechanism but can also
serve as a starting point for the design of new inhibitors of carboxyltransferase.
Materials and Methods
All the tetracycline derivatives were purchased from Sigma/Aldrich except
anhydrotetracycline which was purchased from Cole-Parmer. Heparin, coupling
enzymes, NAD and DMSO were also from Sigma/Aldrich. The 30-nt single-stranded
sequence (5´-TGACCATGATTACGCCAAGCTATTTAGGTG-3´) used in the multiple
inhibition studies, was obtained from MWG BioTech. The bisubstrate analog (BiSA)
was synthesized according to Levert and Waldrop (Levert and Waldrop, 2002).
Purification and Enzymatic Assay of Carboxyltransferase
102
Carboxyltransferase from E. coli was purified from a strain of E. coli transformed
with an overexpression plasmid containing a mini-operon with the genes for the α and β
subunits of the enzyme (Blanchard and Waldrop, 1998). Carboxyltransferase from S.
aureus was purified as described by Bilder et al. (Bilder et al., 2006).
Carboxyltransferase activity was measured in the reverse direction1 with a
spectrophotometric assay in which the production of acetyl-CoA was coupled to the
combined citrate synthase-malate dehydrogenase reaction requiring NAD+ reduction
(Blanchard and Waldrop, 1998). NADH formation was followed spectrophotometrically
at 340 nm using a Uvikon 810 (Kontron Instruments) spectrophotometer interfaced to a
PC equipped with a data acquisition program. Since the crystal structure showed
carboxyltransferase contained two active sites, the initial velocities were calculated per
active site using a molecular weight of 68.5 kDa for each αβ dimer (i.e. active site).
Measuring the activity of carboxyltransferase in the presence of tetracyclines with
the above assay presented two problems that limited the concentration of tetracyclines
that could be tested. First, tetracyclines are sparingly soluble in water; therefore, all
tetracyclines were dissolved in DMSO (dimethyl sulfoxide). Thus, all velocities were
measured with solutions containing 10% DMSO. It is important to point out that 10%
DMSO decreased the catalytic rate 32% , which does not agree with previously published
reports (Santoro et al., 2006). Second, NADH and tetracycline have an overlap in their
absorbance spectra such that high concentrations of tetracycline masked the formation of
NADH. Therefore, a reference cuvette, containing all reagents in the sample cuvette
except for carboxyltransferase, was used. Both of these physical constraints placed an
upper limit on the concentrations of tetracyclines that could be tested. Consistant with
103
previous reports (Vanek et. al., 1973), tetracyclines were not observed to inhibit the
activity of either coupling enzyme. This was confirmed by measuring the reduction of
NAD+ after adding a subsaturating concentration of acetyl-CoA rather than
carboxyltransferase to a reaction mixture in the presence and absence of the tetracycline.
Electrophoretic Mobility Shift Assays
EMSAs were performed as described by Benson et al., (Benson et al., 2008).
Briefly, the binding assays (total reaction volume of 10 µL) included 50 fmol of 300bp
DNA, from the promoter region of the β gene of carboxyltransferase (Benson et al.,
2008), while carboxyltransferase was held at 0.8 µM. The binding buffer was: 20 mM
Tris-HCl (pH 8), 0.1 mM Na2EDTA, 0.075% BRIJ58, 50 mM NaCl, 5 mM MgCl2, 50
µg/ml BSA, and 4% (v/v) glycerol. Protein-nucleic acid complexes were equilibrated on
ice for 0.5 h in the presence of oxytetracycline. Samples were resolved on prerun 6.0%
(w/v) native polyacrylamide gels (39:1 acrylamide:bisacrylamide) at room temperature in
TBE buffer (45 mM Tris borate (pH 8.3), 1.25 mM Na2EDTA), and samples were loaded
with the power on. After 2.0 h of electrophoresis, the gel was dried and complexes were
visualized by phosphorimaging.
Data Analysis
Competitive inhibition data were fitted to equation 1, using the programs of
Cleland (Cleland, 1979). In equation 1, v is the initial velocity, Vm is the maximal
velocity, A is the substrate concentration, I is the concentration of inhibitor, Km is the
Michaelis constant, and Ki is the inhibition constant. Data for multiple inhibition were
fitted to equation 2 where v is the initial velocity, I and J are the concentrations of the two
104
inhibitors, v0 is the velocity in the absence of inhibitors, Ki and Kj are the apparent
dissociation constants for the two inhibitors, and β is a measure of the degree of
interaction of the two inhibitors (Cleland 1990).
v = Vm · A / Km (1 + I/Ki) + A
[1]
v = v0 / (1 + I/Ki + J/Kj + IJ/βKiKj)
[2]
Results and Discussion
Tetracyclines Inhibit Carboxyltransferase Activity
Seven commercially available tetracyclines were tested for their ability to inhibit
carboxyltransferase activity. A titration of increasing amounts of each of the
tetracyclines resulted in a corresponding decrease in the initial velocity of
carboxyltransferase (Figure 3-2). It was not possible to calculate an IC50 value for any of
the derivatives in Figure 3-2 because the plateau region at high concentrations of the
tetracyclines could not be determined due to overlap in absorbance spectra between
NADH and the tetracyclines. However, the estimated IC50 values for each of the
derivatives ranges from about 100 µM to 800 µM. Each of the tetracyclines inhibited
carboxyltransferase from both E. coli and S. aureus. Moreover, while it has been
proposed that a magnesium-tetracycline complex is the species that binds to ribosomes
(Chopra and Roberts, 2001) we found that magnesium did not increase the inhibitory
effects of tetracycline (data not shown). Tetracycline was used as a representative
member of this class of antibiotics to determine the type of inhibition with respect to the
substrates malonyl-CoA and biocytin. Tetracycline exhibited competitive inhibition with
respect to both malonyl-CoA and biocytin for E. coli (Figure 3-3 A and B) and S. aureus
105
(Figure 3-3 C and D) carboxyltransferase. Fitting the data to equation 1 gave inhibition
constants (Ki) of 194 ± 22 µM, with respect to malonyl-CoA, and 199 ± 15 µM, with
respect to biocytin, for E. coli carboxyltransferase, while in the case of S. aureus
carboxyltransferase, the Ki values for malonyl-CoA and biocytin were 110 ± 9 µM and
245 ± 22 µM, respectively.
Figure 3-2. A: Concentration Response Curve for Tetracyclines and E. coli
Carboxyltransferase. B: Concentration response curve for tetracyclines and S. aureus
carboxyltransferase.
106
Figure 3-3. Inhibition Patterns for Tetracycline Versus both Malonyl-CoA and Biocytin,
respectively, with E. coli (A and B) and S. aureus (C and D) Carboxyltransferase. When
malonyl-CoA was the variable substrate, biocytin was held constant at 5.0 mM, and when
biocytin was the variable substrate, malonyl-CoA was held constant at 0.1 mM. The
points are the reciprocal of the experimental velocities and the lines are derived from the
best fit of the data to equation 1.
107
(Figure 3-3 continued)
The competitive inhibition patterns indicate that the substrates and tetracycline
compete for binding to the free enzyme. Usually this means the substrates and inhibitor
bind to the same site. However, it can also mean that the inhibitor and substrate do not
compete for the same binding site but that infinite levels of substrate simply prevent
inhibitor binding. Thus, to determine if tetracycline and substrate binding are mutually
exclusive (or conversely can tetracycline and the substrates bind to the enzyme
108
simultaneously) multiple inhibition analyses were carried out using a bisubstrate analog
(BiSA) inhibitor of carboxyltransferase, in which coenzyme A is covalently attached to
carboxybiotin as an active site ligand (Levert and Waldrop, 2002).
Multiple inhibition experiments are carried out by measuring the initial velocity at
increasing concentrations of one inhibitor while the second inhibitor is held constant.
The initial velocities are measured again at higher levels of the second inhibitor and then
plotted as 1/velocity versus the concentration of the first inhibitor (sometimes referred to
as a Yonetani-Theorell plot) (Yonetani and Theorell, 1964). It is important to keep in
mind that unlike the inhibition studies in Figure 3-2 where the substrate concentrations
are extrapolated to infinity, the substrate concentrations in multiple inhibition
experiments are held constant at subsaturating levels which allows both inhibitors to bind
to the enzyme. Double inhibition of carboxyltransferase by BiSA and different fixed
levels of tetracycline yielded intersecting patterns (Figure 3-4A), which indicate that the
two inhibitors can bind to the enzyme simultaneously and suggests the binding site for
tetracycline and the active site of the enzyme are topologically distinct. Fitting the data
to equation 2 yielded a value of 0.3 for the β parameter which is an indication of the
interaction of the two inhibitors. Values of β greater than 1 indicate that the binding of
the two inhibitors interfere with one another, while a value of 1 indicates no interaction
between the inhibitors. A β value less than 1 indicates synergism in the binding of the
inhibitors. Thus, the binding of tetracycline shows very weak synergism with the binding
of BiSA and vice versa.
109
Figure 3-4. Double Inhibition Plots of Tetracycline Versus: DNA (A), Heparin (B), and
BiSA (C) for E. coli Carboxyltransferase. The points are the reciprocal of the
experimental velocities and the lines are derived from the best fit of the data to equation
2. Malonyl-CoA was held constant at 0.1 mM, while biocytin was held constant at 5.0
mM.
110
(Figure 3-4 continued)
Effect of Tetracycline on DNA Binding
The crystal structures of carboxyltransferase from E. coli carboxyltransferase and
S. aureus revealed that the β subunit contains a zinc finger domain on the amino terminus
(Bilder et al., 2006). Zinc fingers are commonly found in proteins that bind DNA and
carboxyltransferase was recently found to bind DNA non-specifically (Benson et al.,
2008). The DNA analog heparin also bound to carboxyltransferase and both DNA and
heparin inhibited enzymatic activity (Benson et al., 2008). Thus, given that the zinc
domain is not found in the Streptomyces genera (Diacovich et al., 2004) which synthesize
tetracyclines perhaps tetracycline inhibited carboxyltransferase by binding to the DNA
binding site. To investigate whether DNA and tetracycline share the same binding site on
carboxyltransferase we utilized an electrophoretic mobility shift assay (EMSA) and
multiple inhibition analyses. As shown in the EMSA in Figure 3-5, increasing amounts
111
of oxytetracycline did not inhibit formation of the carboxyltransferase-DNA complex
suggesting that tetracycline and DNA do not compete for the same binding site. These
results are consistent with multiple inhibition analyses with DNA and heparin. Both
DNA and heparin exhibited intersecting patterns with tetracycline (Figure 3-4 B and C)
indicating that both tetracycline and DNA or heparin can bind to the enzyme
simultaneously. Moreover, the β values (1.08 and 0.95 for DNA-tetracycline and
heparin-tetracycline, respectively) indicate there is no interaction between DNA or
heparin and tetracycline.
Figure 3-5. Oxytetracycline Effects on the DNA Binding Ability of Carboxyltransferase.
Reactions in lanes 1 - 7 have 50.0 fmol of 300bp radio-labeled DNA carboxyltransferase
β promoter DNA (Benson 2007), reactions in lanes 2 - 7 have 800.0 nM
carboxyltransferase, oxytetracycline is titrated in the reactions from lanes 3 - 7 (from 35.0
µM to 700.0 µM).
While the multiple inhibition and EMSA studies indicate that tetracycline does
not bind to either the active site or the DNA binding site it is not clear where tetracycline
binds to carboxyltransferase. What is known is that the binding of tetracycline to
carboxyltransferase is prevented at saturating levels of either substrate, therefore, any
112
tetracycline binding site must be affected by substrate binding. A possible binding site
for tetracycline may be the interface between the αβ dimers (Figure 3-6). Binding at the
αβ dimer interface would allow access to the active site and the zinc domain and
saturation with either substrate could induce a conformational change that prevents
tetracycline binding. Tetracycline binding would also disrupt communication between
the two αβ dimers that comprise the active sites. It has already been shown that the two
active sites communicate at least with respect to DNA binding (Benson et al., 2008).
Figure 3-6. Ribbon Drawing of Carboxyltransferase from S. aureus. The α-chain is in
purple and the β-chain is in gold. The zinc atom is depicted as a blue sphere in the βchain.
Does Tetracycline Bind Carboxyltransferase in Vivo?
For many years, authors have stressed the similarities between tetracycline or
polyketide biosynthesis and biosynthesis of fatty acids. Similarities respective of both
biosynthesis mechanisms and primary sequences of critical enzymes in both pathways
have caused investigators to postulate that the enzymes evolved from a common origin
113
after early gene duplication (Revill et al., 1996); a brief overview of the most pertinent
reactions in both pathways follows.
In Streptomyces, fatty acid synthase catalyzes the formation of fatty acids (Revill
et al., 1995). Ketoacyl carrier protein synthase (FabH in initiation and FabB in
elongation) catalyzes a decarboxylative reaction with acyl-ACP substrate and malonylACP. The resulting 3-ketoacyl-ACP product is then modified via reduction, dehydration,
and enoyl reduction, leading to an extended acyl-ACP product. This product serves as
the substrate for subsequent elongation by FabB. The malonyl-ACP extender unit, acting
as a substrate for FabB and FabH, is generated from malonyl-CoA by the action of
malonyl-CoA:ACP transacylase (FabD).
Similarly in polyketide biosynthesis, malonyl-CoA is used. Specifically, nine
molecules of malonyl-CoA and a series of oxytetracycline biosynthesis gene cluster
products catalyze the formation of oxytetracycline, one of the earliest derivatives of
tetracycline to be produced. This reaction requires a minimal polyketide synthase,
namely: ketosynthase, chain length factor, and an acyl-carrier protein, named OxyA,
OxyB, OxyC (also called TcmM), respectively; and a malonyl-CoA:ACP transacylase,
FabD (Florova et al., 2002; Hertweck et al., 2007; Zhang et al., 2006), borrowed from
fatty acid synthesis. In an iterative process, these enzymes, along with malonyl-CoA,
generate the polyketide backbone of tetracycline.
Tang and co-workers showed that OxyD, an amidotransferase homolog,
incorporates an amide unit into the polyketide backbone. The structure generated
spontaneously cyclizes, via C11 – C16 regioselectivity, to produce a molecule Tang
termed WJ85 (Pickens and Tang, 2009, Figure 3-7), which at this point resembles both
114
tetracycline and a polymerized moiety of malonyl-CoA. Subsequent enzymatic reactions
produce anhydrotetracycline, followed by oxytetracycline, and finally tetracycline.
Figure 3-7. Stick Rendering of an Early Minor Product, Resembling Malonyl-CoA in the
Tetracycline Synthesis Pathway (Pickens and Tang, 2009).
Florova and colleagues noted that in Streptomyces, the acyl carrier proteins
carrying malonyl-CoA, FabC (in fatty acid synthesis) and OxyC (in polyketide
biosynthesis), are interchangeable. However, it appears that only FabD covalently links
malonyl-CoA to the carrier proteins FabC and OxyC. Having similar affinities for both,
FabD thus provides another tangible link between the two pathways (Revill et al., 1996).
However, Florova also noted that the KCAT for the enzyme in fatty acid synthesis is nearly
four times that of the comparable enzyme in polyketide biosynthesis reaction. Thus, it is
possible that, under normal conditions, malonyl-CoA is primarily utilized in fatty acid
synthesis, but under conditions of nutrient stress, i.e., phosphate deprivation (Votruba and
Běhal,1984), malonyl-CoA is diverted to polyketide biosynthesis and tetracycline is
115
made. Potentially, tetracycline acts here to tip the scale towards polyketide biosynthesis
and away from fatty acid synthesis.
fatty acid biosynthesis
bacterial growth
polyketide (TET) biosynthesis
territory establishment
abundant nutrients
malonyl-CoA
limited nutrients
Figure 3-8. Two Possible Fates of Malonyl-CoA in Streptomyces
Literature suggests that during the stationary phase when fewer fatty acids are
needed, secondary metabolites increase, thus tetracycline biosynthesis increases.
Interestingly, cultures of Streptomyces incubated for long periods show an increase in the
production of tetracycline just prior to autolysis. The addition of tetracycline to cultures
during exponential growth phase abolishes tetracycline biosynthesis, while only partially
reducing protein synthesis (Vanek et al., 1973). Moreover, Vanek and coworkers
observed inhibition of ACC by chlorotetracycline in Streptomyces; therefore tetracycline
potentially contributes to the regulation of both pathways at lower concentrations, prior to
bacteriostatic effects at higher concentrations.
Interestingly, and perhaps counterintuitively, in vitro transcription assays using
ribosomes isolated from tetracycline-producing organisms revealed increased sensitivity
in organisms producing the antibiotic when compared to those incapable of its
116
production. This is likely due to the absence of protection/resistance mechanisms present
in vivo (Mikulik et al.,1983).
As the work discussed in this manuscript was conducted on carboxyltransferase
from E. coli and S. aureus, tetracycline’s effects on carboxyltransferase from
Streptomyces remain unknown. If tetracycline were also to weakly inhibit
carboxyltransferase from Streptomyces, one may wonder if this were part of a survival
mechanism employed by the organism. Conceivably, if tetracycline strongly inhibited
carboxyltransferase, fatty acid synthesis would decrease too quickly upon the onset of
tetracycline biosynthesis. The potential for tetracycline to regulate fatty acid synthesis
and polyketide biosynthesis in Streptomyces remains, however testing this theory should
prove difficult without understanding the in vivo protection mechanisms involved in fatty
acid synthesis, analogous to the in vitro transcription assays discussed.
It is worth noting that, unlike in Streptomyces, the only known fate of malonylCoA in E. coli is the synthesis of fatty acids (James and Cronan, 2004). Perhaps in E.
coli tetracycline does not serve as a switch to direct malonyl-CoA between fatty acid
synthesis and polyketide biosynthesis; rather, it only inhibits fatty acid synthesis, aside
from inhibiting translation.
Structural analysis and comparison of the ribosome, carboxyltransferase, acetylCoA, and tetracycline may be telling. Analysis of the ribosome-tetracycline complex
reveals extensive hydrogen bonding between tetracycline hydroxyls and the phosphate
backbone of RNA at the A-site, in addition to hydrophobic interactions between the
tetracycline rings and the RNA bases (Pioletti et al., 2001). The ribosomal A-site and
acetyl-CoA binding site on carboxyltransferase are both solvent exposed clefts.
117
Preliminary superpositions of tetracycline interacting with the substrate-occupied acetylCoA binding site of carboxyltransferase suggest a possible fit whereby, acetyl-CoA
presents phosphate oxygen atoms that interact with tetracycline in much the same way
that RNA-presented phosphate oxygen atoms in the ribosomal A-site of the ribosometetracycline complex.
This observation is supported by our data, suggesting that both tetracycline and
DNA (or presumably RNA) can bind to the enzyme simultaneously while not occupying
the same site, and tetracycline and the substrates can bind to the enzyme simultaneously.
Another possibility is that the tetracycline disturbs the placement of the aliphatic tail of
acetyl-CoA. Although this disruption could allow acetyl-CoA binding via the nucleotide
binding portion of the molecule, the active part of acetyl-CoA (namely, the pantetheine
moiety sitting in a hydrophobic cleft) at the interface between the alpha and beta subunits
of carboxyltransferase, could be filled by tetracycline.
Additionally, just as there are hydrophobic interactions between tetracycline and
the RNA bases of the bacterial ribosome, there may be equivalent interactions between
the tetracycline and the hydrophobic residues in the active site of carboxyltransferase.
Lastly, the reduced affinity of tetracycline for carboxyltransferase compared to the
ribosome may be supported by the loss of hydrophobic interactions and fewer, lessspecific hydrogen bonding interactions. In the context of these potential mechanisms of
inhibition, it is important to point out that the mechanism of action for tetracyclines has
been questioned, and the actual mechanism of action is still unknown (Schnappinger and
Hillen, 1996).
118
Pharmaceutical Relevance
The fact that tetracyclines inhibit carboxyltransferase is not only helpful for
understanding the mechanism of action of this class of antibiotics but it also provides a
starting point for developing more potent inhibitors with pharmacological properties.
There is precedent for carboxyltransferase as a target for antibiotics. For instance, the
antibiotic moiramide B was found to inhibit carboxyltransferase (Freiberg et al., 2004);
however, moiramide B is not used clinically presumably due to toxicity effects. Thus,
there is a need to find other more specific potent inhibitors of carboxyltransferase that
could serve as antibiotics. The tetracyclines could serve as a lead compounds in the
design of more effective pharmaceutical agents targeting the carboxyltransferase
component of acetyl-CoA carboxylase.
References
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the carboxyltransferase subunit of acetyl-CoA carboxylase: implications for active site
communication. Protein Sci 17:34-42.
Bilder PS, Lightle G, Bainbridge J, Ohren B, Finzel F, Sun S, Holley L, Al-Kassim C,
Spessard M, Melnick M, Newcomer M, and Waldrop GL. (2006) The structure of the
carboxyltransferase component of acetyl-CoA carboxylase reveals a zinc-binding motif
unique to the bacterial enzyme. Biochemistry 45:1712-1722.
Blanchard CZ, and Waldrop, GL. (1998) Overexpression and kinetic characterization of
the carboxyltransferase component of acetyl-CoA carboxylase. J Biol Chem 273:1914019145.
Campbell JW, and Cronan JE Jr. (2001) Bacterial fatty acid biosynthesis: targets for
antibacterial drug discovery. Annu Rev Microbiol 55:305-332.
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Chopra I and Roberts M. (2001). Tetracycline antibiotics: mode of action, applications,
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Cleland WW. (1979) Statistical analysis of enzyme kinetic data. Methods Enzymol
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Cleland WW. (1990) Steady-State Kinetics, p. 117-119. In: D. S. Sigman and P. D. Boyer
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Cronan J E Jr, and Waldrop GL. (2002) Multi-subunit acetyl-CoA carboxylases. Prog
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Diacovich L, Mitchell D L, Pham H, Gago G, Melgar MM, Khosla C, Gramajo H, and
Tsai SC. 2004. Crystal structure of the beta-subunit of acyl-CoA carboxylase: structurebased engineering of substrate specificity. Biochemistry 43:14027-14036.
Duggar BM. (1948) Aureomycin. Annals of the New York Academy of Sciences 51:177181.
Florova G, Kazanina G, and Reynolds KA. (2002) Enzymes involved in fatty acid and
polyketide biosynthesis in Streptomyces glaucescens: role of FabH and FabD and their
acyl carrier protein specificity. Biochemistry 41:10462-10471.
Freiberg C, Brunner NA, Schiffer G, Lampe T, Pohlmann J, Brands M, Raabe M,
Habich D, and Ziegelbauer K. (2004) Identification and characterization of the first class
of potent bacterial acetyl-CoA carboxylase inhibitors with antibacterial activity. J Biol
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Guchhait RB, Polakis SE, Dimroth P, Stoll E, Moss J, and Lane MD. (1974) Acetyl
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Hertweck C, Luzhetskyy A, Rebets Y, and Bechthold A. (2007) Type II polyketide
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James ES, and Cronan JE. (2004) Expression of two Escherichia coli acetyl-CoA
carboxylase subunits is autoregulated. J Biol Chem 279:2520-2527.
Levert KL, and Waldrop G L. (2002) A bisubstrate analog inhibitor of the
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Nelson M. (2001) The chemistry and cellular biology of tetracyclines., p. 3-64. In
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121
CHAPTER 4
CONCLUSION
Fatty acids are the most energetically expensive modules to biosynthesize; thus,
the regulation of fatty-acid production is tightly controlled via multiple mechanisms to
match the growth rate of cells (Zhang and Rock, 2009). And though it is unlikely that a
single paradigm exists for regulation of fatty-acid synthesis in prokaryotes, this work
reveals an interesting mechanism enabling Escherichia coli to maintain stoichiometric
amounts of both subunits of carboxyltransferase despite the separate chromosomal
locations of the two genes.
Much of this work was inspired by the three-dimensional structural information of
E. coli and Staphylococcus aureus carboxyltransferase revealing a zinc finger, or
presumptive DNA-binding domain (Bilder et al., 2006). After I confirmed DNA binding,
much to my surprise, I discovered that DNA and heparin inhibit the reaction catalyzed by
carboxyltransferase, as does a class of antibiotics, tetracycline. The results from this
tetracycline study led me to propose a regulatory role for tetracycline in fatty-acid
biosynthesis in the tetracycline-producing organisms. Furthermore, I suggest that
carboxyltransferase is an alternate, albeit minor, target in the mechanism of action for the
antibiotic tetracycline. The final aspect of my work revealed RNA was both an inhibitor
and a ligand of carboxyltransferase.
Though DNA-binding studies revealed low-affinity, cooperative and nonspecific
interactions, a RNA-binding study revealed non-cooperative binding with a 7-fold higher
affinity to the transcripts of AccA/D. Just as this work built upon the carboxyltransferase
structural information and was made possible by the overexpression, purification and
122
real-time spectrophotometric assay, it provided the lead-in for further characterization of
RNA binding and the discovery of the biological target.
Subsequent Studies
Upon completion of my work, RNA binding and the carboxyltransferase zinc
finger became the focus of future investigations. Meades, et. al. concluded that the
enzyme interacts with RNA in situ via the zinc finger and that this motif is necessary for
catalysis (Meades, et. al., 2010). He demonstrated that in vitro translation of AccA/D is
inhibited upon the addition of carboxyltransferase, and this inhibition is reversed with
acetyl-CoA. These results were consistent with DNA-binding studies that showed a loss
of DNA binding by the enzyme in the presence of the substrates. Likewise, and again
consistent with DNA-binding studies, mRNA encoding either subunit of
carboxyltransferase inhibits catalysis. Consequently, Meades, et. al. proposed these
seemingly independent occurrences are actually part of a coordinated negative feedback
mechanism of control for acetyl-CoA carboxylase in E. coli that hinges on concentrations
of acetyl-CoA (Figure 4-1).
The model demonstrates that under conditions of limited nutrient growth and low
intracellular acetyl-CoA the enzyme binds mRNA from both subunits, attenuating
translation and catalysis (Meades et. al., 2010). Conversely, in the presence of high
nutrients the intracellular concentration of acetyl-CoA rises and mRNA is displaced from
the enzyme, providing for rapid translation of previously synthesized transcripts. Most
significantly, this translational regulation model illustrates the mechanism whereby E.
123
coli produces stoichiometric amounts of both subunits of carboxyltransferase despite the
fact that these genes are within separate operons.
Figure 2.9 Proposed mechanism for autoregulation of CT enzymatic activity and mRNA
Figure translation.
4-1. Negative
Feedback Model
for Autoregulation
Translation
and Catalysis
is
The RNA-binding
and catalytic
functions of CT areof
sensitive
to the metabolic
state
Centered
oncell.
the During
Intracellular
of Acetyl-CoA.
Theisleft
half of
the figure
of the
stationaryConcentration
phase when de novo
fatty acid synthesis
unneeded,
glucose
and
thuslimited
acetyl-CoA
levels growth
are low, allowing
CT to and
bind low
the mRNA
for its subunits,
attenuating
represents
nutrient
conditions,
intracellular
acetyl-CoA
expressionwhereas
of the enzyme
as wellhalf
as enzymatic
activityrepresents
on remaininghigh
acetyl-CoA
stores.
concentration,
the right
of the figure
nutrient
andDuring
log phase growth membrane biogenesis is needed. Glucose is abundant, increasing the
intracellular
acetyl-CoA
levels. for fatty acid synthesis. Acetyl-CoA binds to CT permitting
availability
of acetyl-CoA
translation of the mRNA for the ! and " subunits. This "dimmer switch" model gives the cell a
means of rapid response to changes in cellular metabolic state.
Given the high affinity carboxyltransferase demonstrated toward its own
is not found
in eukaryotic
! and "virtually
subunits are
to one
transcripts
and the
fact thatcarboxyltransferase
the enzyme wasbecause
noted the
to have
nofused
affinity
for another
another as well as to the BC and BCCP components to form one polypeptide chain and because
mRNA (encoding an EF-Ts), an obvious question arose: What recognizable features in
the zinc domain is absent (Bai et al, 1986). The one exception to the multidomain homomeric
the AccA/D transcripts permit tight binding? The answer came after extrapolating the
ACCase found in eukaryotes is the heteromeric ACCase found in plastids. The chloroplast
chemical
symmetry
in the
carboxyltransferase-catalyzed
tofour
thesubunits
structural
ACCase
of all plants
except
for the grasses (Graminae) consistsreaction
of the same
as E.
coli ACCase,
butthe
theirsubunits
genetic origins
are different
(Nikolau
et al,catalysis,
2003). Theenolate-like
BC, BCCP, and
symmetry
between
(Waldrop,
2011).
During
anions
CT ! peptides originate from nuclear-encoded genes and are shuttled to the chloroplast (Bao et
form on acetyl-CoA and biotin. Both subunits of carboxyltransferase have oxyanion holes
al, 1997; Sun et al, 1997; Ke et al, 2000). The CT " protein, however, is expressed from a
(Bilder et al., 2006) that stabilize these enolate species. Likewise, within both genes,
63
124
similar regions exist that code for the oxyanion holes, and it is these “symmetrical boxes”
along with coding regions upstream of these boxes that are described as necessary for
high-affinity mRNA binding by carboxyltransferase (Meades et. al., 2010).
Future Directions
The overall goal of this research is to fully understand the mechanisms of control
for acetyl-CoA carboxylase in E. coli, yet questions warranting further investigation
remain. Over the next few paragraphs, several analytical techniques that may provide
answers to these questions are discussed. However, the utility of these experiments lies in
their capacity to provide structural information for both transcripts and to trap
carboxyltransferase bound to nucleic-acid substrate(s).
The first unanswered questions are: Does carboxyltransferase bind the AccA/D
transcripts in vivo? If so, under what environmental conditions? And, does the enzyme
bind additional nucleic-acid ligands? Though it has been shown, via an in vitro
transcription/ translation assay, that carboxyltransferase does inhibit translation of
AccA/D, and not translation of a 32kDa EF-Ts (Meades et. al., 2010), it has not been
shown that the enzyme binds AccA/D in vivo. Thus, a gentle purification of a
carboxyltransferase:RNA complex, using the incorporation of high-affinity epitope tags
engineered into AccA/ AccD, may help to elucidate the in vivo nucleic-acid binding
target(s). The co-purified complex can be UV crosslinked and the bound mRNA
sequenced (Puig et. al., 2001).
Another curious question is: If carboxyltransferase were exposed to a large pool
of random sequence RNA molecules, what sequence(s) would yield the tightest binding?
125
Systematic Evolution of Ligands by EXponential enrichment, or SELEX, may help
answer this question. Though this experiment is an in vitro selection technique used to
screen for nucleic-acid ligands (Klug and Famulok, 1994), a high-affinity RNA could be
sequenced and compared against the genome, thus producing likely in vivo substrates. It’s
possible that a high-affinity RNA molecule(s) amplified by SELEX would be similar in
sequence to AccA and/ or AccD, thus reinforcing the theory that carboxyltransferase binds
the transcripts that encode both subunits in vivo.
Obtaining co-crystal structures of carboxyltransferase and the transcript for either
subunit appears unlikely given the large size of the transcripts; therefore, other means of
obtaining three-dimensional structural information is necessary. Insights regarding the
interaction of the zinc-finger domain and mRNA may be gleaned with hydrogenexchange mass spectrometry, as this technique may help map the nucleic-acid-binding
site(s) on the enzyme (Hoofnagle, et al., 2003). This experiment may reveal whether one
mRNA spans both zinc-fingers or if one molecule of enzyme binds a single or multiple
(separate) transcript(s).
To further support the theory that the symmetry boxes within the AccA/D
transcripts are the structurally recognizable features that drive high-affinity binding by
carboxyltransferase, a better characterization of these regions is necessary. In doing so,
comparing the secondary structures of these symmetry boxes and their upstream regions
remains essential. Two experiments that may help elucidate secondary structure
characteristics in these regions are: in-line probing (Gopinath, 2009) and selective 2'hydroxyl acylation analyzed by primer extension, or SHAPE analysis (Merino et al.,
2005). Hopefully, results from these experiments may allow for the design of RNA
126
oligomers, identical to the portions of the transcripts thought to interact with
carboxyltransferase, for co-crystallographic studies.
Follow-up investigations shedding light on the biological significance of
tetracycline-inhibiting carboxyltransferase remain relevant. If obtaining novel derivatives
of tetracycline is possible, inhibition studies may uncover better enzyme inhibitors.
Furthermore, if a tight-binding inhibitor were discovered, solving a co-crystal structure
may allow for visualization of the atomic contacts. Taken together, these results could
guide the design of subsequent-generation, high-potent inhibitors of the enzyme, while a
better understanding of the mechanisms of inherent or acquired bacterial resistance to
tetracycline may help us design lasting antibiotics in this class.
References
Bilder P, Lightle S, Bainbridge G, Ohren J, Finzel B, Sun F, Holley S, Al-Kassim L,
Spessard C, Melnick M, Newcomer M, and Waldrop GL. (2006). The structure of the
carboxyltransferase component of acetyl-CoA carboxylase reveals a zinc-binding motif
unique to the bacterial enzyme. Biochemistry 45, 1712-1722.
Gopinath SC. (2009). Mapping of RNA-protein interactions. Anal Chim Acta 636, 117128.
Hoofnagle AN, Resing KA, and Ahn NG. (2003). Protein analysis by hydrogen exchange
mass spectrometry. Annu Rev Biophys Biomol Struct 32,1-25.
Klug SJ, and Famulok M. (1994). All you wanted to know about SELEX. Mol Biol Rep
20, 97-107.
Meades G Jr, Benson BK, Grove A, and Waldrop GL. (2010). A tale of two functions:
enzymatic activity and translational repression by carboxyltransferase. Nucleic Acids Res
38, 1217-1227.
Merino EJ, Wilkinson KA, Coughlan JL, and Weeks KM. (2005). RNA structure analysis
at single nucleotide resolution by selective 2'-hydroxyl acylation and primer extension
(SHAPE). J Am Chem Soc 127, 4223-4231.
127
Puig O, Caspary F, Rigaut G, Rutz B, Bouveret E, Bragado-Nilsson E, Wilm M, and
Seraphin B. (2001). The tandem affinity purification (TAP) method: a general procedure
of protein complex purification. Methods 24, 218-229.
Zhang YM and Rock CO. (2009). Transcriptional regulation in bacterial membrane lipid
synthesis. J Lipid Res 50, S115-S119.
Waldrop GL (2011). The Role of Symmetry in the Regulation of Bacterial
Carboxyltransferase. BioiMol Concepts 2, 47-52
128
APPENDIX A
LETTER OF PERMISSION
The second chapter of this dissertation was published (in modified form) in the
journal Protein Science. Permission to reprint was requested via email and is provided,
along with the approval, below.
-----Initial Message----From: Brian Benson [mailto:pooh8honey@gmail.com]
Sent: Tuesday, June 10, 2008 5:08 PM
To: 'proteinscience@proteinsociety.org'
Subject: request for manuscript to be included in dissertation
GreetingsI am requesting permission to include a manuscript that was printed in your journal, as a
chapter in my dissertation. I, Brian K. Benson, am the first author and the manuscript
was printed in January of 2008; the reference is Protein Sci. 2008 17: 34-42.
Please let me know if you are able to accommodate this request or if you need any
additional information.
Thanks and have a great day----- Message Response----From: Brian Matthews [mailto:brian@uoxray.uoregon.edu]
Sent: Wednesday, June 11, 2008 5:24 PM
To: pooh8honey@gmail.com
Subject: Copyright Permission
RE: Benson, B.K., Meades Jr., G., Grove, A. and Waldrop, G.L. (2008) DNA inhibits
catalysis by the carboxyltransferase subunit of acetyl-CoA carboxylase: Implications for
active site communication. Protein Sci.
17, 34-42.
Dear Mr. Benson:
In reply to your request of June 10, 2008, permission is hereby given to reprint the abovereferenced material. Please include the acknowledgment "Courtesy of Protein Science".
With best regards,
Brian Matthews
Editor, Protein Science
129
APPENDIX B
ABBREVIATIONS
ACC
acetyl-CoA carboxylase
Ala
alanine
AOPP
aryloxyphenoxypropionate
Arg
arginine
Asp
aspartic acid
BCCP
biotin carboxyl carrier protein
BirA
biotin holoenzyme synthetase
BiSA
bisubstrate analog
C2′
carbon within ureido ring, see Figure 1-1.
C-terminal
carboxyl-terminal
CHD
cyclohexanediones
EF-Ts
elongation factor thermo stable
E. coli
Escherichia coli
EMSA
electrophoretic mobility shift assay
FAS-I
type I fatty acid synthase
FAS-II
soluble enzymes in prokaryotes and plant chloroplasts
IHF
integration host factor
I, Ile
isoleucine
Lys
lysine
Met
methionine
N1′
nitrogen within urido ring, see Figure 1-1.
130
N3′
nitrogen within urido ring, see Figure 1-1.
N-terminal
amino-terminal
NMR
nuclear magnetic resonance
Pi
Inorganic phosphate
T, Thr
threonine
Tyr
tyrosine
Val
valine
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APPENDIX C
ANALYTICAL TECHNIQUES APPLIED TO NUCLEIC-ACID BINDING PROTEINS
It became clear at the beginning of this work that studying DNA-protein
interactions via an agrose gel would be insufficient, so we pursued a more acceptable
method. As electrophoretic mobility shift assays, or EMSAs, were essential to this thesis,
yet not previously performed in our laboratory, a literature review concerning the
detection and study of DNA-binding proteins was necessary and proved insightful.
Several methods have been developed and enhanced to the point of current gel shift
assays. Initially, two groups (Coombs and Pearson, 1978; Riggs et al., 1970) developed
the filter assay to retain DNA-protein complexes on a nitrocellulose filter. Subsequently,
an abortive initiation assay was developed to visualize an RNA polymerase reaction in
1978 using paper chromatography (McClure et al., 1978). Finally, in 1979, one group
(Chelm and Geiduschek, 1979), in an agrose gel electrophoresis assay, studied the very
stable ternary transcription complexes. These complexes were composed of RNA
polymerase, DNA and radioactively labeled nascent RNAs. This method provided
information on polymerase binding and on promoter mapping by identification of DNArestriction fragments. These DNA-restriction fragments generated measurable ternary
complexes.
Though several groups prior to 1981 had reported methods for studying
protein:DNA complexes, it was Garner and Revzin (Garner and Revzin, 1981) who first
described a new protein:DNA gel shift, or EMSA, as “a new gel electrophoresis
technique for more quantitative studies of specific protein:DNA interactions.” They
explained that this method could separate unbound DNA fragments from complexes by
132
electrophoresis in polyacrylamide gels. The amount of unbound DNA is then determined
by densitometry of the gel. They proclaimed that this fast and easy method used small
amounts of reagents.
Around the same time, Fried and Crothers also described the utility of EMSA. In
an article also published in Nucleic Acids Research only months after Garner and Revzin
published their findings, Fried and Crothers (Fried and Crothers, 1981), who have since
been credited with developing this technique, explained that “the protein is mixed with
one or more DNA fragments, and protein:DNA complexes are resolved as discrete bands
by polyacrylamide gel electrophoresis, with mobilities decreasing as a function of the
number of proteins bound to a given DNA fragment.” They suggested that, under
minimal ionic strength conditions, the gel matrix itself stabilizes the complex against
dissociation. Potential factors for strengthening the complex include: volume exclusion,
direct interaction with gel matrices, and the reduction of water activity by the gel. While
one may wonder if this technique could cause artificially high affinities for protein:DNA
complexes, the group has reported that the parameters appear no different in the gel than
in an ordinary buffer (Fried and Bromberg, 1997). Fried and Crothers added that the
benefit of the EMSA technique was the measuring, via microdensitometry, of stained gels
and gel autoradiograms for the quantification of the protein and DNA present in each
complex (Fried and Crothers, 1981). Furthermore, the technique permitted determination
of the stoichiometric ratio of protein specifically bound to DNA, and it facilitated precise
measurement of the relative binding affinities of a given protein for two different nucleic
acids. Next, dissociation kinetics is readily obtainable for long-lasting protein:DNA
complexes. Most importantly, in addition to use in quantitative binding studies, this
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technique should simplify identification of proteins that bind specifically to a given DNA
sequence or of restriction fragments containing sequences recognized by a particular
protein.
References
Chelm BK and Geiduschek EP. (1979). Gel electrophoretic separation of transcription
complexes: an assay for RNA polymerase selectivity and a method for promoter
mapping. Nucleic Acids Res 7, 1851-1867.
Coombs DH, and Pearson GD. (1978). Filter-binding assay for covalent DNA-protein
complexes: adenovirus DNA-terminal protein complex. Proc Natl Acad Sci USA 75,
5291-5295.
Fried M and Crothers DM. (1981). Equilibria and kinetics of lac repressor-operator
interactions by polyacrylamide gel electrophoresis. Nucleic Acids Res 9, 6505-6525.
Fried MG and Bromberg JL. (1997). Factors that affect the stability of protein-DNA
complexes during gel electrophoresis. Electrophoresis 18, 6-11.
Garner MM and Revzin A. (1981). A gel electrophoresis method for quantifying the
binding of proteins to specific DNA regions: application to components of the
Escherichia coli lactose operon regulatory system. Nucleic Acids Res 9, 3047-3060.
McClure WR, Cech CL, and Johnston DE. (1978). A steady state assay for the RNA
polymerase initiation reaction. J Biol Chem 253, 8941-8948.
Riggs AD, Suzuki H, and Bourgeois S. (1970). Lac repressor-operator interaction. I.
Equilibrium studies. J Mol Biol 48, 67-83.
134
VITA
Brian Kimbrough Benson was born in Lafayette, Louisiana, on March 12, 1976.
He attended high school at Scotlandville Magnet in Baton Rouge, Louisiana, graduating
in 1994. After serving in the infantry of the United States Marines, he owned and
operated a small contracting company. Shortly after his wife became a nurse, he began to
understand the limitations of cancer treatments and was then inspired to learn biology and
medicine. At this time, he enrolled in the biochemistry program at Louisiana State
University and was awarded a Bachelor of Science in 2002. Directly thereafter, he began
graduate school at Louisiana State University and a year later began working in the
laboratory of Dr. Grover Waldrop. In the summer of 2004 Benson was diagnosed with
Hodgkin’s lymphoma and underwent six months of chemotherapy, after which he was
considered cancer-free for four years. In the summer of 2008, he was diagnosed with a
presumptive relapse of cancer. For staging and biopsies, he underwent surgery while
being readied for a bone marrow transplant. To his great surprise and relief, no cancer
was found. His experiences hardened his determination to thoroughly understand cancer.
In 2007, after submitting manuscripts for publication, he enrolled in medical school at
Tulane University. He will complete the requirements for the Doctor of Philosophy in
biochemistry in July 2011 and will complete a residency in rural family medicine in
2015. Shortly after starting the practice, he plans to partner with a postdoctoral researcher
in oncology and another in immunology, and together they will work toward developing
future oncology diagnostics and treatments.
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