U N I V E R S I T Y O F C O P E N H A G E N D E P A R T M E N T O F B I O L O G Y
PhD thesis Rasmus Nielsen Klitgaard, M.Sc.
Antibiotic Drug Discovery Potentiation of the quinolones and targeting the initiation of DNA replication
This thesis has been submitted to the PhD School of The Faculty of Science, University of Copenhagen,
Denmark, 28. February 2018.
Dan Andersson
Department of Medical Biochemistry and Microbiology
University of Uppsala, Sweden.
Mogens Kilstrup
Department of Biochemistry and Biomedicine
Metabolic Signaling and Regulation
Danish Technical University, Denmark.
Signe Lo Svenningsen
Department of Biology
Biomolecular Sciences
University of Copenhagen, Denmark.
Submitted: 28.02.2018
Academic advisor Anders Løbner-Olesen
Department of Biology
Functional Genomics
University of Copenhagen, Denmark.
Assessment committee
1
Acknowledgements
First, I would like to thank my supervisor Anders Løbner-Olesen for his excellent support and
guidance throughout my PhD. I have highly appreciated that Anders has been available more or
less every day and gladly discussed any questions I might have had.
I would also like to thank Godefroid Charbon, not only for our collaboration on paper II
presented in this thesis, but also for always taking time to discuss and give advice on my other
projects. It has been greatly cherished.
Furthermore, I would like to thank the staff at Naicons srl. and all of the people who have been
part of the ALO lab: Thomas T. Thomsen, Jakob Frimodt-Møller, Maria S. Haugan, Christoffer
Campion, Anna E. Ebbensgard, Michaela Lederer, Henrik Jakobsen, Leise Riber and Belén M.
Chamizo.
A thanks, should also be given to my bachelor student, Anne Kristine Schack, who contributed
to the construction of the screening system presented in paper III.
Finally, I would like to thank my girlfriend, Marie, my family and my friends for their great
support and interest in my work.
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Abstract – English
Antibiotic resistance has been deemed as one of the biggest threats to the global public health by the
World health Organization. In 2050, an estimated 10 million deaths per year will be attributed to
antimicrobial resistance, thus proper action needs to be taken to stop this negative development. An
important mean in the arms race against antibiotic resistance is the discovery and development of novel
antibiotics, but also preserving the efficacy of the antibiotics that are already in clinical use.
In paper I, we search for ciprofloxacin helper drug targets in an effort to preserve the use
of this widely applied antibiotic. Using a combined genetic and transcriptomic approach, the AcrAB-TolC
efflux pump and the SOS response genes, RecA and RecC, are identified as potential targets for helper
drugs in Escherichia coli strains with low-level ciprofloxacin resistance. In addition, our results also
indicate that reversing high-level ciprofloxacin resistance is likely not plausible.
In paper II, we present two novel cell based screens for identifying inhibitors of the
chromosomal DNA replication initiation in bacteria. The screens are based on growth rescue of cells that
rigorously over-initiate the DNA replication, due to either increased regeneration of the active ATP
bound form of the replication initiator protein DnaA, or by being deficient in the process known as
regulatory inactivation of DnaA (RIDA). Screening a library of 400 microbial extracts, revealed the iron
chelator deferoxamine as a compound that rescues the growth of over-initiating cells. Albeit not by
decreasing the replication initiation frequency, but by reducing the production of reactive oxygen
species. Substantiating the model that oxidative DNA damage and its repair promotes the lethal action
of hyper-replication.
In paper III, we constructed and verified a novel high throughput, cell based, fluorescence
screen for inhibitors of chromosome replication initiation in bacteria. The screen utilizes an E. coli
mutant that is resistant to replication initiation inhibitors and holds a fluorescence reporter system for
DNA replication inhibitors. This screen was also subjected to the above-mentioned library of microbial
extracts, though it did not lead to any positive hits.
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Abstract – Danish
Verdens Sundheds organisationen, WHO, har udnævnt antibiotika resistens til at være en af de største
trusler mod det globale sundhedssystem. Det er blevet estimeret at i 2050 vil ca. 10 millioner dødsfald
årligt være associeret med antibiotika resistens. Det er derfor yderst vigtigt at der allerede nu tages de
nødvendige initiativer til at begrænse denne negative udvikling. En af de væsentligste faktorer i kampen
mod antibiotika resistens er udviklingen af nye antibiotika, samt at præservere virkningen af de antibiotika
som allerede bruges i klinikken.
I et forsøg på at præservere den kliniske anvendelighed af det ofte benyttede antibiotika
ciprofloxacin. Søger vi i artikel I efter gener i Escherichia coli hvis deletion reverserer ciprofloxacin resistens
og dermed kan bruges som mål for ciprofloxacin hjælpestoffer. Ved hjælp af genetisk deletions analyse
identificerede vi efflux pumpen, AcrAB-tolC, samt SOS-respons proteinerne, RecA og RecC som mulige mål
for ciprofloxacin hjælpestoffer i lav-resistente stammer af E. coli. Ydermere viste vores resultater også at
det formentlig ikke er muligt at reverserer ciprofloxacin resistens i høj-resistente stammer af E. coli.
I artikel II præsenterer vi to nye screeningssystemer til at identificere inhibitorer af
initieringen af kromosomal DNA replikation i bakterier. Disse to screeningssystemer er baseret på celler der
over-initierer DNA replikationen, via henholdsvis forhøjet regenerering af den ATP bundne form af
initieringsproteinet DnaA eller mangel på processen kendt som regulativ inaktivering af DnaA (RIDA).
Denne over-initiering er lethal for cellerne. Under screening af et bibliotek bestående af 400 mikrobielle
ekstrakter, identificerede vi jern chelatoren deferoxamine, som et stof der kan redde væksten af celler der
over-initierer replikationen. Dog ikke ved at nedsætte initierings frekvensen, men ved at reducere
produktionen af reaktive oxygen radikaler. Hvilket ydermere fast slår modellen, at oxidativ DNA skade og
dets reparation medierer celledød i bakterier det over-initierer DNA replikationen.
I artikel III konstruerede og verificerede vi endnu et nyt screeningssystem til inhibitorer af
DNA replikations initieringsprocessen. Denne screen består af en E. coli mutant der er resistent over for
stoffer der blokerer replikations initierings processen og samtidig indeholder et fluorescens baseret
reporter system der aktiveres af replikations initierings inhibitorer. Denne screen blev også testet mod det
ovennævnte bibliotek af mikrobielle ekstrakter, men gav ingen positive hits.
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List of papers
Paper I Can Ciprofloxacin Resistance be Reversed by Helper Drugs? Rasmus N. Klitgaard, Bimal Jana, Luca Guardabassi, Karen Leth Nielsen and Anders Løbner-Olesen.
Paper II A strategy for finding DNA replication inhibitors in E. coli identifies iron chelators as molecules that promote survival of hyper-replicating cells. Godefroid Charbon, Rasmus Nielsen Klitgaard, Charlotte Dahlmann Liboriussen, Peter Waaben
Thulstrup, Sonia Ilaria Maffioli, Stefano Donadio and Anders Løbner-Olesen.
Paper III A Novel Fluorescence Based Screen for Inhibitors of the Initiation of DNA Replication in Bacteria. Rasmus N. Klitgaard and Anders Løbner-Olesen.
Papers not included in the thesis Ciprofloxacin intercalated in fluorohectorite clay: Identical pure drug activity and toxicity with higher adsorption and controlled release rate. E. C. dos Santos, Z. Rozynek, E. L. Hansen, R. Hartmann-Petersen, R. N. Klitgaard, A. Løbner-Olesen, d L.
Michels, A. Mikkelsen, T. S. Plivelic, H. N. Bordallo and J. O. Fossum.
Mutations in the Bacterial Ribosomal Protein L3 and Their Association with Antibiotic Resistance. Rasmus N. Klitgaard, Eleni Ntokou, Katrine Nørgaard, Daniel Biltoft, Lykke H. Hansen, Nicolai M.
Trædholm, Jacob Kongsted, Birte Vester.
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Table of contents
ACKNOWLEDGEMENTS ................................................................................................... 2
ABSTRACT – ENGLISH ..................................................................................................... 3
ABSTRACT – DANISH ....................................................................................................... 4
LIST OF PAPERS ............................................................................................................... 5
TABLE OF CONTENTS ...................................................................................................... 6
A BRIEF HISTORY OF ANTIBIOTICS ................................................................................ 9
The early days ................................................................................................................................................................... 9
The golden age of antibiotics ........................................................................................................................................... 9
The present and future of antibiotics ............................................................................................................................ 10
PART I: POTENTIATION OF THE QUINOLONES ........................................................... 11
Discovery and development of the quinolones ............................................................................................................. 11
The quinolone targets ..................................................................................................................................................... 12
Mechanism of action ....................................................................................................................................................... 14 Fragmentation of the bacterial chromosome ................................................................................................................ 14
Reactive oxygen species and quinolone lethality .......................................................................................................... 15 Are ROS involved in quinolone lethality? ................................................................................................................... 15
The SOS response, an endogenous defense against quinolones .................................................................................. 16 Regulation and induction of the SOS response ............................................................................................................ 17 Repair of quinolone mediated double stranded DNA breaks by the SOS response ..................................................... 17
Quinolone resistance ....................................................................................................................................................... 18 Target site mutations .................................................................................................................................................... 18 Non-target site mutations involved in quinolone resistance ........................................................................................ 19 Plasmid mediated quinolone resistance ....................................................................................................................... 19
Reversing antibiotic resistance by helper drugs .......................................................................................................... 22 Potential targets for potentiation of quinolones ........................................................................................................... 23
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PART II: TARGETING THE INITIATION OF CHROMOSOMAL DNA REPLICATION IN BACTERIA ........................................................................................................................ 23
Initiation of chromosomal DNA replication in E. coli ................................................................................................. 24 DNA replication and the cell cycle. ............................................................................................................................. 24 Initiation of replication ................................................................................................................................................ 25 The origin of replication .............................................................................................................................................. 26 The initiator protein DnaA ........................................................................................................................................... 27
Replication initiation by DnaAATP ................................................................................................................................. 29 Formation of the DnaAATP initiation complex ............................................................................................................. 29 DUE unwinding ........................................................................................................................................................... 30 DnaB helicase loading ................................................................................................................................................. 31
Regulation of the replication initiation ......................................................................................................................... 31 The dual role of DiaA in regulating replication initiation ............................................................................................ 32 Regulatory inactivation of DnaAATP (RIDA) ............................................................................................................... 33 datA-dependent DnaAATP-hydrolysis (DDAH) ............................................................................................................ 33 Regulation of DDAH activity ...................................................................................................................................... 34 SeqA, a negative regulator of the replication initiation ............................................................................................... 35 Rejuvenation of the cellular DnaAATP pool .................................................................................................................. 36
The lethal action of severe over-initiation of the DNA replication ............................................................................. 39
Targeting the Initiation of replication .......................................................................................................................... 40
PAPER I: CAN CIPROFLOXACIN RESISTANCE BE REVERSED BY HELPER DRUGS?............................................................................................................................ 42
PAPER II: A STRATEGY FOR FINDING DNA REPLICATION INHIBITORS IN E. COLI IDENTIFIES IRON CHELATORS AS MOLECULES THAT PROMOTE SURVIVAL OF HYPER-REPLICATING CELLS. ....................................................................................... 57
PAPER III: A NOVEL FLUORESCENCE BASED SCREEN FOR INHIBITORS OF THE INITIATION OF DNA REPLICATION IN BACTERIA. ....................................................... 94
DISCUSSION .................................................................................................................. 102
Potentiation of the quinolones ..................................................................................................................................... 102
Targeting the commencement of DNA replication in bacteria ................................................................................. 104
Why is severe over-initiation of the DNA replication lethal? ................................................................................... 106
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CONCLUSIONS .............................................................................................................. 107
FUTURE PERSPECTIVES .............................................................................................. 107
BIBLIOGRAPHY ............................................................................................................. 109
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A brief history of antibiotics
The early days
Most people, even without a background within life science, have heard the intriguing story of how
Alexander Fleming by coincidence contaminated his agar plates with mould and discovered penicillin
back in 1929 (1). Of less common knowledge is the pioneering work of Alexander Ehrlich and Sahachiro
Hata, which led to the discovery of salvarsan, in 1909, a novel drug for treating the sexual transmitted
disease syphilis that is caused by the spirochete Treponema pallidium (2). Salvarsan and its derivative
neosalvarsan, were the most prescribed drugs until they were replaced by penicillin in the 1940s (3).
The large-scale screening method used by Ehrlich and Hata in the discovery of salvarsan, became the
gold standard for identifying novel drugs and led to the discovery of the first sulfa drug in 1934,
sulfonamidochrysoidine, a precursor of the active compound sulfanilamide, which inhibits folic acid
synthesis in bacteria (3, 4).
The golden age of antibiotics
The discovery of the sulfa drugs and the release of penicillin for clinical use kick-started a period of 30
years known as the golden age of antibiotics (1940-1970), in which almost all of the antibiotic drug
classes used in the clinic today were discovered (see Figure 1) (5, 6). Most of the antibiotics discovered
in this period were isolated from natural extracts from different microorganisms. Following the isolation
Figure 1: The top panel indicates the time at which different antibiotics and classes of antibiotics were discovered. The bottom
panel, indicates when resistance was observed for the given antibiotics. Modified from Clatworthy et al., 2007.
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of streptomycin, in 1944, from the soil growing filamentous bacteria Streptomyces griseus. Soil samples
were collected from around the world and in 1952 the vancomycin producing Streptomyces orientalis
was isolated from a soil sample from Borneo, leading to the release of vancomycin for clinical use in
1958 (5).
Despite of its name it was also in the golden age that it became evident that clinical
antibiotic resistance would become a problem. In 1945, Alexander Fleming, during his Nobel lecture,
warned that underdosing of penicillin could potentially lead to the development of resistance (7). In the
decade following Flemings warning, it became apparent that antibiotic resistance was a problem. To
overcome resistance scientists started to make derivatives of already know drugs, this led to the
development of antibiotics that were impervious to the resistance mechanisms and in some cases
improved the pharmacodynamics and pharmacokinetics of the drugs (5). However, it was also the start
of a race between the evolution of antibiotic resistance and the development and discovery of
antibiotics. A race that currently seems to be led by the bacteria.
The present and future of antibiotics
In the last 40 years, the only truly novel class of antibiotics that has been introduced into the clinic are
the oxazolidinones, initially represented by the synthetic compound linezolid that was released in 2000
(8). Due to its synthetic nature it was anticipated that linezolid resistance would evolve slowly (9). This
presumption unfortunately turned out to be wrong, as soon after its release, linezolid resistance was
identified in clinical isolates of Staphylococcus aureus and several enterococcus species (10).
As of December 2017 an estimated 48 antibiotics are in phase I to III clinical trials. Most
of these antibiotics are derivatives of known antibiotics, almost half do not target pathogens listed as
being a critical threat by the World Health Organisation (WHO) and even fever are expected to display
activity against the multi drug resistant group of Gram negative ESKAPE pathogens (11). Considering
that on average only one third of these antibiotics will make it through the clinical trials and become a
marketable product, the current antibiotic pipeline is not robust enough to support the current and
future clinical need (12). In addition, a report commissioned by the government of the United Kingdom
in 2014, estimated that the annual number of deaths attributable to antimicrobial resistance would be
10 million by 2050 and that it will generate a loss of 100 trillion dollars globally (13). Even though these
numbers are only estimates, there is no doubt; antibiotic resistance is a major global health care
problem and it will only become more evident with time, if proper action is not taken.
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Part I: Potentiation of the quinolones
The quinolone class of antibiotics includes some of the most widely used and prescribed antibiotics (14-
16). Due to their popularity and misuse, quinolone resistance has become a major problem in the clinic
(17, 18). In context of the current lack of development of novel classes of antibiotics, potentiation of
already known antibiotics will be essential. At least until the antibiotic development pipeline has
become more robust. The following sections will introduce the reader to the quinolone class of
antibiotics, quinolone resistance and how quinolones might be potentiated to overcome resistance.
Discovery and development of the quinolones
In 1964, Sterling Drugs released the first compound
of a novel class of antibiotics for use in the clinic,
named nalidixic acid. Though nalidixic acid is based
on a 1,8-naphthyridone core and therefore
technically not a quinolone (see Figure 2A), it is in
general acknowledged as the first quinolone
antibacterial. The events that led to the discovery of
nalidixic acid are somewhat unclear. The story goes
that a by-product of the synthesis of the antimalarial
drug chloroquine, made at Sterling Drugs inc.,
showed antibacterial properties and contained a
quinolone core. Sterling has newer commented on
why the quinolone core was substituted for a 1,8-
napthyridone core in nalidixic acid. However, it was
likely because there had already been filled a patent,
by Imperial Chemical Industries in 1960, on a
compound similar to nalidixic acid, but with a
quinolone core. In the years after the release of
nalidixic acid, a number of follow-up drugs were
released (19, 20). The first generation of quinolones
were mainly used in treating uncomplicated urinary
Figure 2: A) Comparison of the quinolone core with
the 1,8-naphthyridone core of nalidixic acid.
Adapted from Bisacchi et al., 2015. B) The structure
of the second-generation quinolone, ciprofloxacin.
The fluorine at position C6 is marked by a blue
circle and the piperazine substituent at position C7
by a red circle.
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tract infections, as their systemic absorption was poor. In the early 1980s, the first second-generation
compounds were released, including ciprofloxacin and norfloxacin. The major differences from the first-
to the second-generation compounds were the addition of a fluorine at position C6 and a piperazine or
methyl-piperazine substituent at C7 (See Figure 2B). The addition of the fluorine, led to the term
fluoroquinolones. These two additions to the quinolone core, improved both the bacterial spectrum,
but also the pharmacokinetic and pharmacodynamics significantly (21). Since then both third and fourth
generation fluoroquinolones has made its way into the clinic. The third generation fluoroquinolones like,
levofloxacin, sparfloxacin and grepafloxacin expanded the bacterial spectrum to include streptococci
and had prolonged half-lives. The fourth generation fluoroquinolones, was the first generation with
activity against anaerobes like, bacteroides fragilis, in addition to an enhanced activity against Gram-
positives (22). Furthermore, the 8-methoxy group possessed by two of the fourth generation drugs,
gatifloxacin and moxifloxacin, eliminated the phototoxicity observed for earlier generations (22).
Throughout the rest of this thesis the term quinolone, will be
used for both first generation quinolones and the
fluoroquinolones, unless differences are specified.
The quinolone targets
The cellular pathway targeted by nalidixic acid was revealed
already in 1964. By measuring the incorporation of C14-labeled
thymine in DNA, it was shown that it inhibited the DNA
synthesis (23). Five years later, in 1969, Hane et al. genetically
mapped mutations in two distinct genes, nalA and nalB, that
conferred different levels of nalidixic acid resistance (24). nalA
was subsequently identified as gyrA, encoding the subunit of
the DNA gyrase responsible for nicking and re-ligation of the
DNA (25, 26). The DNA gyrase is not the sole target of the
quinolones, in 1990 a novel topo-isomerase, topo-isomerase IV
(topo IV), was discovered (27). Topo IV is the gene-product of
parC and parE, which have a high degree of sequence homology
with gyrA and gyrB, especially in the regions where there have
been identified mutations conferring quinolone resistance (28).
The DNA gyrase and topo IV are both type II
topoisomerases, essential for numerous processes involving
Figure 3: A, B) Crystal structure of a
topo IV cleavage complex bound by
two molecules moxifloxacin. In
green: the ParE subunits. In blue: the
ParC subunits. In red: moxifloxacin
and in yellow: the DNA strand.
Modified from Aldred et al. 2013
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nucleic acids, including DNA replication and chromosome
segregation. They modulate the topology of the DNA by
controlling the level of under- and over-winding and are able to
sort tangles and knots in the DNA. The modulation of the DNA
topology is achieved by inducing transient double-stranded
brakes in the DNA, thereby releasing the torsional stress. To
maintain the integrity of the chromosome during the opening
of the double-strand, the gyrase and topo IV binds covalently to
the generated 5´-DNA ends, creating a so-called “cleavage
complex” (see Figure 3AB). Following the cleavage reaction the
DNA is re-ligated again by the bound enzyme(21).
The gyrase and topo IV are both heterotetramers
with an A2B2 quaternary structure. The gyrase consists of two
GyrA and two GyrB subunits, while Topo IV contains two ParC
and two ParE subunits. The GyrA/ParC subunits holds the active
site tyrosine residues and are responsible for the DNA cleavage
and ligation reactions, while GyrB and ParE both contain an
ATPase domain delivering the energy for the cleavage and
ligation reaction by hydrolysis of ATP (29). Studies of crystal
structures of type II topoisomerases bound by different
quinolones have revealed that the binding is mediated by a
water-metal ion bridge (29-31), between the C3/C4 keto acid of
the quinolone and Ser83, Asp87 in GyrA, or Ser80 and Glu84 in
ParC (E. coli numbering, see Figure 4A). These findings are
supported by the fact that the most common amino acid
substitutions conferring resistance to quinolones, have been
identified at these specific amino acids (32). Notefully, the
human type II topoisomerases , hTIIα and hTIIβ, do not contain
these residues (see Figure 4B). It has therefore been proposed
that lack of the necessary amino acids to mediate the water-
metal bridge, is one of the main reasons why quinolones do not target hTIIα and hTIIβ (21).
Earlier, based on studies in E. coli, S. aureus and Streptococcus pneumoniae, it was
pressumed that the DNA gyrase was the primary target in Gram-negative bacteria, while topo IV was the
primary target in Gram-positive bacteria (33-35). However, this presumption turned out to be incorrect,
Figure 4: A) Structure of the water-metal
ion bridge between the C3/C4 keto acid of
the quinolone, the serine and either
aspartic acid or glutamic acid. B) Alignment
of GyrA and ParC (GrlA) from Acinetobacter
baumanii (Ab), Bacillus anthracis (Ba),
Escherichia coli (Ec), Staphylococcus aureus
(Sa) and Streptococcus pneumonia (Sp). In
red the serine and the acidic amino acid
that forms the water-metal ion bridge.
Note that the human homologs hTIIα and
hTIIβ do not contain the serine or the acidic
amino acid. Modified from Aldred et al.,
2014.
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as later studies have shown that the target specificity is both species and drug dependent (21). For
instance, in S. aureus nalidixic acid was shown to target the gyrase and norfloxacin preferentially topo
IV, while ciprofloxacin targeted both the gyrase and topo IV (36).
Mechanism of action
Fragmentation of the bacterial chromosome
In 1979 Kreuzer et al. proposed that quinolones were acting as poisons, corrupting the function of the
gyrase, rather than directly targeting the catalytic effect of the gyrase(37). This hypothesis turned out to
be true. Quinolones act by blocking the ability of topo IV and the gyrase to re-ligate the cleaved DNA in
the cleavage complex, in turn leading to fragmentation of the bacterial chromosome (21). The exact
events that leads to the fragmentation is still under debate. Earlier, it was believed that the quinolone
bound cleavage-complex was converted to a permanent break if hit by a replication fork or other
complexes moving along the DNA. This idea originated from the findings that eukaryotic topoisomerase
I, trapped on the DNA, created double stranded breaks when colliding with a replication fork (38, 39).
However, no one have been able to show that this is also the case in bacteria. Though it was shown that
collision between the quinolone bound cleavage-complex and the replication fork, stalled the
replication fork and rendered the quinolone-cleavage-complex in an irreversible state (40-43). These
findings spawned the idea, that stalling of the replication fork was followed by endonuclease mediated
clevage of the DNA at the replication fork (44). This model was later challenged, as halting DNA
replication, by using a temperature sensitive DnaB helicase mutant, did not affect quinolone lethality
(45). Studies of the lethal action of nalidixic acid and gatifloxacin treatment in combination with
chloramphenicol, a protein synthesis inhibitor, revealed that the lethal action of nalidixic acid was
blocked without ongoing protein synthesis, while the lethality of gatifloxacin was retained (39, 46).
These findings indicated the existence of two pathways leading to the lethal action of quinolones; a
protein synthesis dependent pathway and a pathway independent of protein synthesis. It has been
proposed that binding of quinolones to the cleavage complex destablizes the complex, thereby releasing
the double stranded DNA break from the cleavage complex (39). This model is independent of protein
synthesis and is supported by the fact, that gatifloxacin can fragment chromosomes in vitro in the
presence of purfied gyrase (39). Additionally, it has also been shown that an E. coli mutant, where the
gyrase has been destabilized by introduction of a GyrA A67S mutation, is killed by nalidixic acid in the
presence of chloramphenicol, in contrary to the wild type (39). The chromosome fragmentation
pathway dependent on protein synthesis is less clearly understood. However, protease digestion of the
14
gyrase or nuclease-mediated cleavage on either side of the cleavage complex, have been suggested to
mediate the release of the DNA from the cleavage complex (47).
Reactive oxygen species and quinolone lethality
In E. coli, deleterious reactive oxygen species (ROS) are continuously formed during respiration, when
auto-oxidation of its redox enzymes generates superoxide (O2.-) (48). To prevent accumulation of O2
.- it
is converted by superoxide dismutases to oxygen and hydrogen peroxide (H2O2). Intracellularly H2O2 can
react with iron (II), leading to generation of highly reactive hydroxyl radicals (OH•) through Fenton
chemistry (49):
+ → + + •
The generated hydroxyl radicals can essentially react with and damage most biomolecules, including
DNA, proteins and lipids. As there are no known cellular pathways degrading hydroxyl radicals, its
generation is limited by peroxidases and catalases that degrade H2O2 (50).
From 2002 to 2006, a number of papers reported that treatment of bacteria with
bactericidal antibiotics lead to heightened levels of ROS and that ROS was involved in the lethal action of
bactericidal antibiotics (51-54). In 2007, the first model for a ROS mediate cellular death pathway
induced by bactericidal antibiotics was published. It was proposed that bactericidal antibiotics stimulate
oxidation of NADPH to NAD+ by the electron transport chain, leading to a boost in superoxide
production. Superoxide mediated damage of iron-sulphur-cluster proteins then releases iron (II), which
reacts with H2O2 and generates hydroxyl radicals through the Fenton reaction. At the time, cell death
was explained by general hydroxyl radical mediated damage to proteins, lipids and DNA (55, 56). Later,
oxidation of the cells nucleotide pool, specifically generation of 8-oxo-dGTP and its incorporation into
DNA was proposed as the dominant mechanism by which ROS mediates cell death by bactericidal
antibiotics (57, 58). Proposing that ROS significantly contributed to the lethal action of bactericidal
antibiotics was controversial and it is still a matter of debate.
Are ROS involved in quinolone lethality?
The first clues indicating that quinolone treatment of E. coli led to an increase in ROS
production, came from the observation that nalidixic acid significantly increased the expression of the
superoxide dismutase, encoded by sodA (59). The increase in expression of sodA was later shown to be
mediated by activation of the soxRS regulon (60, 61), a major oxidative stress response system fund in
most Gram-negative bacteria (62, 63). Investigations of the involvement of the soxRS regulon in
quinolone resistance revealed that over-expression of soxS in E.coli and constitutive activation of the
15
soxRS regulon in salmonella enterica increased the level of quinolone resistance. However, it should be
noted that the observed resistance was likely, due to the fact that activation of SoxRS results in
posttranscriptional negative regulation of the OmpF porin, involved in quinolone transport into cells (64)
and overproduction of the AcrAB-TolC efflux pump (65).
Several different quinolones have been shown to increase ROS production, as detected
by both chemiluminescence and fluorescence methods, in E. coli, S. aureus and Enterococcus faecalis
(51-53, 55, 56). Furthermore, blocking ROS generation by either antioxidants or iron chelators lowers
the susceptibility of E. coli to some quinolones (54, 66). In addition, deletion analysis, in E. coli, of the
genes involved in H2O2 metabolism, katG, ahpCF and katE , showed that a katG, ahpCF double mutant
and a katG, ahpCF, katE triple mutant were more susceptible to ciprofloxacin than the wild-type(54).
To challenge the proposed model for ROS mediated killing by bactericidal antibiotics
described above. The efficacy of a number of quinolones was investigated under anaerobic conditions,
where ROS cannot be generated. This included the first generation quinolone, nalidixic acid and the
fluoroquinolones norfloxacin, ciprofloxacin and ofloxacin (67-69). The results showed that anaerobic
growth did not lead to an increase in MIC. However, anaerobic conditions blocked the killing by nalidixic
acid, but not by norfloxacin, ciprofloxacin or ofloxacin, though higher concentrations of norfloxacin and
ciprofloxacin were required to kill the cells when grown anaerobically (67-69). Furthermore, quenching
of ROS production by treatment with the iron chelator, dipyridyl and the reducing agent thiourea,
blocked the lethality of oxolonic acid, but only partially reduced the lethal action of moxifloxacin, while
the C8-methoxy fluorquinolone, PD161144, was unaffected. Interestingly, there is an inverse correlation
between the lethal action of quinolones under anaerobic conditions and the observed degree of protein
synthesis dependency for lethality (66, 67). Indicating that the protein dependent pathway relies on
generation of ROS, while the protein synthesis independent pathway does not (66). However, further
research is needed to elucidate the exact mechanism that connects ROS with the lethal action of the
protein synthesis dependent pathway.
The SOS response, an endogenous defense against quinolones
Maintaining genome integrity is vital for bacteria, therefore most bacteria express an inducible DNA
damage repair system termed the SOS response(70). As quinolones fragments the chromosome, they
are strong induceres of the SOS response (55, 71), which acts as a first line of defence against this group
of antibiotics. In addition, the activation of the SOS response leads to high mutation rates, which in turn
can result in occurrence of mutations conferring resistance to quinolones (32, 72). Therefore the SOS
response is a key process in both quinolone susceptibility and in the evolution of quinolone resistance.
16
Regulation and induction of the SOS response
The SOS response is regulated by two key proteins, the LexA repressor and the activator; RecA. During
regular cell growth, the LexA repressor binds to a specific sequence in the promoter regions of the SOS
response genes called the SOS box. The binding of LexA to the SOS box blocks the expression of the SOS
response genes. In addition, the binding of LexA to the SOS box also regulates the sequence by which
the SOS response genes are expressed during DNA damage. Genes expressed early in the SOS response
have a low affinity SOS box, while the SOS box in the promoter region of late SOS genes has a high
affinity for LexA. When the DNA is damaged, filaments of activated RecA are assembled on persisting
regions of single stranded DNA. The assembly of the RecA filaments facilitates the autocleavage of the
LexA repressor, thereby leading to expression of the SOS response genes. In E. coli more than 40 genes
are regulated by LexA cleavage in response to DNA damage, including genes responsible for DNA repair
and cell cycle control (73-75).
Repair of quinolone mediated double stranded DNA breaks by the SOS response
One of the major tasks carried out by the SOS response genes
is DNA damage repair. The DNA repair systems that are part
of the SOS response can repair a number of different types of
DNA damage. Repair of double stranded breaks (DSB) in
bacteria, like those caused by quinolones, is achieved by
homologous recombination (HR). In E. coli there are two
known pathways of HR , the RecBCD- and RecF-pathway,
where RecBCD is the predominate one (see Figure 5) (76).
RecBCD is a multi-functional enzyme complex, having both
nuclease and helicase activity, and is responsible for
processing the open DNA ends formed at DSBs in the DNA.
RecBCD initiates the DSB repair by binding to the open DNA-
end at the DSB and starts unwinding the DNA. Hereafter, a
combination of helicase and nuclease activity leads to
formation of a single stranded 3´-overhang. When the
RecBCD complex have reached a so-called chi site on the
strand with the open 3´-end, it loads RecA onto the 3`-tail,
creating a RecA filament, and dissociates from the DNA (77).
RecA then catalyzes strand invasion of a homologous dsDNA,
Figure 5: Schematic of DNA double stranded
break repair by homologous recombination
via the RecBCD pathway. Modified from
Wyman et al. 2004
17
creating a displacement loop (D-loop). Hereafter the intact homologous DNA strands are used as
templates for the DNA polymerase. DNA crosses termed Holiday junctions now physically link the
hetero duplex DNA strands. To resolve the Holiday junctions, the RuvAB protein complex extends the
heteroduplex DNA region by migrating the Holiday junctions in an outward direction. Following the
hetero-duplex extension, the RuvC protein associated with RuvAB, resolves the Holiday junctions by
nicking the crossed strands. A DNA ligase then ligates the nicks in the DNA, reconstituting the two
double strands (78).
Quinolone resistance
Quinolones have become one of the most prescribed antibacterial drugs in the world today (14-16), It is
therefore not surprising that quinolone resistance has been identified in almost all bacterial species of
clinical interest (17, 18). A number of different resistance mechanisms conferring quinolone resistance
have been identified this far, including; target site mutations, enzymatic inactivation, target protection
and efflux systems.
Target site mutations
Quinolone resistance is most frequently caused by target site mutations in the gyrase and topo IV,
eventhough the mutations confering quinolone resistance have been mapped to wide range of positions
in both subunits of the gyrase and topo IV. The most frequent mutations are found at the serine and
acidic residues of gyrA and parC that are critical for binding of quinolones via the water-metal-ion bridge
(21). Studies have revealed that more than 90% of all clinical isolates and laboratory derived strains with
lowered susceptibility to quinolones generally have a mutation at the specific serine residue, and that
85% of these also have parC mutations (79). In vitro selection of quinolone resistant mutants have
shown that the mutations in the gyrase and topo IV are selected for in a stepwise manner (34, 80). In
most cases multiple mutations are needed to confer clinical quinolone resistance. In an E. coli
background without any other quinolone confering mechanisms, two amino acid substitutions in gyrA
and one in parC is needed for the ciprofloxacin MIC to exceed the CLSI clinical breakpoint of 1 µg/mL
(81, 82).
Often, target site mutations confering antibiotic resistance have a negative impact on
the fitness of the bacteria (83, 84). It has therefore, to some degree, been surprising that quinolone
resistance caused by target site mutations, has become such a serious problem in the clinic. An
explanation to this paradox is likely that third-step quinolone resistance mutations have been shown to
both restore fitness and increase resistance significantly (80, 81). The increase in fitness could thereby
catalyze the selection of mutants highly resistant to quinolones, without exposure to high quinolone
18
concentrations (81). Furthermore, there is evidence that accumulation of quinolone resistance
mutations lead to increased mutation rates (85). Taken together with the fact that quinolones, as
mentioned earlier, are induceres of the SOS response and thereby mediates expression of the error-
prone DNA polymerase IV (86). The frequent occurrence of quinolone resistance might not be so
suprising after all.
Non-target site mutations involved in quinolone resistance
A number of different non-target site mutations often occur in quinolone resistant bacteria. Including,
deleterious mutations in acrR and marR, a direct and an indirect repressor of the expression of the
endogenous AcrAB-TolC efflux system in E. coli (87), thus leading to elevated levels of quinolone efflux.
MarR acts by repressing expression of marA, a global transcriptional activator, that activates expression
of acrAB (88). In addition, MarA also activates transcription of micF, encoding an antisense RNA that
post-transcriptionally inhibits the outer membrane porin, OmpF (89). OmpF is important for quinolone
entry into the cell in E. coli and its inhibition leads to lowered quinolone susceptibility (90, 91). Other
Gram-negative bacteria have similar efflux systems. For instance, Pseudomonas aeruginosa expresses
the MexAB-OprM efflux system that is repressed by MexR (92). Quinolone resistant clinical isolates of P.
aeruginosa often have mutations in mexR leading to overexpression of the MexAB-OprM efflux pump
(93, 94). The Gram-positive bacteria where quinolone efflux is best characterized is S. aureus. Here
overexpression of three different efflux pumps; NorA, NorB and NorC, have been shown to lower the
quinolone susceptibility. The regulation of these three efflux pumps is somewhat more complex than for
AcrAB-TolC and MexAB-OprM, as some transcriptional regulators, like GntR, is both an activator of norA
and norB, but a repressor of norC (95). A number of other efflux systems in both Gram-negative and
Gram-positive bacteria have been linked to quinolone resistance (95), but these will not be discussed
here.
Plasmid mediated quinolone resistance
Eventhough target site mutations are the most frequent cause of quinolone resistance, a number of
different plasmid mediated resistance mechanisms have also been identified in clincal isolates. These
mechanisms do generally not confer clinical quinolone resistance, but have been shown to facilitate
selection of high level quinolone resistance (96-98). The first claims of a plasmid mediated quinolone
resistance (PMQR) mechanism were reported back in 1987 (99), but was later withdrawn. Though, it
was first over 10 years later, by Martinez-Martines et al., that the existence of PMQR was confirmed by
transfer of a plasmid that lowered the susceptibility for nalidixic acid and ciprofloxacin in an otherwise
susceptible E. coli strain (96).
19
The qnr genes; a DNA mimic.
The gene identified by Martinez-Martinez et al. was named qnr (later qnrA) (96), encoding an 218 amino
acid long protein, belonging to the pentapeptide repeat protein (PRP) family. The PRP family contains
more than a 1000 proteins, many of which are of unknown function (100). The PRPs are defined by
being composed of or having domains of tandem peptide repeats with the consensus sequence;
[S,T,A,V], [D,N], [L,F], [S,T,R] and [G] (101). It was the function of two other members of the PRP familly,
MfpA and McbG, that led the way to the discovery of the function of QnrA. MfpA and McbG are both
encoded on the chromosome and protect the DNA gyrase from ciprofloxacin and the natural DNA
gyrase poison microcin B17, respectively (98). Knowing this, the in vitro supercoiling activity of DNA
gyrase in the presence of ciprofloxacin and purified QnrA was assesed. The results reveald that QnrA
protected the DNA gyrase from inhibition by ciprofloxacin, retaining its ability to supercoil DNA (102).
The discovery of qnrA was followed by the discovery of six other families of plasmid born
qnr genes; qnrS (103), qnrB (104), qnrC (105), qnrD (106), qnrE (107) and qnrVC (108). These six qnr
families generally have around 65%, or less, sequence homology with qnrA and each other(100). Crystal
structures of QnrB1 and a Qnr protein from the Gram-negative bacteria Aeromonas hydrophila showed
that they are dimers linked at the C-termini, folding into a right-handed β-helix (see Figure 6). This
strutucture resembles the size, shape and charge of β-DNA, which has led to the current opinion; that
Qnr proteins are DNA mimics that bind to and destabilize quinolone bund cleavage-complexes, leading
to release of the bund quinolone and reactivation of the topoisomerase (100, 109, 110). It still remains
to be resolved, how Qnr proteins can compete with DNA for binding to the DNA gyrase without
significantly inhibiting the gyrase actitivty in the bacteria.
Figure 6: Structure of the QnrB1 dimer. The two QnrB1 monomers are linked at the C-termini and fold into a right-handed
quadrilateral β-helix, mimicking the size, structure and charge of β-DNA. Deletion of loop A´or B´ leads to lowered
protection of DNA gyrase from ciprofloxacin. Modified from Vetting et al., 2011.
20
Inactivation by AAC(6´)-lb-cr mediated acetylation
The AAC(6´)-lb protein family consists of 6´-N-
acetyltransferases that can inactivate a number of
aminoglycoside antibiotics by acetylation (98). It was therefore
surprising when disruption of an aac(6´)-lb gene, on a multiple
resistance plasmid from a clinical isolate of E. coli, led to
increased ciprofloxacin susceptibility (111). An acetylation
assay showed that this novel member of the AAC(6´)-lb family
was able to N-acetylate ciprofloxacin at the amino nitrogen on
its piperazinyl substituent (see Figure 7). The enzyme was
therefore called AAC(6´)-Ib-cr, where “cr” stands for
ciprofloxacin resistance (111). AAC(6)-Ib-cr also confers
resistance to norfloxacin, but not other quinolones as they lack
the unsubstituted amino nitrogen group (111). As with Qnr,
AAC(6)-Ib-cr does not, by it self, cause clinical quinolone
resistance. It increases the MIC by three- to four-fold in wild
type E. coli, but more interestingly it increases the mutation
prevention concentration significantly. Thus, it likely plays an
important role in selecetion of higher level resistance
mutations (98, 111). In addition to the seven different allelic AAC(6´)-Ib-cr variants that have been
identified this far. An 24 amino acid longer variant, termed AAC(6´)-Ib-cr4, was discovered in a clinical
isolate of Salmonella typhimurium (112).
QepA and OqxAB efflux pumps
QepA and OqxAB are the major types of efflux pumpes that are involved in PMQR. QepA is part of the
14-transmembrane-segment family of the major facilitator superfamily transporters and was discovered
in an E. coli isolate from Japan with lowered susceptibility to quinolones (113). QepA is able to actively
pump out hydrophilic fluoroquinolones, especially norfloxacin and ciprofloxacin. The increase in MIC
conferred by QepA varies from 2-64 fold, this wide range is most likely caused by differences in QepA
expression (112).
The OqxAB efflux system is a member of the resistance-nodulation-cell division family of
transporters and is able to pump out a range of different antibiotics, including; quinolones,
chloramphenicol and trimethroprim. OqxAB is highly associated with extended spectrum beta lactamase
(ESBL) producing Klebsiella pneumoniae, where it is found on the chromosome and on plasmids. Like
Figure 7: AAC(6´)-Ib-cr acetylation of the
amino nitrogen of the piperazinyl
substituent in ciprofloxacin.
21
QepA, the expression level of OqxAB varies widely, hence the change in MIC differs from strain to strain
(112).
Reversing antibiotic resistance by helper drugs
In an effort to overcome the current crisis with treating infections caused by multi-drug resistant
bacteria, many different treatment types have been investigated. One of them is the reversal of
antibiotic resistance by combining antibiotic treatment with administration of a potentiating compound
also known as a helper drug. A helper drug is a compound that does not have an antibiotic effect in
itself, but is able to reverse the antibiotic resistance against a given antibiotic. In general helper drugs
can act by either directly targeting resistance mechanisms or by targeting intrinsic mechanisms
protecting the bacteria from the antibiotic, like; efflux pumps, cell membranes and repair systems. An
example of a helper drug that have been used with great success in the clinic is the combination of
clavulanic acid and the beta-lactam, amoxicillin. Clavulanic acid is a Beta-lactamase inhibitor (114), that
reverses resistance by competitively binding to beta-lactamases (115), thereby blocking the inactivation
of amoxicillin by the beta-lactamase.
Another type of helper drugs that have been heavily investigated are efflux pump
inhibitors (EPI), as many multi drug resistant pathogens have acquired mutations that elevates the
expression of their endogenous efflux pump systems (116). Inhibitors of the resistance nodulation
family (RND) of efflux pumps in Gram negative bacteria are especially interesting, as this family of efflux
pumps is able to pump out a wide variety of antibiotics, including; fluoroquinolones (ciprofloxacin and
levofloxacin), β-lactams, tetracyclines and oxazolidinones. Several compounds that inhibits the RND
family, including AcrAB-TolC and MexAB-OprM, are described in the literature, but so far, none of these
compounds have been licensed for medical use (117).
In addition to the AcrAB-TolC and MexAB-OprM inhibitors mentioned above, celecoxib, a
non-steroidal anti-inflammatory drug, has been shown to increase the susceptibility to ciprofloxacin in S.
aureus. In silico screening of a small library of celecoxib analogues identified a compound that inhibited
the NorA efflux pump and in vitro lowered the MIC for ciprofloxacin in a S. aureus strain overexpressing
NorA(118). Furthermore, multiple compounds have been proposed to inhibit RecA in vitro and thereby
prevent repair of quinolone mediated DSBs by HR and activation of the SOS response (119-125).
However, only suramin and copper phtalocyanine-3, 4´, 4´´,4´´´-tetrasulfonic acid were shown to
potentiate ciprofloxacin in vivo, albeit only weakly(119, 125).
22
Potential targets for potentiation of quinolones
A higher number of potential targets for quinolone helper drugs have been identified through genetic
screens. Specifically, screening the entire Keio collection (126) of close to 4000 single non-essential gene
deletion mutants of E. coli, revealed in excess of 25 genes, which when deleted significantly increased
the susceptibility to ciprofloxacin. Unsurprisingly, genes involved in DNA replication, recombination and
DNA repair counted for almost half of the total number of genes identified, though genes with a broad
variety of other cellular functions were also represented (127, 128). These findings were obtained in an
E. coli wild type strain, susceptible to ciprofloxacin. Paper I of this thesis addresses the question if
deletion of any of the identified genes renders high- and low-level ciprofloxacin resistant E. coli strains
clinically susceptible to ciprofloxacin, in an attempt to identify targets for ciprofloxacin helper drugs.
During the preparation of the manuscript for paper I, it was reported that single gene deletion of more
than 24 genes, identified as being involved in ciprofloxacin resistance, did not lower the MIC of a high-
level ciprofloxacin resistant E. coli strain beneath the clinical break point. Conversely, deletion of acrB in
combination with any of the SOS response genes recB, recC, recG or uvrD decreased the MIC of the
same E. coli strain beneath the clinical breakpoint (129). Furthermore, deletion of recA was reported to
render a low-level ciprofloxacin resistant strain clinically susceptible and to increase the in vivo efficacy
of ciprofloxacin against the same strain in a peritoneal sepsis murine model (130).
Part II: Targeting the initiation of chromosomal DNA replication in bacteria
Potentiation of existing antibiotics is one of many methods that have been deployed to overcome
antibiotic resistance. Another approach is the discovery of novel drugs that target unexploited processes
essential for bacterial growth and viability. A target that is underexploited is the chromosomal DNA
replication, and more specifically its initiation (131). Currently, the only antibiotics that targets the DNA
replication and are used in the clinic are the quinolones and novobiocin (132). The DNA replication is an
attractive target for novel antibiotics for numerous reasons. The proteins that are involved in DNA
replication are conserved in prokaryotes, but differ greatly with respect to their eukaryotic
counterparts. Furthermore, the number of replisomes per cell is low; hence, the quantity of a given
target that needs to be inhibited to block replication is correspondingly low (132). The following sections
will introduce the reader to the replication initiation process and how it is regulated in E. coli, where it is
best characterized.
23
Initiation of chromosomal DNA replication in E. coli
DNA replication and the cell cycle.
Early studies of the DNA content in bacteria growing at different rates showed that fast growing bacteria
contained more DNA per cell, relative to slower growing ones (133). A logic explanation to this fact
would be that replication forks move more rapidly during fast-growth. However, studies of DNA
replication and cell cycle in balanced cultures of E. coli demonstrated that the average rate of DNA
elongation is constant in cells with doubling times between 20 and 100 minutes (134). Furthermore, it
was demonstrated that the duplication of the chromosome took approximately 40 minutes (the D
period), independent of the growth rate, and that an additional 20 minutes was needed to complete
septum formation and cell division (the C period). However, the so-called I-period, which defines the
time it takes to prepare initiation of a round of chromosome replication is strictly dependent on the
growth rate (135). Interestingly, the sum of C + D being equal to 60 minutes means that in cells with a
doubling time below 60 minutes, replication and division is not completed before a new round of DNA
replication is initiated. As replication forks in E. coli move bidirectional from a single and fixed origin of
replication (136), newly divided cells, with a doubling time of less than 40 minutes, will therefore inherit
branched chromosomes with multiple origins and ongoing replication forks (see Figure 8). The fact that
fast growing cells contain branched chromosomes explained why they have a higher DNA content per
cell relative to slower growing ones. In addition, it also revealed that timeous replication initiation was a
key factor in the bacterial cell cycle (137, 138).
24
Initiation of replication
In short, the chromosomal DNA replication in E. coli is initiated by binding of the initiator protein DnaA,
in its active ATP-bound form, to the origin of replication, oriC. The formation of oriC-DnaAaTP
nucleoprotein complex triggers the separation of the DNA double strand (139, 140). Following opening
of the DNA double strand the nucleoprotein complex loads the DnaB helicase, with help from the
helicase loader protein DnaC (141, 142). Loading of DnaB triggers the assembly of the remaining parts of
the replication machinery necessary for DNA synthesis(143).
Multiple regulatory systems have been identified, ensuring that replication initiation is
triggered in a timely manner and only once per cell cycle for each origin. Following replication initiation,
regulatory inactivation of DnaA (RIDA) stimulates the autohydrolysis of DnaA bound ATP to ADP,
increasing the level of the inactive DnaAADP (144). A similar, but RIDA independent hydrolysis of ATP
bound to DnaA is mediated by a mechanism referred to as, datA-dependent DnaAATP-hydrolysis (DDAH)
(145). Furthermore, the negative initiation regulator protein SeqA sequesters hemi-methylated DNA in
Figure 8: Replication of the E. coli chromosome during moderate growth. On top, a cell with one chromosome and four
origins (green). On the right, the oldest replication forks terminates at the terminus (red) and the chromosomes are
segregated. Replication is then initiated from the four origins followed by cell division. On the left, the cells have divided and
now contain a single chromosome with four origins and four ongoing replication forks. Inspired by Fossum et al. 2007
25
the oriC, sterically hindering replication initiation by DnaAATP (146, 147). When time comes to reinitiate
DNA replication, DnaAADP is reactivated by oligomerization at two DnaA-activating sequences (DARS1
and DARS2), which triggers the release of ADP. The nucleotide free apo-DnaA is then activated by
binding of ATP and ready to initiate the DNA replication (148).
The origin of replication
In E. coli, the minimal oriC is a 245 bp DNA element containing two regions with distinct functionality;
one is the DnaA oligomerization region (DOR) and the other the DNA unwinding element (DUE) (see
Figure 9) (139). The DUE is defined by three 13-mer sequences (L, M and R) with the consensus
sequence 5´-GATCTnTTnTTTT-3´ (149). DnaAATP complexed at the DOR unwinds the DUE region, which is
susceptible to duplex unwinding due to its high AT-content. The DOR contains twelve DnaA binding
sites, known as DnaA boxes, with the 9-mer consensus sequence 5´ TTATnCACA-3` (139, 150). The DOR
can be divided into three sub-regions; the left- and right-halfs and a middle. The six DnaA boxes in the
left-half (R1, τ1, R5M, τ2, I1 and I2) all point in the same direction, i.e. they are situated on the same
strand. The remaining five DnaA boxes in the right-half (C3, C2, I3, C1 and R4) and the one in the middle
(R2) share directionality, but in thee opposite direction of the DnaA boxes in the left-half (151-153). The
R1, R2 and R4 DnaA boxes are moderate to high affinity DnaA boxes bound by either DnaAATP or DnaAADP
throughout most of the cell cycle, While the remaining DnaA boxes are low affinity boxes (154, 155).
The left-half DOR also contains a binding site (IBS) for the integration host factor (IHF) between DnaA
box R1 and τ1.
Figure 9: Structure of the minimal oriC in E. coli. The twelve DnaA boxes and their directionality are shown
by blue triangles. The IHF binding site is marked by a square between DnaA box R1 and τ1. Red arrows
mark the three 13-mer AT-rich sequences of the DUE. Katayama et al., 2017.
26
The initiator protein DnaA
The master replication initiator DnaA is a conserved 473 amino acids (aa) long protein composed of four
domains (156, 157) (see Figure 10). Domain I covers the first 87 aa in the N-terminal and is important for
protein-protein interactions (158). Specifically, mutational studies have shown that Trp-6 is essential for
the oligomerization of DnaA at the oriC by promoting domain I-domain I interactions between
neighboring DnaAATP molecules (159-161). Furthermore, substitution of either Glu-21 or Phe-46 with
alanine results in failure of DnaB helicase loading and for Phe-46 also binding of DiaA (161, 162), a
stimulator of DnaAATP assembly on the oriC and DUE unwinding (163). Finally, Asn44 has been shown in
vitro to be essential for RIDA, but not for initiation of replication (164).
Domain II of DnaA is the least conserved domain and varies significantly in both length
and sequence among bacterial species (157). It is usually described as flexible linker that connects
domain I and domain III. Systematic deletions studies showed that having either the 21 N-terminal
residues or the 27 C-terminal residues of the domain is sufficient for correct DnaA function, though
replication initiation in the deletion mutants was less efficient than in wild type cells (158).
Figure 10: Overview of the four domains of DnaA and its functions.
27
Domain III is the
largest domain of DnaA and
contains the AAA+ (ATPases
associated with diverse cellular
activities) region, making DnaA part
of the AAA+ superfamily of proteins.
The AAA+ module of DnaA can be
divided into two subdomains; a αβα-
nucleotide binding core and a
smaller C-terminal α-helical bundle,
known as the “lid”. The αβα-core is
composed of several signature motifs holding residues that are important for ADP/ATP binding and ATP
hydrolysis, while the sensor 2 motif is found in the “lid” (See Figure 11). The Walker A element forms a
loop structure, important for ATP/ADP binding, while residues of the Walker B motif interacts with the
magnesium ion that is crucial for ATPase activity (165). Lys-178 is an essential residue in the walker A
element that is highly acetylated in stationary growth phase cells, preventing binding of DnaA to ATP
and has therefore recently been proposed as novel regulatory mechanism of replication initiation (166).
Asp-269 and Arg-334 of the sensor 1 and 2 motifs, respectively, are required for high affinity ADP/ATP
binding (167, 168). Furthermore, Arg-334 is essential in DnaA ATP auto hydrolysis by both RIDA and
DDAH, most likely due to direct interactions with the ϒ-phosphate of the bound ATP (169, 170). Box IV
contains an arginine finger (Arg-285) that is exposed upon binding of ATP to DnaA. It is believed that the
exposed Arg-285 is able to interact with the ATP in the neighboring DnaAATP molecule in the DnaAATP-
oriC- complex, thereby facilitating the assembly of an active initiation complex (171). This kind of
assembly is shared between all AAA+ oligomers that have been structurally characterized (165). In
addition to Arg-285, four other residues, Lys-243, Arg-227, Arg-281 and Leu-290, are required for
Domain III-Domain III interactions(139) and contribute to DnaA oligomerization and DUE unwinding
(172-174). It as has recently been shown that Lys-243 can be acetylated in vivo, which blocks binding to
the low affinity DnaA boxes I3, C1 and C3 in vitro, though its significance for replication initiation is still
unsure (175). During DUE unwinding Val-211 and Arg-245 are believed to bind ssDUE, as in vitro assays
have shown that alanine substitution mutants of any of the two residues leads to deficiency in both DUE
unwinding and ssDUE binding.
Domain IV in the C-terminal of DnaA contains a typical helix-turn-helix motif (HTH), which
binds specifically to the DnaA box 9-mer consensus sequence. Crystallography studies of domain IV
complexed with the R1 DnaA box revealed that the binding leads to a 20O degree bend in the DNA. A α-
Figure 11: The ATPase module of DnaA from Aquifex aeolicus bound by the
ATP analog β,ϒ-methylene-ATP. Modified from Snider et al. 2008.
28
helix in the HTH-motif, constituted by residue 434-451, is inserted into the major grove of the DnaA box,
recognizing the 5’-TnCACA-3’ part of the consensus sequence (176). In addition, several residues of
domain IV interacts with the phosphate backbone of the DnaA box, specifically mutations in Arg-407
and Lys-417 leads to DNA binding deficiencies (176, 177). A single residue of the HTH-motif, Arg-399,
mediates base pair recognition by domain IV in the minor groove of the DnaA box (176). The importance
of Arg-399 is emphasized by the fact that mutations in this specific residue leads to loss of sequence
recognition and DNA binding (177). As for the major grove, multiple residues of the HTH-motif also
interacts with the phosphate backbone of the minor groove (176). Molecular dynamic simulations and
crystallography studies have shown that a short flexible loop connecting domain IV with domain III,
allows for pivoting of domain IV, and indicated that this is an important feature in DnaA oligomerization
(178, 179). Besides its essential function in DNA-binding, two residues of domain IV, Leu-422 and Pro-
423, contributes to binding of Hda, which is essential for RIDA activity (180).
Replication initiation by DnaAATP
The hallmarks of the replication initiation process is binding and oligomerization of DnaAATP on the oriC,
DUE unwinding and DnaB helicase loading. Although the process of replication initiation has been
investigated for decades, the exact structural and dynamic events that leads to replication initiation still
remains to be fully elucidated, due to its complex nature.
Formation of the DnaAATP initiation complex
Studies of the orientation of DnaAATP molecules in complex with oriC have shown that structurally
distinct complexes are formed on the left-half, right-half and middle DOR. As described above the DnaA
boxes within each DOR are orientated in the same direction, hence the DnaAATP molecules bound to
each box are also orientated in the same direction and interact in a head to tail manner (152, 173, 181).
Truncation studies of the DOR regions revealed that the left-half DOR complexed with DnaAATP and
bound by IHF is capable of mediating DUE unwinding, independently of the right-half and middle DOR
(182). In the right half DOR, DnaAATP is believed to initially bind the high affinity box R4, which then
triggers sequential binding of CI, I3, C2 and C3 (152). A similar binding order has also been proposed for
the left-half DOR, where R1 binding is followed by binding at τ1, R5M, τ2, I1 and I2 (152). However,
recently DnaA assembly studies on the left-half DOR revealed that deletion of R1 did not have a
significant effect on DnaAATP assembly at the remaining DnaA boxes. Whereas, deletion of the low
affinity box R5M severely impaired complex formation. Indicating, that R5M acts as the core assembly
point in the left-half DOR (153). These findings fits well with the fact that sequential binding of DnaAATP
molecules is most effective if the distance between the binding sites is 2-5 bp (152), which is the case
29
for R5M with respect to τ1 and τ2, but not for R1 that is situated 33 bp from its nearest neighbor, τ1 (E.
coli, MG1655) (183). DnaAATP occupying the middle DOR DnaA box, R2, has been suggested to interact,
via domain I-domain I interactions with DnaAATP occupying the I2 box, thereby stabilizing and promoting
assembly of the initiation complex on the left-half DOR (153, 184).
DUE unwinding
Three different models have been proposed for the events leading to DUE unwinding (see Figure 12).
The first is known as the continuous filamentation model or the two-state model. In this model, DnaAATP
can take two forms; an extended dsDNA binding state and a closed ssDNA binding state. Initially,
DnaAATP in its extended state binds to the DOR and a continuous DnaAATP filament is branched into the
DUE, where a combination of ATP-dependent unwinding by DnaAATP and torsional stress starts to open
the DNA duplex. As the DUE is unwound the conformational state of the DnaAATP molecules in the DUE
shifts to the closed confirmation allowing them to bind and stabilize the ssDUE (185-187). The two other
models for DUE unwinding are variants of the so-called loop back model. In the first variant, DnaAATP
assembly in the right-half DOR starts at R4 and ends at C3. In the left-half DOR DnaAATP binds to R1,
followed by binding of IHF. The IHF induced bend in the DNA loops back the R1-DnaAATP complex to the
low affinity boxes in the left-half DOR and triggers the assembly of DnaAATP on the remaining left-half
DnaA boxes. A combination of DNA bending by IHF and/or interactions with the DnaAATP oligomer on the
left-half DOR unwinds the DUE, which is then bound by DnaAATP in its closed ssDNA binding state (188).
The second variant of the loop back model differs from the first variant in two key points; i) R5M is
proposed as the core assembly site of the DnaAATP oligomer in the left-half DOR, though the R1- DnaAATP
still interacts with the DnaAATP oligomer in the left-half DOR. ii) The ssDUE directly interacts with the
DnaAATP oligomer in the left-half DOR, through Val-211 and Arg-245 of domain III, known as the H/B
motifs (153, 179, 182).
Figure 12: Current models for DUE unwinding. Only DnaA domain III and IV are shown. Sakiyama et al., 2017.
30
DnaB helicase loading
The next step in the replication initiation process, following DUE unwinding, is loading of the DnaB
helicase. The functional DnaB helicase is a hexamer of identical DnaB monomers that forms a barrel
shaped toroid structure (189-191). Replicative helicases, like DnaB, are molecular motors driven by ATP
hydrolysis that are able to translocate along ssDNA and induce unwinding of duplex DNA in front of the
moving replication fork (192). Loading of the DnaB hexamer onto the ssDUE is chaperoned by the DnaC
helicase loader. Recent evidence suggests that the DnaB ring structure opens and closes and that
binding of three to six DnaC molecules traps it in its open conformation, ready for loading onto the
ssDUE (193). DnaB helicase loading has been proposed to happen independently for the left- and right-
half DOR-DnaAATP complex, creating two distinct DOR-DnaAATP-DnaB complexes (173, 179, 182). The
loading of the DnaB is mediated by interactions between DnaA domain I, including Glu-21 and Phe-46,
and DnaB (161, 162, 194). For the second variant of the loop back model, it has been suggested that a
DnaB-DnaC complex is initially loaded onto the ssDUE opposite of the DNA strand that interacts with the
left-half DnaAATP-DOR complex. The loaded helicase then moves forward in the direction of the right-half
DnaAATP-DOR complex, revealing a stretch of ssDNA available for DnaB loading, by the right-half DnaA-
DOR complex, in the opposite direction and on the opposing strand of the other DnaB helicase (182,
195). Following loading of the DnaB helicase onto the ssDUE the DnaC molecules dissociates from the
DnaB hexamer. The release of DnaC is suggested to be mediated by interactions between DnaB and the
DnaG primase, stimulating the ATPase function of DnaC (196).
Regulation of the replication initiation
As mentioned above, the replication initiation is regulated to happen only once from each origin during
a cell cycle. Even in rapidly dividing cells, where the oriC copy number per cell is higher than two,
replication initiation at sister origins is triggered simultaneously and only once per cell cycle (139).
Several regulatory systems are deployed during the cell cycle to ensure that replication initiation is
31
triggered in a timely manner (See Figure 13). These regulatory systems are described in the sections
below.
The dual role of DiaA in regulating replication initiation
DiaA is a 196 amino acid long protein that forms homo-tetramers in which each monomer holds a DnaA-
binding site (163, 197). Observations that DiaA mutants initiates DNA replication asynchronously and
that DiaA in vitro promotes replication of mini-chromosomes, led to the conclusion that DiaA is a DnaA
associated factor that ensures timely initiation of the DNA replication process (163). A combination of
mutational and crystallography studies revealed that the DiaA homo-tetramer can bind multiple DnaA
molecules at once, and thereby stimulate DnaA oligomerization and DUE unwinding (197). The
stimulatory effect of DiaA on replication initiation has been explained by a linker effect observed for
several DNA binding proteins. By themselves the DNA binding proteins has a moderate affinity for DNA,
but when they are linked, through a linker protein, their DNA affinity increases dramatically (198).
Hence, DiaA linkage of DnaA molecules is suggested to increases the affinity of DnaA for DnaA boxes.
Interestingly, both DiaA deletion and overproduction inhibits replication initiation in vivo, indicating that
DiaA both has a positive and a negative effect on the initiation process. As described earlier, Phe-46, of
DnaA domain I is both involved in binding of DiaA and the DnaB helicase, thus DiaA proposedly blocks
the loading of the DnaB helicase by hindering the interaction between DnaA domain I and DnaB (162).
Figure 13: An overview of the regulatory systems that ensures timely initiation of the DNA replication during the cell cycle.
Katayama et al., 2017.
32
DiaA is believed to dissociate from the oriC-DnaA complex during DUE unwinding or closely after.
However, the dissociation mechanism still needs to resolved (139).
Regulatory inactivation of DnaAATP (RIDA)
Following a successful round of replication initiation DnaAATP is converted to its inactive form DnaAADP.
As described earlier, RIDA and DDAH are the two regulatory processes that are responsible for this
conversion, though RIDA is the predominant one (199). In RIDA, the activity of the DnaA ATPase is
stimulated by the DnaA homologue, Hda, in complex with the DNA loaded β sliding clamp (DnaN) of the
DNA polymerase III holoenzyme (144, 200). Like DnaA, Hda is member of the AAA+ superfamily of
proteins and holds a AAA+ module in its C-terminal (200), while the N-terminal is responsible for
interactions with the β-clamp. The DnaA ATPase stimulatory effect of the DnaN-Hda complex is only
active when Hda is bound by ADP (201). Arg-153 constituting the Arg-finger of the Hda AAA+ module is
crucial for the function of RIDA (202). The binding of ADP to Hda likely triggers a conformational change
in the Arg-finger, enabling interaction and activation of the ATPase region in domain III of DnaAATP (201).
In addition, interactions between DnaA domain I and the C-terminal of Hda seems to stabilize the
contact and promote the conversion of DnaAATP to DnaAADP (164). Recently, a crystal structure of a β-
clamp-Hda complex from E. coli revealed insight into how the activation of RIDA might be regulated.
Interestingly, the β-clamp- Hda complex was shown to form an octamer, where two pairs of Hda dimers
were sandwiched by two β-clamp ring structures. Based on these findings, and additional biochemical
and genetic evidence, it was proposed that the octameric complex negatively regulates RIDA, by
encaging Hda. Additionally, it was suggested that loading the β-clamp with DNA, by the clamp loader,
leads to dissociation of the octamer and formation of a DNA-β-clamp-Hda complex that is active in RIDA
(203). The requirement for a DNA loaded β-clamp in activating RIDA neatly couples active DNA
elongation with inhibition of the replication initiation (204).
datA-dependent DnaAATP-hydrolysis (DDAH)
In 1996, Kitagawa et al. identified a novel high affinity DnaA binding region (later datA) at 94.7 min. on
the E. coli chromosome, relatively close to the oriC (84.6 min.) (205). Shortly after its discovery, it was
reported that deletion of the datA locus led to asynchronous initiation of replication and that a DnaA
titrating plasmid suppressed the mutant phenotype. It was therefore proposed that datA repressed
untimely initiation by titrating high amounts of DnaAATP following a round of replication initiation (145).
In addition, an IHF binding site within the datA locus was shown to be important for maintaining a
proper timing of replication initiation (206). It was first over a decade after the initial discovery of datA
that the true mechanism by which datA regulates the timing in replication initiation was revealed.
33
Through a series of experiments, it was shown
that datA in complex with IHF promotes
hydrolysis of ATP bound to DnaA, through
inter-DnaA interactions at the datA locus
(170).
DDAH functionality depends on
a minimal datA locus of 183 bp containing the
two high affinity DnaA boxes 2 and 3, the low
affinity DnaA box 7 and a single IHF binding
site (see Figure 14A) (145, 170, 206-208). In
similarity to the individual DOR regions in the
oriC, the essential DnaA boxes in datA are all
orientated in the same direction, suggesting
that DnaAATP bound at these sites interacts in a
head to tail manner (208). Due to the long
distance between DnaA box 2 and 3, DnaAATP
at these two sites cannot interact without
binding of IHF. The binding of IHF to the IBS, which is situated between DnaA box 2 and 3, bends the
DNA and brings DnaAATP bound to box 2 and 3 in close proximity, thereby enabling their interaction (see
Figure 14B ) (208). The interaction between DnaAATP at box 7, 2 and 3 is mediated by domain III AAA+
Arg-finger and is further stabilized by Arg-281 and Leu-290. Furthermore, negative supercoiling of the
DNA stabilizes DnaAATP- DnaAATP interactions and the binding of IHF (170, 208). It has been suggest that
activation of DnaAATP hydrolysis at datA is promoted by conformational changes to the nucleotide
binding pocket of the AAA+ module, induced by inter DnaAATP interactions via Arg-281 (208). Conversion
of DnaAATP to DnaAADP is thought to destabilize the domain III-doamin III interaction mediated via Leu-
290, leading to release of the DnaAADP molecule from datA and loading of a new DnaAATP molecule.
Current evidence supports two distinct models for the conversion of DnaAATP to DnaAADP at datA. In one
model, DnaAATP hydrolysis is only activated in DnaAATP bound to DnaA box 7. In the other, DnaAATP is
hydrolysed at both DnaA box 7 and 2 (208). However, further research is needed to determine which of
the two models that may be correct.
Regulation of DDAH activity
Binding of IHF to datA is essential for the timely activation of DDAH. Cell cycle analysis of IHF-datA
complex formation indicated that IHF dissociates from datA before replication initiation and temporarily
Figure 14: A) Schematic of the minimal datA locus needed for
DDAH activity (DnaA box 4 is not essential). B) IHF induced
bending of datA leads to interactions between DnaATP bound
to DnaA box 2 and 3, which lead to activation DDAH. From
Katayama et al., 2017.
34
binds to datA shortly after the DNA replication is initiated. This suggests that the activation of DDAH by
IHF binding is tightly regulated by specific cell cycle events (139). Inhibiting transcription by treatment
with rifampicin is suggested to hinder dissociation of IHF from datA. As IHF is abundant in cells growing
exponentially (209), the inhibitory effect of rifampicin treatment on IHF dissociation from datA,
indicates that transcription in general or transcription of a specific factor is needed for inhibition of IHF-
datA complex formation (139). This hypothesis is further backed by the fact that moving datA to a highly
transcribed region on the chromosome inhibits the activity of DDAH (210). Hence, the timely binding of
IHF to datA might be regulated by changes in transcription through datA from adjacent genes (139).
Due to datAs close proximity to the oriC on the chromosome, it isreplicated shortly after
replication initiation, leading to a temporary increase in copy number. The increase in datA copy number
is believed to be important for repression of untimely replication initiation (205), which is in agreement
with the observation that a four-fold increase in the datA copy number delays replication initiation (211).
In contrast, datA deletion or transversal of the datA locus to the terminus region allows for untimely
replication initiation (145, 210). As mentioned above DNA supercoiling of datA promotes the activity of
DDAH (170, 208). In addition, an increase in untimely replication is observed for a datA deletion mutant
grown in nutrient poor-medium, relative to rich-medium (145). Indicating that the nutritional state of the
cell might influence DDAH activity. This theory was further established by analysis of the chromosomal
conformation during amino acid starvation, where datA was shown to interact with the oriC (212).
SeqA, a negative regulator of the replication initiation
In addition to the conversion of DnaAATP to DnaAADP by RIDA and DDAH, replication initiation is also
negatively regulated by the SeqA protein, which sequesters hemi-methylated GATC sites DNA in the oriC
after DNA replication has been initiated. In E. coli the Dam adenine methylase (Dam) methylates GATC
sites in the DNA. In newly replicated DNA only the parental strand is fully methylated while the daughter
strand is unmethylated, referred to as hemi-methylated DNA. Based on the early findings that Dam
deficient cells could not be efficiently transformed with mini-chromosomes unless it was methylated
and that fully methylated plasmids were only replicated one round in dam- cells (213, 214). It became
evident that the hemi-methylated state of the DNA somehow was involved in the regulation of the DNA
replication (215). Interestingly, it was shown that hemi-methylated DNA could be replicated in vitro.
Indicating that in vivo replication of hemi-methylated DNA was inhibited by an unknown factor, rather
than the hemi-methylation itself (216, 217). The unknown factor was later identified as SeqA in screens
for dam- mutants that could be transformed with, and maintain, a fully methylated mini-chromosome
(146, 147).
35
SeqA sequestrates the oriC for approximately one-third of the cell cycle and ensures that
a new round of replication is not triggered untimely at the newly replicated origins (217). The ratio of
DnaAATP to DnaAADP reaches its maximum at initiation and gradually decreases due to RIDA and DDAH
activity. In cells with a doubling time of 30 minutes, it takes approximately ten minutes to decrease the
DnaAATP to DnaAADP ratio to a level that prevents replication initiation. Hence, SeqA sequestrates the oriC
for a period equal to the time needed by RIDA and DDAH to prevent initiation by decreasing the
DnaAATP/ DnaAADP ratio to a certain threshold value (218-220). The minimal oriC contains 11 GATC sites,
which is significantly more than the average random distribution of GATC sites on the rest of the
chromosome. In vitro the binding of SeqA to hemi-methylated GATC sites in the oriC blocks DnaAATP
binding to DnaA box R5M, I2 and I3 that are all three overlapping with GATC sites (221, 222).
Conversely, the sequestration of the oriC by SeqA does not interfere with binding of DnaAATP/ADP to the
high affinity DnaA boxes R1 and R4 and the moderate affinity DnaA box R2 (222), which is in agreement
with the observation that DnaA occupies these three sites throughout most of the cell cycle(154, 155).
Furthermore, evidence show that the period of hemi-methylation of the oriC is reduced when the
available amount of DnaA is decreased (223).Though, the molecular mechanism that links the
sequestration period with the DnaA concentration is still not known. However, direct interaction
between SeqA and DnaA has been proposed to stabilize the sequestration of the oriC, though such an
interaction remains to be proven (223). Alternatively, DnaA binding to DnaA boxes overlapping GATC
sites in newly replicated origins might protect from methylation by Dam. A DnaA-SeqA exchange at the
oriC is then suggested to be mediated by an increase in allosteric DnaA binding sites due to ongoing
replication (217). At high concentrations, SeqA binding to DNA inhibits the formation of negative
supercoils, this inhibitory effect has been proposed to counteract unwinding of the DUE by DnaAATP
(224). In addition to the sequestration of the oriC, SeqA also binds to GATC sites in the dnaA promoter
following its replication (215). This binding inhibits the transcription of dnaA and thereby contributes to
the accurate timing of the replication initiation (225, 226).
Rejuvenation of the cellular DnaAATP pool
When it is time for the cell to prepare a new round of replication initiation, the DnaAATP level is
increased by three mechanisms. One is de novo synthesis of DnaA, which then bind ATP readily available
in the cytosol. The second and third are distinct pathways that lead to dissociation of ADP from DnaAADP
and subsequent binding of ATP. This process is mediated by either phospholipids in the cell membrane
or a pair of specific chromosomal DNA elements known as DnaA-reactivating sequences, DARS1 and
DARS2 (139).
36
DARS1 and DARS2
As mentioned, the dissociation of
ADP from DnaAADP is promoted by
DARS1 and DARS2, subsequently
leading to regeneration of DnaAATP
and stimulation of replication
initiation (148, 227). Even though
DARS2 is more than four times the
length of DARS1, 455 bp versus
101 bp, both elements contain a
similar core region of three DnaA
boxes (I, II and III) that are bound
mainly by DnaAADP. The regulatory
region is the major factor that
differentiates the two DARS
elements. DARS1 has a small ≈50
bp regulatory region, in contrast to the ≈400 bp in DARS2 (139). The differences in the regulatory
regions leads to an important functional difference of DARS1 and DARS2. In vitro, DARS1 is able to
mediate the dissociation of ADP from DnaAADP without any additional factors, while DARS2 activity is
significantly stimulated by interaction of its regulatory region with IHF and Fis (see Figure 15AB) (148,
227). In vivo, both DARS regions promotes DnaAATP production and replication initiation, as the deletion
of either delays the commencement of the replication (148, 227). However, deletion of DARS1 effects
the timing of the replication initiation less than deletion of DARS2, indicating that DARS2 promotes the
production of DnaAATP to a higher degree than DARS1. In addition, increasing the copy number of DARS2
leads to a more severe over-initiation, than an increase in DARS1 (148, 227). The observed difference
between the activity of DARS1 and DARS2 is likely, due to a promoting effect of IHF and Fis on the
number of DnaAADP molecules that oligomerizes at DARS2, as shown by pull-down assays (227). Owing
to the difference in the activity of DARS1 and DARS2, it is believed that DARS2 is important for timing
the replication initiation, while DARS1 might act to maintain a basal level of DnaAATP in the cell (139).
In both DARS1 and DARS2, DnaA box I is orientated in the opposite direction of DnaA box
II and III. Therefore, DnaAADP at DnaA box I and II interacts in a head to head manner, in contrast to the
head to tail interactions observed at oriC and datA. Even though the DnaA box core region of both
DARS1 and DARS2 is arranged in a similar manner, the events leading to oligomerization and ADP
dissociation are likely not identical. DnaA mutant analysis demonstrated that a D269N mutant was
Figure 15: A) Schematic of the DARS2 region, In light blue DnaA box I-IV, in
green the IHF binding site (IBS) and in orange the Fis binding sites (FBSs). B) The
DnaAADP-IHF-Fis complex at DARS2. Modified from Katayama et al., 2017.
37
deficient in both DnaAADP oligomerization and ADP dissociation at DARS1, but not at DARS2 (148, 227).
This difference is likely caused by unknown functions of IHF and Fis at DARS2 (139) Conversely, the ADP
dissociation via DARS1 was unaffected by a R334A mutation, while DARS2-mediated ADP dissociation
was moderately impaired by this mutation (148, 227). Like for inter DnaAATP-DnaAATP interactions at the
oriC Leu-290 is essential for the the oligomerization of DnaAADP at DARS2 (227). It remains to be
investigated, if Leu-290 is essential for DnaAADP oligomerization at DARS1. In the current mechanistic
model for DARS2-mediated ADP dissociation from DnaAADP. A DnaAADP oligomer forms at the DnaA box
core region, while IHF and Fis binds to their respective sites in the regulatory region. The binding of IHF
bends the DNA, which promotes the interaction between Fis and the DnaAADP oligomer at the core
region. The resultant DnaAADP-IHF-Fis complex (see Figure 15B) induces conformational changes in
DnaAADP leading to dissociation of ADP. The apo-DnaA then dissociates from the DARS2 complex and
binds to free ATP in the cytosol (139).
The activation of DARS2-mediated rejuvenation of the cellular DnaAATP pool is regulated
in a cell cycle coordinated manner by binding of IHF. IHF binds DARS2 in the pre-initiation period and
dissociates again just before the replication is initiated (227). Unlike IHF dissociation from datA, IHF
binding and release from DARS2 is resistant to inhibition of the transcription. Furthermore, the binding
of IHF is not coupled to replication initiation, as the binding and dissociation still occurs under conditions
where replication initiation is blocked. In light of these observations it is suggested that IHF-DARS2
interactions are regulated by an unknown cell-cycle dependent pathway that is uncoupled from the
regulation of the replication initiation (227). In contrast to IHF, Fis binds DARS2 throughout the cell cycle
and proposedly couples the replication initiation with the growth phase of the cell(227). As Fis is
abundant in cells growing exponentially, but scarce in stationary phase cells (209).
Phospholipid mediated reactivation of DnaAADP
The first evidence that phospholipids were involved in reactivation of DnaAADP, by mediating the release
of ADP, was published in 1988. Here it was shown that in vitro cardiolipin, an acidic phospholipid fund in
the E. coli cell membrane, interacted with DnaA and mediated the release of both ADP and ATP (228).
Subsequently it was demonstrated that mixtures of phospholipids and fluidic membranes also promoted
nucleotide dissociation from DnaA (229, 230). Furthermore, DnaA/oriC independent replication, known
as constitutive stable DNA replication (cSDR), suppressed the growth arrest observed in an E. coli strain
depleted for acidic phospholipids (231). In the same strain, it was demonstrated that the growth arrest
was also suppressed by expression of a DnaA L366K mutant (232). However, it remains unknown how
DnaA L366K suppresses the growth arrest. Flow cytometry analysis revealed a simultaneous shutdown
of the DNA replication and the protein synthesis in cells depleted of acidic phospholipids. Indicating,
38
that phospholipid mediated regulation of replication initiation might be part of a globular response
system (233). Nonetheless, more research is required to elucidate the mechanisms that lead to
regulation of replication initiation by phospholipids.
De novo synthesis of DnaA
The final known mechanism that is involved in increasing the DnaAATP level is de novo synthesis of DnaA.
The newly synthesized DnaA molecules bind to ATP that is abundant in the cytosol and are thereby
ready to partake in initiating the DNA replication. As described above, SeqA sequestration of the dnaA
promoter inhibits its transcription shortly after it has been replicated (215, 225). The sequestration of
the dnaA promoter by SeqA is proposedly auto-regulated by DnaA binding to DnaA boxes in the
promoter region, which stabilizes the sequestration (217). The dnaA promoter stays hemi-methylated
for approximately one sixth of the cell cycle following replication initiation; permitting initiation of RIDA
and DDAH activity and replication of DnaA titration sites on the chromosome (215, 234). Furthermore,
translocation of dnaA further away from the oriC leads to asynchronous replication initiation, explained
by an increase in the available amount of DnaA at the end of the oriC sequestration period. Hence, the
coordination between the periods of SeqA sequestration of both the oriC and the dnaA promoter is
crucial in timing the replication initiation (226).
The lethal action of severe over-initiation of the DNA replication
The importance of RIDA and DARS in regulating the replication initiation is emphasized by the severe
growth retardation and over-initiation observed in hda mutants, deficient in RIDA, and cells carrying
multiple copies of DARS2 (200). Evidence show that Hda deficient cells are viable under anaerobic
conditions or if the GO repair system is impaired (235). The GO system is involved in prevention and
repair of 8-oxo-dGTP incorporation in the DNA. Incorporation of 8-oxo-dGTP in the DNA is potentially
mutagenic due to its ability to form base pairs with both cytosine and adenine (236).The GO repair
system consists of at least three proteins, MutM, MutT and MutY. MutT acts by hydrolyzing 8-oxo-dGTP
to 8-oxo-dGMP, disabling its incorporation into the DNA. The excision of 8-oxo-dGTP already
incorporated into the DNA is mainly carried out by the formamidopyrimidine glycosylase, encoded by
mutM. Finally, the glycosylase activity of MutY enables it to remove adenines inserted opposite
incorporated 8-oxo-dGTPs (236). If 8-oxo-dGTPs are closely spaced in the DNA or encountered by
replication forks during repair, they may cause DSBs in the DNA (57). Based on the observations
described above and the fact that over-initiating cells have an increased number of ongoing replication
forks. It was suggested that the lethal effect of over-initiating replication is due to the formation of DSBs
when replication forks encounters 8-oxo-dGTP lesions that are under repair by the GO system (235).
39
Two other models have been suggested for how over-initiation leads to accumulation DSBs in the DNA.
In one model, it is proposed that ongoing replication forks collide with forks that have been stalled,
which leads to replication fork collapse and DSBs (237). In the second model, dNTP starvation, due to a
high number of replication forks, is suggested to lead to accumulation of DSBs in the DNA (238, 239).
In hda- mutants secondary mutations quickly arises. These mutations are known as hda
suppressor mutations (hsm), as they suppress the over-initiating phenotype of hda- cells. Several of the
identified hsm directly affects the replication initiation or the oriC itself, thereby diminishing the over-
initiation of the hda- cells (125, 240-242), while others permit growth despite of over-initiation (235,
243, 244). The later includes mutations in iscU and fre encoding an iron-sulfur cluster assembly protein
and the flavin reductase, respectively (244). Differential gene expression analysis of the iscU and fre
mutants by micro-array demonstrated a down regulation of genes involved in the TCA cycle and the
aerobic respiratory chain, while genes involved in the micro-aerobic respiratory chain were up-
regulated. Indicating a rerouting of the electron flow from the aerobic respiratory chain to the micro-
aerobic respiratory chain (243). The effect of such a rerouting is a decrease in the generation of ROS,
which in turn leads to a decrease in 8-oxo-dGTP formation and its repair. Hence, decreasing the ROS
production enables unhindered progression of replication forks in over-initiating cells (235, 243). In
paper II of this thesis, we further verify the proposed model for the lethal action of over-initiating the
DNA replication. As during a screen for replication initiation inhibitors, using over-initiating cells, we
identify the iron chelator deferoxamine, a known inhibitor of ROS production via Fenton chemistry (245,
246), as a compound that rescues the growth over-initiating cells by enabling fork progression during
hyper-replication.
Targeting the Initiation of replication
Multiple compounds have been identified that target different parts of the DNA replication machinery,
including, DNA ligase (247, 248), DNA polymerase III (249, 250), the β-sliding clamp (251, 252) and
single-stranded DNA-binding proteins (253). However, screening for putative inhibitors of the
replication initiation process have been limited and so far unsuccessful. A single screen for replication
initiation inhibitors has been published. This screen is based on a conditional lethal, cold sensitive DnaA
E. coli mutant that over-initiates replication. Thus, inhibition of replication initiation, at non-permissive
conditions, restores growth (254). Subjecting the screen to a library of pharmacological active
compounds (LOPAC), did not lead to the discovery of any replication initiation inhibitors. However, the
benzazepine derivative, (±)-6-Chloro-PB hydrobromide (S143), was identified as a novel gyrase inhibitor
that rescues the growth of over-initiating cells (255). Despite the current lack of success in identifying
compounds that blocks replication initiation, there is evidence that the initiation of chromosomal DNA
40
replication is a druggable process. As an inhibitor has been identified for the distinct replication
initiation process of the second chromosome in the Vibrionaceae family of bacteria (256). In addition,
expression of a cyclic DnaA domain I or over-expression of DnaA domain IV and I lead to inhibition of the
replication initiation, most likely by interfering with DnaA oligomerization at the oriC (252, 257). In
paper II and III of this thesis, we present two distinct strategies for identifying replication initiation
inhibitors.
41
Paper I: Can Ciprofloxacin Resistance be Reversed by
Helper Drugs?
Currently in review at: Annals of Clinical Microbiology and Antimicrobials.
42
Can Ciprofloxacin Resistance be Reversed by Helper 1
Drugs? 2
Rasmus N. Klitgaard 1, Bimal Jana2, Luca Guardabassi2, Karen Leth Nielsen3 and Anders Løbner-3
Olesen 1,* 4
1 Department of Biology, Section for Functional Genomics, University of Copenhagen, Copenhagen, 5
Denmark. 6
2 Department of Veterinary and Animal Sciences, Section for Veterinary Clinical Microbiology, University of 7
Copenhagen, Denmark. 8
3 Department of Clinical Microbiology, Center for Diagnostics, Rigshospitalet, Copenhagen, Denmark. 9
* [email protected]; Tel: +4535322068 10
Academic Editor: name 11
Received: date; Accepted: date; Published: date 12
Abstract 13
Background 14
Fluoroquinolones such as ciprofloxacin are potent antibacterial drugs that are widely used in the 15
clinic. As a consequence of their extensive use, resistance has emerged in almost all clinically 16
relevant bacterial species. In an attempt to reverse ciprofloxacin resistance, we searched for potential 17
helper drug targets in Escherichia coli strains with different levels and mechanisms of ciprofloxacin 18
resistance. 19
Methods 20
The search for ciprofloxacin helper drug targets was conducted by a combined transcriptomic and 21
genetic approach. Differential gene expression (DGE) analysis of the high-level ciprofloxacin 22
resistant E. coli sequence type (ST) 131 UR40 strain, treated with 2 µg/ml ciprofloxacin, was done by 23
43
RNA-Seq. In the genetic screen 23 single gene deletions were transduced from the Keio collection 24
into a high ciprofloxacin resistant E. coli strain LM693 (carrying gyrAS83L, gyrAD87N and parCS80I 25
mutations), followed by determination of the minimal inhibitory concentration (MIC) by Etest. The 26
seven individual gene deletions that lowered the ciprofloxacin MIC the most were subsequently 27
introduced into strains LM862/pRNK1 and LM862/pRNK9 carrying the aac(6´)-Ib-cr and qnrS genes, 28
which confer low-level ciprofloxacin resistance by drug modification and target protection, 29
respectively. The ciprofloxacin MICs were then determined for these two strains by broth micro-30
dilution. 31
Results 32
Differential gene expression analysis of ST131 UR40 treated with ciprofloxacin, showed that the 33
transcriptome was similar to that of untreated samples, i.e. no genes were found to be significantly 34
upregulated. The genetic screen of the 23 single gene deletions in LM693 identified a number of 35
genes that significantly lowered the ciprofloxacin MIC, including genes encoding the AcrAB-TolC 36
efflux pump, SOS-response genes and the global regulator fis. However, none of the deletions 37
lowered the MIC beneath the clinical breakpoint. In the low-level resistant strains carrying aac(6´)-38
Ib-cr and qnrS, respectively, deletion of acrA, tolC, recC or recA all rendered the strains clinically 39
susceptible to ciprofloxacin. 40
Conclusions 41
The results of the combined transcriptomic and genetic approach show that it is not straightforward 42
to reverse ciprofloxacin resistance in high-level ciprofloxacin resistant E. coli strains. On the other 43
hand, components of AcrAB-TolC efflux pump and the SOS response proteins, RecA and RecC were 44
identified as possible helper drug targets in E. coli strains with a MIC closer to the clinical 45
breakpoint. 46
Keywords: Antibiotic resistance, ciprofloxacin, helper drugs, RNA-Seq, transcriptomics. 47
44
48
Background 49
Fluoroquinolones are some of the most prescribed antibacterial drugs in the world [1-3], but this 50
has not always been the case. For the first two decades after the discovery of nalidixic acid in 1962, 51
and its introduction into the clinic in 1964, the quinolones were only used to treat uncomplicated 52
urinary tract infections. This changed with the release of the second generation quinolones, 53
including ciprofloxacin, which showed significant activity outside the urinary tract and against a 54
broad spectrum of both Gram-negative and Gram-positive bacteria. Ciprofloxacin acts by binding 55
to its targets, gyrase and topoisomerase IV, inhibiting the native ability of these two enzymes to re-56
ligate double stranded DNA breaks, in turn leading to fragmentation of the chromosome. Due to its 57
mechanism of action it is sometimes referred to as topoisomerase poison[4]. Inevitably, considering its 58
extensive use and misuse, resistance towards ciprofloxacin has arisen in almost all clinically 59
relevant bacteria [5, 6]. One method to overcome antibacterial resistance is by combinatorial 60
treatment with a potentiating compound, also known as a helper drug. A helper drug is by 61
definition non-antibacterial when administered alone, but it enhances the activity of the antibiotic 62
when used in concert. The potentiating effect of a helper drug can be achieved by either direct 63
inhibition of the resistance mechanism or by targeting endogenous cellular components and 64
pathways like, cell membranes, efflux pumps and cellular repair systems. A classic example of 65
targeting the resistance mechanism is the combination of amoxicillin and the β-lactamase inhibitor 66
clavulanic acid [7]. In Gram-negative bacteria, high-level ciprofloxacin resistance is mainly 67
associated with multiple target site mutations in gyrA and parC, encoding subunits of the DNA 68
gyrase and topoisomerase IV, respectively. Since 1998 three different plasmid-mediated 69
ciprofloxacin resistance mechanisms have been identified; i) target protection (Qnr proteins), ii) 70
efflux pumps (QepA and OqxAB) and iii) drug modification (AAC(6´)-Ib-cr acetyltransferase)[8]. 71
45
Here, we used a combined transcriptomic and genetic approach to identify potential helper drug 72
targets in Escherichia coli strains with different levels and mechanisms of ciprofloxacin resistance. 73
Methods 74
Bacterial strains and plasmids 75
Strains LM693 and LM862 were obtained from Diarmaid Hughes from Uppsala University. 76
LM693 is isogenic to MG1655 besides two gyrA mutations, S83L and D87N, and one parC mutation, 77
S80I. LM862 is also isogenic to MG1655, but with one gyrA S83L mutation and one parC S80I 78
mutation. ST131 UR40 has two gyrA mutations, S83L and D87, and two parC mutations, S80I and 79
E84V, and carries aac-6´-Ib-cr on a plasmid[9]. The aac-6´-Ib-cr carrying plasmid pRNK1 was 80
constructed as follows: aac-6´-Ib-cr gene was amplified by PCR from ST131 UR40, using the 81
following primers: GATCGGATCCATGAGCAACGCAAAAACAAAGTTAGGC and 82
CATCGAATTCTTAGGCATCACTGCGTGTTCGC, and cloned into pMW119 (Nippon Gene, 83
Toyama, Japan) using BamHI and EcoRI. The qnrS-carrying plasmid pRNK9 was constructed as 84
follows: qnrS was amplified by PCR from the clinical E. coli isolate EC38 using the following 85
primers: GATCGGATCCATGGAAACCTACAATCATACATATCGGC and 86
GATCAAGCTTTTAGTCAGGATAAACAACAATACCCAGTGC, and cloned into pMG25 using 87
BamHI and HindIII (M. Mikkelsen and K. Gerdes, unpublished). Strain EC38 was isolated from a 88
patient with a urinary tract infection at Hvidovre Hospital, Denmark. 89
Genetic screening and MIC tests 90
For the genetic screen, P1 phage lysates were prepared from the relevant Keio collection strains 91
[10] and used for transduction into LM693 and LM862. All the transduced strains were verified by 92
PCR. Theciprofloxacin MICs for LM693 and derived strains were determined using E-tests (0.002-32 93
µg/ml, BioMerieux) and according to the manufactures guidelines. The MICs for LM862 and derived 94
46
strains were determined by broth micro-dilution using cation adjusted Mueller Hinton broth II with 95
1mM and 10 µM IPTG for pRNK1 and pRNK9 respectively. The reference E. coli strain ATCC 25922 96
was used as standard in all MIC tests and the susceptibility was evaluated according to CLSI 97
breakpoints. 98
Checkerboard assay 99
All wells in a micro-titter plate were filled with 100 µl cation adjusted Mueller Hinton broth II (200 100
µL in the negative control wells). Copper phtalocyanine-3, 4´, 4´´,4´´´-tetrasulfonic acid, was added 101
to the first row, followed by serial dilution along the abscissa, leading to a start concentration of 100 102
µM. Hereafter ciprofloxacin was serial diluted along the ordinate, giving a start concentration of 2 103
µg/ml and 64 µg/ml for LM862 and LM693, respectively. 100 µl diluted culture with an OD600 of 104
0.001 was then inoculated in each well and the plates were incubated at 370C for 24 hours. 105
RNA-sequencing 106
ciprofloxacin was added to a balanced ST131 UR40 culture to a final concentration of 2 µg/ml. 107
Samples for RNA isolation were taken at 0 minutes (prior to ciprofloxacin addition) and 30 and 90 108
minutes after ciprofloxacin addition. Total RNA was isolated using a Thermo Scientific GeneJET 109
RNA isolation kit. Dnase treated with TURBO DNA-free kit from Ambion. rRNA was depleted using 110
an Illumina Ribo-zero rRNA removal kit, followed by RNA-Seq library prep using an Illumina 111
TruSeq Stranded mRNA Library Prep Kit. Sequencing was performed on an Illumina Miseq with a 112
Miseq reagent kit v3. (75bp paired-end) from Illumina. Data analysis was performed in Rockhopper 113
ver.2.03[11]. E. coli NA114 (ST131) (accession number: NC_017644) was used as reference genome[12]. 114
Results 115
Identification of helper drug targets by genetic screening 116
47
More than 25 single gene knockouts have already been shown to increase ciprofloxacin 117
susceptibility in wildtype E. coli strains [13-16]. Here, 23 of these deletions were introduced into the 118
high ciprofloxacin resistant strain LM693 [17] and tested for hyper-susceptibility towards 119
ciprofloxacin (Table 1). LM693 is isogenic to the commonly used laboratory strain MG1655 besides 120
two gyrA mutations; S83L and D87, and one parC mutation; S80I. Even though nine of the mutant 121
strains showed a three to four fold reduction in MIC , none of them were lowered beneath the CLSI 122
clinical breakpoint of 1 µg/ml. Our results therefore indicate that none of the tested gene-knockouts 123
identify valid helper drug targets in high-level ciprofloxacin resistant E. coli strains but could 124
potentially be used as helper drug targets to reverse low-level resistance. To create low-level 125
ciprofloxacin resistant strains, we constructed plasmids pRNK1 and pRNK9 carrying the 126
ciprofloxacin resistance determinants aac-6´-Ib-cr and qnrS, respectively. AAC-6´-Ib-cr inactivates 127
ciprofloxacin by N-acetylation of the amino nitrogen of its piperazinyl substituent [18], while QnrS 128
acts as a DNA mimic, binding to and protecting the gyrase from the action of ciprofloxacin[8]. 129
Introduction of pRNK1 and pRNK9 into strain LM862, which carries gyrA S83L and parC S80I 130
mutations, increased the MIC from 1 to 2 µg/ml, i.e. above the clinical breakpoint. We then evaluated 131
the ability of seven of the most promising of the 23 gene deletions described above to reduce 132
ciprofloxacin resistance. Four of the deletions (acrA, tolC, recA and recC) lowered the MIC beneath the 133
clinical break point. (Table 2). To assess whether inhibition of RecA was an amenable strategy for 134
potentiation of ciprofloxacin, synergy between ciprofloxacin and a RecA inhibitor, copper 135
phtalocyanine-3, 4´, 4´´,4´´´-tetrasulfonic acid[19], was tested by a checkerboard assay. No reductions 136
of the ciprofloxacin MICs were observed for either of the high- (LM693) -or low-level (LM862) 137
resistant strains. 138
48
139
Identification of helper drug targets by RNA sequencing 140
The E. coli clonal group sequence type (ST) 131 has become the predominant E. coli lineage 141
isolated from extra-intestinal infections and is currently regarded a global problem in hospitals and 142
clinical practices. Two independent studies have shown that more than 90% of ESBL-producing 143
ST131 isolates are also resistant to ciprofloxacin [20, 21]. Strain ST131 (UR40) is resistant to high levels 144
of ciprofloxacin due to gyrA mutations S83L and D87, and parC mutations S80I and E84V [9]. Here 145
we used RNA-Seq to map the transcriptomic changes during treatment of ST131 UR40 with a 146
clinically relevant concentration of ciprofloxacin (2 µg/ml). The rationale behind this was to identify 147
potential helper drug target genes that are over-expressed upon ciprofloxacin exposure and 148
putatively involved in ciprofloxacin resistance. In contrast to the genetic screen, the RNA-Seq analysis 149
would also reveal targets encoded by essential genes and non-coding RNA. The transcriptomic 150
analysis did not show any non-ribosomal transcripts to be significantly upregulated in the presence 151
of ciprofloxacin, i.e with a false discovery rate of <1% and more than 2-fold expression change. 152
Strain/Single deletions MIC (µg/ml)
LM693 24-32
tolC 1.5 acrA, acrB and fis 2
recC, xseA, xseB, uvrD and recA 4
ruvC and dksA 6 recG and hlpA 8
pgm, ybgF and ybgC 12 deoR, ydcS, yciT, ybjQ 16
ygcO and nlpC 24 rimK 24-32
Table 1. MIC values for the single gene
deletions in LM693.
MIC(µg/ml) Strain pRNK1 pRNK9
LM862 (No plasmid) 1 1 LM862/Empty vector 1 1
LM862 2 2 tolC 0.25 0.5 acrA 0.25 0.5 recA 0.5 0.5 recC 0.5 0.5 uvrD 2 1 xseA 1 1
fis 2 4
Table 2. MIC values for the single gene
deletions in LM862/pRNK1 and LM862/pRNK9
49
Discussion 153
By utilizing a combination of “direct genetic screening “and differential gene expression analysis, we 154
have attempted to identify genes suitable as targets for ciprofloxacin potentiating compounds. We 155
did not find any genes to be significantly up-regulated by ciprofloxacin, indicating that the 156
transcriptome of ST131 UR40 was fairly unaffected by treatment with a sub-inhibitory and yet 157
clinically relevant concentration of ciprofloxacin. The ciprofloxacin is most likely not binding to its 158
target, pumped out by efflux pumps or inactivated by Aac-6’-Ib-cr. The lack of an upregulation of the 159
SOS response genes in the transcriptomic analysis clearly shows that the ciprofloxacin exposure did 160
not cause sufficient DNA damage to induce a SOS response; hence it was not necessary for ST131 161
UR40 to up-regulate any specific genes to cope with the presence of ciprofloxacin at a sub-inhibitory 162
concentration. 163
The screening of selected mutant strains revealed a number of genes, which when deleted, 164
lowered the MIC for ciprofloxacin significantly. These findings are in accordance with genes reported 165
to contribute to high-level ciprofloxacin resistance by Tran et al. [22]. Treatment of bacteria with 166
ciprofloxacin generates double stranded breaks in the DNA of the bacteria [23], which in turn 167
activates the SOS response. Seven of the tested gene deletions; recA, recC, recG, uvrD, xseAB and ruvC, 168
which all significantly reduced the MIC of LM693, are part of the SOS response and involved in DNA 169
damage repair [24-26]. The results from the MIC analyses indicate that deletion of any of these seven 170
genes lowers the ability of the bacteria to cope with ciprofloxacin induced DNA damage. Deletion of 171
genes encoding the AcrAB-TolC efflux pump, or the global regulator Fis (Factor for inversion 172
stimulation) showed the largest decreases in MIC values relative to LM693. The Fis protein has been 173
50
shown to repress the gyrA and gyrB promoters, thereby reducing the expression of the DNA gyrase 174
[27]. Deletion of fis therefore increases DNA gyrase expression and the number of ciprofloxacin 175
targets. As ciprofloxacin works as a topoisomerase poison, an increase in ciprofloxacin bound DNA 176
gyrase could potentially lead to an increase in double stranded breaks, and this could explain the 177
decrease in MIC for the fis deletion strain. The fis deletion did not have the same effect in the low-178
resistant strains LM862/pRNK1 and LM862/pRNK9, which may be explained by the relatively higher 179
affinity of ciprofloxacin for its target in LM862, compared to that of LM693. Hence, the increase in 180
expression of the DNA gyrase might lead to an increase in ciprofloxacin-gyrase complexes, but if the 181
ciprofloxacin induced DNA damage already is at a level, where the DNA repair mechanisms cannot 182
keep up, the fis deletion does not have a dramatic effect on the MIC. 183
Individual deletions of acrA, acrB or tolC genes encoding the AcrAB-TolC efflux pump had a 184
large effect on the ciprofloxacin susceptibility of both LM693 and LM862 strains. This was not 185
surprising as overexpression of the AcrAB-TolC efflux system has been connected to ciprofloxacin 186
resistance numerous times [28]. The deletion of acrA or tolC in the LM862 strains lowered the MIC 187
beneath the clinical breakpoint indicating that AcrAB-TolC efflux system is a potential target for 188
ciprofloxacin potentiating compounds in low level resistant E. coli. A number of AcrAB-TolC 189
inhibitors have been identified [29-33], two of which have been shown to decrease the MIC of 190
ciprofloxacin in susceptible E. coli strains [29, 30], but none of them are used in clinical practice so far. 191
Inhibition of RecA and thereby of the SOS response has been proposed as a strategy to fight 192
antibiotic resistance numerous times [19, 34, 35]. Our finding; that deletion of RecA in low-level 193
resistant strains of E. coli lowers the MIC beneath the clinical break-point, is in accordance with recent 194
51
observations by Recacha et al.[36]. Combined, this indicates that RecA could be a potential 195
ciprofloxacin helper drug target. 196
Even though deletion of AcrAB-TolC or RecA rendered LM862/pRNK1 and LM862/pRNK9 197
clinically susceptible to ciprofloxacin, the respective MICs were only 2 to 4-folds lower than the 198
clinical break-point. It therefore seems reasonable to assume that a given inhibitor should completely 199
block the activity of either RecA or AcrAB-TolC in order for it to be an efficient helper drug. This 200
hypothesis is backed by the failure of lowering the ciprofloxacin MIC of LM862 and LM693 with the 201
relatively poor RecA inhibitor phtalocyanine-3, 4´, 4´´,4´´´-tetrasulfonic acid. Overall, it might 202
therefore prove difficult to reverse ciprofloxacin resistance by helper drugs targeting the proteins 203
encoded by the genes tested in this study. 204
Conclusions 205
The combined transcriptomic and genetic approach show that it may be difficult to reverse 206
ciprofloxacin resistance in high-level resistant E. coli strains. However, the components of the AcrAB-207
TolC efflux pump along with the SOS response proteins RecA and RecC were identified as putative 208
targets for reversing resistance in low-level ciprofloxacin resistant strains. The only described RecA 209
inhibitor working in vivo, phtalocyanine-3, 4´, 4´´,4´´´-tetrasulfonic acid, was found unable to reverse 210
resistance, suggesting that it did not inhibit RecA to a degree sufficient to re-sensitize cells to 211
ciprofloxacin. 212
Abbreviations 213
MIC: Minimal inhibitory concentration, ST: Sequence type. 214
Declarations 215
52
Ethics approval and consent to participate 216
Not applicable. 217
Consent for publication 218
Not applicable. 219
Availability of data and materials 220
The RNAseq datasets generated and analyzed during the current study are available in the Gene 221
expression Omnibus, https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE89507 222
Competing interests 223
The authors declare that they have no competing interests. 224
Funding 225
Study was funded with financial support from the University of Copenhagen Centre for Control of 226
Antibiotic Resistance (UC-Care) and by the Center for Bacterial Stress Response and Persistence 227
(BASP) funded by a grant from the Danish National Research Foundation (DNRF120). 228
Authors´ contributions 229
RNK carried out all experimental work, designed the study, analysed the data and prepared the 230
final manuscript. ALO supervised all aspects of the study and helped prepare the final manuscript. 231
BJ assisted and supervised the experimental part of the RNAseq. LG supervised and delivered the 232
ST131 UR40 strain. KLN performed genomic analyses and delivered the EC38 strain carrying the qnrS 233
gene. All authors read and approved the final manuscript 234
Acknowledgments 235
53
We acknowledge the financial support from the University of Copenhagen Centre for Control of 236
Antibiotic Resistance (UC-Care) and by the Center for Bacterial Stress Response and Persistence 237
(BASP) funded by a grant from the Danish National Research Foundation (DNRF120). 238
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354
56
Paper II: A strategy for finding DNA replication
inhibitors in E. coli identifies iron chelators as
molecules that promote survival of hyper-replicating
cells.
Currently in review at: Molecular Microbiology.
57
A strategy for finding DNA replication inhibitors in E. coli identifies iron chelators as molecules 1
that promote survival of hyper-replicating cells 2
3
Godefroid Charbon1†, Rasmus Nielsen Klitgaard1†, Charlotte Dahlmann Liboriussen1, Peter Waaben 4
Thulstrup2, Sonia Ilaria Maffioli3, Stefano Donadio3 and Anders Løbner-Olesen1* 5
6
From the 1University of Copenhagen, Dept. of Biology, Ole Maaløes Vej 5, 2200 Copenhagen N, 7
Denmark. 2University of Copenhagen, Dept. of Chemistry, Universitetsparken 5, 2100 Copenhagen 8
Ø, Denmark. 3NAICONS Srl, Viale Ortles 22/4, 20139 Milano, Italy. 9
10
Running title: Screens for DNA replication inhibitors 11
12
† Equally contributing authors. 13
14
*To whom correspondence should be addressed: Anders Løbner-Olesen: University of 15
Copenhagen, Dept. of Biology, Ole Maaløes Vej 5, 2200 Copenhagen N, Denmark. Phone: +45 16
3532 2068 . 17
18
Keywords: Escherichia coli, anti-replication initiation compounds, deferoxamine, iron chelation, 19
oxidative stress, DNA damage, DNA replication, drug screening. 20
58
21
22
Summary 23
DNA replication is often considered an attractive target for new antibacterial compounds. Here we 24
present a strategy to select molecules that inhibit initiation of chromosome replication. We made 25
use of two Escherichia coli strains that display hyper-initiation of replication by keeping the DnaA 26
initiator protein in its active ATP bound state. While viable under anaerobic growth or when grown 27
on poor media, theses strains become inviable when grown in rich media. Our strategy relies on the 28
ability of putative anti-replication initiation molecules to restore their survival. Extracts from 29
actinomyces strains were screened, leading to the identification of deferoxamine (DFO) as the 30
active compound in one of them. However, rather than inhibit replication initiation, we suggest that 31
DFO chelates cellular iron to limit the formation of reactive oxygen species and promote 32
processivity of DNA replication. We also argue that the benzazepine derivate (±)-6-Chloro-PB 33
hydrobromide acts in a similar manner. 34
Introduction 35
Duplication of the genetic material is essential for bacterial proliferation. Targeting DNA 36
replication for inhibition by new antimicrobials is attractive because the many factors contributing 37
to this process are conserved between prokaryotes, but differ significantly from their eukaryotic 38
counterparts (Robinson et al., 2012). Yet, only DNA topoisomerase inhibitors such as quinolones 39
are currently used in the clinic. Other molecules have been found to directly target components of 40
the DNA replication machinery as reviewed in (Robinson et al., 2012) but status for clinical use is 41
uncertain at this stage. 42
59
In Escherichia coli, like most bacteria, the commencement of DNA replication is controlled by 43
DnaA. DnaA is a conserved protein that binds to the chromosomal origin of replication, oriC, 44
promotes strand opening and loads the replication machinery (for recent reviews see (Leonard & 45
Grimwade, 2015, Riber et al., 2016, Skarstad & Katayama, 2013)). In E. coli, DnaA activity is 46
controlled by multiple regulatory pathways to ensure that it starts DNA replication only once per 47
cell cycle and at a defined cellular mass (Donachie, 1968, Cooper & Helmstetter, 1968). Deviations 48
from this once-and-only-once rule has fatal consequences for cell survival (Kellenberger-Gujer et 49
al., 1978, Hirota et al., 1970) . An increased frequency of initiations, such as provoked by hyper-50
activation of DnaA, leads to accumulation of strand breaks and cell death in a manner somewhat 51
resembling the mode of action of quinolones (Simmons et al., 2004, Charbon et al., 2014). 52
Inactivating DnaA on the other hand leads to an arrest in cell proliferation due to the absence of 53
duplication of the genetic material (Hirota et al., 1970). Slight deviations in the timing of initiation 54
that are seemingly inconsequent for bacterial growth in a laboratory setting affect competitiveness 55
in the host digestive tract (Frimodt-Moller et al., 2015). Thus compounds that affect DnaA function 56
and/or the replication initiation frequency holds promise for therapeutic use. 57
DnaA is composed of four domains performing distinct functions in the initiation process (Messer 58
et al., 1999), and domain I, III and IV functions could serve as putative targets for inhibition. 59
Domain I interacts with the DNA helicase to commence the assembly of the DNA replication 60
machine at the origin of replication and is involved in oligomerization of the protein. Domain II is a 61
flexible linker region that shows little conservation between DnaA proteins from different bacterial 62
species (Messer, 2002) . Domain III is an AAA+ ATPase domain which is often found in initiator 63
proteins. Domain III has a crucial function in promoting formation of a DNA bound DnaA polymer 64
necessary to induce DNA duplex opening and to interact with single stranded DNA (Erzberger et 65
al., 2006). Finally, binding of DnaA to oriC is ensured by a helix-turn-helix motif in Domain IV. 66
60
The regulation of DnaA is quite complex, but in essence, DnaA bound to ATP is the active form 67
that accumulates prior to initiation when a DnaAATP-oriC nucleoprotein complex is formed at the 68
origin. This complex triggers strand opening, helicase loading and assembly of the DNA replication 69
machinery to commence DNA replication. This multimeric DnaAATP assembly on oriC is regulated 70
by binding and hydrolysis of ATP in the AAA+ domain and is the key regulatory feature that 71
ensures proper timing of initiation (Sekimizu et al., 1987, Kurokawa et al., 1999). Following 72
initiation, and to prevent a new cycle of initiation, DnaAATP is inactivated, i.e. converted to 73
DnaAADP. This inactivation is triggered by regulatory inactivation of DnaA (RIDA) (Kato & 74
Katayama, 2001) and datA-dependent DnaA-ATP hydrolysis (DDAH) (Kasho & Katayama, 2013) 75
process. RIDA is performed by the Hda protein in complex with the -clamp loaded on the 76
chromosome. In this complex, Hda directly stimulates the ATPase activity of the DnaAATP 77
complex; DnaA now bound to ADP is inactive. Inactivation of DnaA by DDAH is achieved by the 78
formation of a DnaAATP nucleo-protein complex on the non-coding DNA element datA, which 79
stimulate ATP hydrolysis. Several factors stimulate the DnaA dependent initiation process without 80
being essential. These include DiaA, H-NS, IHF etc. (For review see (Riber et al., 2016, Skarstad & 81
Katayama, 2013)). 82
Prior to a new initiation event the pool of active DnaA molecules is increased by de novo synthesis 83
of DnaA and by rejuvenation of DnaAADP into DnaAATP. This rejuvenation is controlled by the 84
binding of DnaAADP to two DNA elements called DARS1 and DARS2 (Fujimitsu et al., 2009). 85
DnaAADP binding to DARS promotes the release of ADP which permits DnaA to rebind ATP and 86
be active for initiation. 87
Cells deficient in Hda and cells carrying a multi-copy DARS2 plasmid, have an increased 88
DnaAATP/DnaAADP ratio (Kato & Katayama, 2001, Fujimitsu et al., 2009) . This results in hyper-89
initiation of replication, also called over-initiation, and in most conditions loss of viability or 90
61
selection of compensatory mutations (Kato & Katayama, 2001, Fujimitsu et al., 2009, Charbon et 91
al., 2011, Riber et al., 2006). The high number of replication forks present in these cells makes 92
them hypersensitive to DNA damages such as those provoked by reactive oxygen species (ROS) 93
(Charbon et al., 2014, Simmons et al., 2004) (for review (Charbon et al., 2017b)). However, hyper-94
initiating cells are viable under growth conditions that reduce conflicts between the elevated 95
number of replication forks and DNA repair processes. These conditions include anaerobic growth 96
to lower oxidative damage to the DNA or slow growth to increase spacing between replication 97
forks. (Charbon et al., 2014, Charbon et al., 2017a). 98
Here we present a screen for inhibitors that target the initial step in the duplication of the bacterial 99
chromosome, i.e. replication initiation at oriC. The screen is based on shifting hyper-initiating cells 100
from permissive conditions to non-permissive conditions, the latter being aerobic growth on rich 101
medium. In principle, a compound that reduces initiations to a level that sustains growth can be 102
selected as it will provide viability to the cells. Such anti-replication initiation compounds are 103
expected to lower DNA replication and thereby viability in wild-type cells (Fig. 1A). 400 extracts 104
of filamentous actinomycetes were screened for containing putative replication initiation inhibitors, 105
a strategy that led to the discovery of -clamp targeting griselimycins antibiotics (Kling et al., 2015, 106
Terlain & Thomas, 1971). We identified deferoxamine (DFO) as being able to restore growth of 107
over-initiating cells. A detailed characterization of its mode of action however points to titration of 108
the cellular iron pool to reduce the Fenton reaction and thereby also ROS inflicted DNA damage. 109
Rather than being a replication inhibitor, DFO thus works by promoting replication elongation in 110
over-initiating cells. The benzazepine derivate (±)-6-Chloro-PB hydrobromide (S143) that was 111
previously identified in a similar screen (Johnsen et al., 2010, Fossum et al., 2008) and proposed to 112
target the DNA gyrase was found to act in a manner similar to DFO. 113
Results 114
62
Screening microbial extracts for inhibitors of the initiation of DNA replication 115
Cells deficient in Hda and cells carrying a pBR322 type plasmid with DARS2 have an increased 116
DnaAATP/DnaAADP ratio and hence over-initiate chromosomal replication, albeit to different extent, 117
with the degree of over-initiation being strongest in the presence of pBR322-DARS2 (Charbon et 118
al., 2014). Both cell types are viable during anaerobic growth or growth on a poor carbon source 119
such as glycerol (referred to as minimal poor medium) i.e. permissive conditions, while inviability 120
is observed during aerobic growth on rich medium, i.e. non-permissive conditions (Charbon et al., 121
2014). When over-initiating cells were plated on minimal medium plates supplemented with 122
glucose and casamino acids (referred to as minimal rich medium) and incubated aerobically no 123
growth was observed. In order to screen for inhibitors of DNA replication initiation, cells plated on 124
minimal rich medium agar plates were exposed to bioactive natural products supplied in small holes 125
punctured in the agar plate. Following overnight incubation at 37 C° the presence or absence of 126
cellular growth can be determined by visual inspection (Fig. 1B). 127
To search for compounds targeting chromosomal replication initiation, 400 microbial extracts 128
derived from a collection of filamentous actinomycetes were screened using the pBR322-DARS2 129
setup. Seven extracts rescued the growth of the pBR322-DARS2 strain on minimal rich medium. 130
These seven hits were then tested in the hda-screen, giving six strong hits and one weaker; judged 131
from the diameter of the growth zone at non-permissive conditions (Fig. 2A). Extract 18C9 derived 132
from a Streptomyces sp. ID. 62762 gave a strong response in both the pBR322-DARS2- and the 133
hda-screen, and was therefore chosen for further characterization and fractionated into 24 fractions 134
by high performance liquid chromatography (HPLC). 135
136
Identifying the active compound of extract 18C9 137
63
To identify the active compound in extract 18C9, the 24 HPLC fractions were screened using the 138
hda-screen. Only fraction five and six rescued the growth of hda mutant cells, indicating that these 139
contained the active compound (Fig. 2B). These two fractions were then analyzed by HPLC and 140
mass spectrometry (MS). Figure 2C depicts the HPLC chromatogram and MS results for fraction 141
five. In the HPLC chromatogram, there is a distinctive peak between five and six minutes that was 142
only abundant in in these two fractions. MS analysis of the HPLC peak revealed a peak at 585 m/z 143
585 [M-2H+Al], with a clear MS fragmentation pattern. Submission of the MS data in the GNPS 144
database identified the compound as deferoxamine (DFO), a known iron-chelator. 145
146
Deferoxamine rescues the growth of the pBR322-DARS2 strain and the hda mutant 147
Iron plays a key role for many important processes in microorganisms, including reduction of 148
oxygen for ATP synthesis and amino acid synthesis (Roosenberg et al., 2000). Although iron is one 149
of the most abundant elements, the most common oxidation state iron (III) is very insoluble under 150
physiological conditions. Therefore, many microorganisms secrete iron-chelators, also known as 151
siderophores, to scavenge and solubilize iron from their environment to be transported across the 152
cell membrane (Hider & Kong, 2010). DFO, the presumed active compound in extract 18C9, is a 153
siderophore that is produced and secreted by different Streptomyces species (Barona-Gomez et al., 154
2004). To assess whether DFO is indeed the active compound that can rescue growth of over-155
initiating cells, five different DFO concentrations were tested in both the pBR322-DARS2 and hda 156
screen. All five DFO concentrations resulted in growth rescue at non-permissive conditions for both 157
types of over-initiating cell types (Fig. 2D). Note that a higher level of DFO was needed to rescue 158
cells carrying a pBR322- DARS2 plasmid in agreement with these cells having the strongest over-159
initiation phenotype. We estimated the minimal hda rescuing concentration of DFO to be at ~8µg 160
ml-1 (Fig. S1). 161
64
162
Deferoxamine does not prevent bacterial growth 163
In cells grown under permissive conditions, i.e. on minimal poor medium, a clearing zone was 164
observed around the point of DFO addition (Fig. 2D), suggesting that DFO can interfere with E. 165
coli growth. To evaluate the antimicrobial activity of DFO against wild-type E. coli, we attempted 166
to determine the minimal inhibitory concentration (MIC) for DFO with 512 µg ml-1 as the highest 167
concentration. Consistent with previous reports (Thompson et al., 2012) we did not observe 168
complete growth inhibition even at concentrations as high as 512 µg ml-1. However, we observed a 169
~20 pct reduction in doubling time of wild-type cells at DFO concentrations ranging from 100 µg 170
ml-1 to 10 µg ml-1 (Fig. S2). This Indicates that the clearing zone observed around the point of DFO 171
addition most likely reflects growth retardation due to iron depletion. 172
173
Deferoxamine does not inhibit initiation of chromosome replication 174
When wild-type cells were grown in minimal poor medium and treated with rifampicin and 175
cephalexin prior to flow cytometric analysis, they were found to contain mainly one, two or four 176
fully replicated chromosomes indicating the same number of origins prior to drug addition (Fig. 3). 177
When shifted to minimal rich medium, the doubling time decreased from 90 minutes to 35 minutes 178
and cells contained mainly two and four replication origins in accordance with the increased growth 179
rate (Cooper & Helmstetter, 1968). The addition of 150 M DFO to the minimal rich medium 180
increased the doubling time from 35 minutes to 43 minutes and the number of origins per cell 181
decreased somewhat consistent with the reduced growth rate. The origin concentration did not 182
change in the presence of DFO suggesting that it does not affect initiation of replication in wild-183
type cells. 184
65
Cells deficient in Hda and cells containing the pBR322-DARS2 plasmid had an increased number 185
of origins per cell when grown in minimal poor medium and over-initiated replication as 186
demonstrated by an increased origin concentration. When these cells were shifted to minimal rich 187
medium for four hours the number of origins per cell increased from an average of 2.9 and 3.0 to >7 188
and >8 for hda mutant and pBR322-DARS2 carrying cells, respectively (Fig. 3). Note that the 189
replication run out following treatment with rifampicin and cephalexin was incomplete and the 190
cellular number of origins is therefore underestimated (Fig. 3). When the same cells were shifted to 191
minimal rich medium in the presence of DFO the situation was different. The number of origins per 192
cell increased somewhat due to the increased growth rate, replication runout was complete and the 193
origin concentration remained the same or was only slightly elevated (Fig. 3). 194
Altogether this suggests that DFO does not reduce initiations from oriC and that this is not the 195
mechanism behind the rescue of over-initiating cells. 196
197
Deferoxamine does not rescue over-initiating cells by reducing their growth rate. 198
We have previously shown that lethal over-initiation in hda mutant cells can be suppressed by slow 199
growth (Charbon et al., 2017a). Because the presence of DFO was found to slow down the growth 200
of wild-type cells we wondered whether this could be the mechanism behind the rescue of hda 201
mutant and pBR322-DARS2 carrying cells. 202
We therefore tested the ability of DFO to rescue the hda mutant in the richer LB medium. Wild-203
type and hda mutant cells were grown exponentially in presence of 150 M DFO for more than 12 204
generations and had doubling times of 28 and 31 minutes, respectively (Fig. 4A). In minimal 205
medium with DFO hda mutant cells had a doubling time of 60 minutes (Fig. 3). 206
We proceeded to shift cells from DFO containing to DFO free medium. During such a shift the 207
doubling time of the hda mutant increased (Fig. 4A insert) and eventually ceased altogether. The 208
66
number of origins per cell increased from ~15 to more than 30, while the origin concentration could 209
not be determined precisely due to an incomplete run-out. This demonstrates that the presence of 210
DFO ensures viability of hda mutant cells even at doubling times as fast as 31 minutes, where cells 211
over-initiate dramatically. The aggravation of the growth and replication phenotypes after DFO 212
removal also indicates that the DFO rescue was not due to accumulation of suppressor mutations 213
(Kato & Katayama, 2001, Fujimitsu et al., 2009, Charbon et al., 2011, Riber et al., 2006). We 214
therefore conclude that DFO does not rescue hda mutant cells by merely reducing their growth rate. 215
216
Deferoxamine increases processivity of replication forks in over-initiating cells 217
The flow cytometry histograms of over-initiating cells at non-permissive conditions indicated that 218
these failed to complete replication in the presence of rifampicin and cephalexin (Figs. 3 and 4). We 219
therefore determined the origin to terminus ratio (ori/ter) for wild-type, hda mutant cells and cells 220
carrying a pBR322-DARS2 plasmid during growth on minimal poor medium and four hours 221
following a shift to minimal rich medium (Fig. 4B). As expected the ori/ter ratio for wild-type cells 222
only increased from 1.2 to 2.4 when shifted from minimal poor to minimal rich medium, as 223
expected from the increase in growth rate (Fig. 4B). On the other hand the ori/ter ratio for hda 224
mutant cells and cells carrying a pBR322-DARS2 plasmid increased from 1.6 and 1.7 to >25 and 225
>75, respectively, following the same shift (Fig. 4B) suggesting that many replication forks initiated 226
at oriC never reach the terminus in these cells. Again this is in agreement with the strongest over-227
initiation phenotype elicited by the pBR322-DARS2 plasmid. The presence of 150 M DFO in 228
minimal rich medium reduced the ori/ter ratio of hda mutant cells and cells carrying a pBR322-229
DARS2 plasmid from >25 and >75 to 2.0 and 5.0 relative to cells without DFO, respectively (Fig. 230
4B). Altogether, this indicates that DFO helps the DNA replication elongation process in over-231
initiating cells. 232
67
233
Optimization of the pBR322-DARS2 and hda screens by addition of excess iron. 234
The ability of DFO to ensure viability of over-initiating cells by promoting replication elongation 235
was not surprising as it is known that oxidative damage to DNA is a main reason for inviability 236
(Charbon et al., 2011, Charbon et al., 2017a, Babu et al., 2017, Charbon et al., 2014). A major 237
source of ROS species that can cause oxidative damage is the iron dependent Fenton reactions 238
which are inhibited by DFO (Imlay et al., 1988, Liu et al., 2011), most likely by its ability to bind 239
iron. 240
In order to reduce the risk of false positives such as iron chelators and reducing agents that lower 241
ROS formation in our screens, we added excess iron (II) or (III), in the form of Fe(ClO4)2 or FeCl3, 242
when performing the hda based screen (Fig. 5). The rationale behind adding excess iron to the 243
plates, was to ensure that a given iron chelator would not deplete iron in the plates to a level that 244
limit the generation of ROS, and rescue the over-initiating cells in this manner. To test the 245
hypothesis, 5 l of 10 mM of the four iron chelators; DFO, phenanthroline, bipyridyl, EDTA and 246
the reducing agent dithiothreitol (DTT) were tested, with iron (II) or (III) at a final concentration of 247
3 or 200 µM in the plates. DFO, phenanthroline and EDTA all rescued the growth at the standard 248
iron (II) or (III) concentration of 3 µM, while DTT and bipyridyl only did at higher concentration 249
(Fig. 5 A). When iron (II) is at a final concentration of 200 µM, the rescuing effect of EDTA, DFO 250
and phenanthroline was no longer observed (Fig. 5A). As expected the DFO effect was also 251
counteracted by iron supplementation in the pBR322-DARS2 screen (Fig. S3). These results are 252
also consistent with the recovery of growth rate observed when wild-type cells treated with DFO 253
are provided with excess iron in the liquid medium (Fig. S4), i.e. DFO treated cells are depleted for 254
iron. This demonstrates that a high level of iron (II) in the agar plates removes falls positives from 255
iron chelators and reducing agents in the screens. 256
68
To verify that a high level of iron (II) in the screen did not negatively interfere with 257
the detection of DNA replication inhibitors, we assessed the IPTG dependent expression of either 258
the negative initiation regulator SeqA (Lu et al., 1994, Campbell & Kleckner, 1990, von 259
Freiesleben et al., 2000, Charbon et al., 2011) or a cyclic DnaA domain I derived peptide inhibiting 260
DnaA activity (Kjelstrup et al., 2013) in the hda based screen (Fig. 5B). Production of either the 261
cyclic peptide or SeqA was able to rescue the hda mutant cells in presence of 200 µM iron (II). A 262
high level of iron therefore did not have a negative effect on the screen. (Fig. 5 A,B). 263
Finally, we determined whether the remaining six positive hits from the initial screen (Fig. 2A) 264
were false positives by subjecting them to the hda screen with 200 µM iron in the agar plates. This 265
time the six extracts did not rescue the growth of the hda mutant, indicating that they were false 266
positives, most probably preventing ROS formation one way or another. 267
268
(±)-6-Chloro-PB hydrobromide (S143) rescues the growth of the hda mutant 269
Previously, Johnsen et al. reported that the benzazepine derivate (±)-6-Chloro-PB hydrobromide 270
(S143) rescued the growth of over-initiating cells (Johnsen et al., 2010). The rescuing effect of 271
S143 was assigned to a partial inhibition of the DNA gyrase, demonstrated by a supercoiling assay 272
and by countering growth inhibition caused by gyrase overproduction (Johnsen et al., 2010). When 273
tested as a 10 mM solution in our hda based screen, S143 rescued growth of the mutant on plates 274
with iron (II) at a final concentration of 3 µM but not 200 µM (Fig. 6A). S143 also gave rise to a 275
clearing zone when tested on wild-type cells (Fig. 6B) suggesting that the compound interfere with 276
bacterial growth. The growth inhibition could be overcome by addition of Iron (II) at final 277
concentration of 200 µM (Fig. 6 B). Taken together these results indicate that S143 affects iron 278
homeostasis. Note that in presence of excess iron in the plate, S143 changes color (Fig. 6 B). 279
280
69
S143 chelates iron 281
The structure of S143 indicates that it may have a catechol type iron chelation activity (Fig. 7). 282
Catechol groups are found in many siderophores such as E.coli’s enterobactin that contains three 283
catechol groups and has an extremely high affinity for chelating iron(III)(Raymond et al., 2003). 284
We first tested the ability of S143 to outcompete the chelation of iron II by phenanthroline using 285
DFO as a control. Phenanthroline complexes with iron (II) (3:1) and absorbs light at 510 nm. We 286
measured absorbance at 510 nm when a limiting amount of iron (II) was mixed with increasing 287
amount of S143 or DFO prior to addition of a fixed amount of phenanthroline (Fig. 6 C). It was 288
clear that both DFO and S143 outcompete phenanthroline, with DFO being more efficient, 289
indicating that both compounds here are able to bind iron (Fig. 6 C and Fig. S5), although both 290
preferably bind iron(III) over iron(II). Because our assay is performed aerobically in unbuffered 291
ddH2O, the assay likely shows in all or in part, binding of S143 to iron (III) due to iron (II) 292
oxidation. When S143 was mixed with iron (II) perchlorate or iron (III) nitrate, the mixture became 293
green. We therefore measured the absorption spectrum of S143 mixed with iron (III) nitrate. The 294
absorption spectrum indicates that iron (III) and S143 forms complexes absorbing at ~450 nm and 295
~700 nm (Fig. 6 D). Altogether, these data indicate that S143 binds iron as expected for a catechol-296
containing ligand, however at tested conditions the mono-complex is formed rather than the bis- or 297
tris-complex (Sever & Wilker, 2004). 298
299
Discussion 300
We designed screens to identify inhibitors of initiation of chromosome replication. We made use of 301
the fact that hda mutants or cells carrying a pBR322-DARS2 plasmid accumulate DnaAATP, hyper-302
initiate replication, accumulate strand breaks and eventually die. Consequently, compounds that 303
reduce the initiation frequency are expected to restore viability. These screens leave the DnaA 304
70
protein intact as opposed to a related screen employing DnaA mutated in the AAA+ domain 305
(Johnsen et al., 2010, Fossum et al., 2008). Our approach uses a dual sensitivity assay, with 306
pBR322-DARS2 cells being the most selective (Fig. 2). Testing hda and DARS2 assays also has the 307
advantage of discarding certain type of compounds that would be false positives in the pBR322-308
DARS2 screen. These include a molecule that inhibits plasmid replication of the pBR322, as this is 309
expected to restore growth of pBR322-DARS2 transformed cells but not hda mutant cells (not seen 310
with the collection of extracts tested insofar). 311
Deferoxamine was identified from an extract of Streptomyces sp. ID. 62762 as a molecule that 312
restores viability of both types of hyper-initiating cells. Deferoxamine is a siderophore produced by 313
actinomycetes that has been in use as therapeutic agent for iron or aluminum poisoning (Barata et 314
al., 1996) . Because of its iron chelation properties, it has also been tested as a bacteriostatic agent, 315
albeit with poor outcome (Thompson et al., 2012). Successful anti-microbial use of siderophores 316
was previously reported (Saha et al., 2016) but only for species unable to use the siderophore in 317
question. For bacteria capable of using DFO as a siderophore, the situation is reversed (D'Onofrio et 318
al., 2010) and DFO enhance the growth of Klebsiella pneumoniae and increase the susceptibility of 319
mice to infections caused by Yersenia enterocolitica (Chan et al., 2009, Robins-Browne & Prpic, 320
1985). 321
We found that although DFO reduced the growth rate of E. coli, the minimal inhibitory 322
concentration was in above 512 µg ml-1 indicating that it failed to display bacteriostatic or 323
bactericidal effects below this concentration in agreement with previous data (Thompson et al., 324
2012). We found no indication that DFO affects DNA replication in wild-type cells since its 325
presence affect neither origin concentration nor initiation synchrony. 326
The reason for identifying DFO in our screens may solely come from its ability to chelate iron and 327
thus inhibit the Fenton reaction such as described previously in vitro (Imlay et al., 1988) and in 328
71
vivo (Liu et al., 2011). This results in reduced generation of ROS and hence a reduced level of 329
oxidative damage in cells treated with DFO. While there is a narrow time window to repair 330
oxidative damage prior to passage of the next replication fork in wild-type cells (Takahashi et al., 331
2017, Charbon et al., 2014, Foti et al., 2012), forks are more frequent and closely spaced in hyper-332
initiating cells where they occasionally encounter a single stranded region resulting from repair of 333
oxidized bases. This results in double stranded DNA breaks, the ultimate reason for cell death 334
(Charbon et al., 2014). Overall, we therefore suggest that DFO acts by binding iron to reduce ROS 335
generated by the Fenton reactions. This results in a reduced level of oxidative DNA damage, which 336
in turn permits closely spaced replication forks to proceed unimpeded in hyper-initiating cells. This 337
explains why hda mutant cells and cells carrying a pBR322-DARS2 plasmid have a close to wild-338
type ori/ter ratio when treated with DFO despite of continued over-initiation. This is also in 339
agreement with data showing that hyper-initiating cells that generate less or no ROS, due to 340
anaerobic growth or due to having their energy metabolism shifted towards fermentation are viable, 341
as these cells has less or no ROS inflicted DNA damage that need repair (Charbon et al., 2017a, 342
Charbon et al., 2014). The iron chelator bipyridyl and other reducing agents were found to have the 343
same effects as DFO 32. 344
Finally, we tested the benzazepine derivate (±)-6-Chloro-PB hydrobromide (S143) that rescued the 345
growth of cells carrying the conditional hyperactive DnaA219 protein at non-permissive conditions 346
(Johnsen et al., 2010, Fossum et al., 2008) and found it capable of rescuing the growth of hda cells. 347
S143 was initially described as a selective agonist of the dopamine D1-like receptor (Weed et al., 348
1993) but has also been proposed to be a partial inhibitor of E. coli DNA gyrase (Johnsen et al., 349
2010, Fossum et al., 2008). However, this seems unlikely as the ability of S143 to rescue the 350
growth of hda cells was counteracted by addition of excess iron. We predict that S143 is able to 351
bind iron up to a 3:1 stoichiometry (Fig. 7) through its catechol group and demonstrated that it 352
72
forms complexes with iron (II or III). We therefore suggest that the S143 mode of action is, like for 353
DFO, explained by its iron chelating properties. This is also in agreement with S143 being selected 354
as a molecule capable to promote survival of myocardial cells exposed to toxic level of H2O2 (Gero 355
et al., 2007). Here it was concluded that S143 is an indirect inhibitor of cellular PARP activity. 356
Viewing our results, another likely explanation can be found in the chelation of iron and thereby 357
reducing the Fenton reactions. We suggest that S143 chelates iron (and likely other metals) and that 358
this activity is responsible in all or in part for the effects previously observed with this drug. 359
It seems clear that a drawback of our screens is that they will identify molecules that limit reactive 360
oxygen species mediated DNA damage. DFO and other siderophores are often co-produced with 361
other metabolites by actinomycetes. As DFO clearly did not represent the type of molecules we 362
(and other) originally pursued, we adapted our screens to avoid “Fenton reaction moderators” by 363
adding iron in excess in the growth medium. This still allowed identification of replication initiation 364
inhibitors because overproduction of SeqA or a DnaA Domain I derived peptide came out positive 365
in the modified screens. With these modified screens we retested all of our original hit extracts and 366
found none of them to be positive, suggesting that naturally occurring replication initiation 367
inhibitors isolated from actinomycetes strains are found at a much lower frequency in natural 368
extracts than iron chelators such as DFO. 369
370
Experimental procedures 371
Medium 372
Cells were grown in Lysogeny Broth (LB) medium or AB minimal medium (Clark & Maaløe, 373
1967) supplemented with 10 µg ml-1 thiamine and either 0.2% glycerol (minimal poor medium) or 374
0.2% glucose and 0.5% casamino acids (minimal rich medium). 375
376
73
Bacterial strains and plasmids 377
All strains used are derivatives of the E. coli strain MG1655 (F- λ- rph-1) (Guyer et al., 1981). The 378
deletion of hda was performed by P1 mediated transduction (Miller, 1972) as described previously 379
(Riber et al., 2006) and plated on minimal poor medium. The pBR322-DARS2 plasmid is described 380
in (Charbon et al., 2014) (Bolivar et al., 1977). pRNK4 is derived from pSC116 (Kjelstrup et al., 381
2013) by digestion with PvuI followed by re-ligation, thereby removing the chloramphenicol 382
resistance gene and reconstituting the ampicillin resistance gene. Plasmid pMAK7 was described 383
previously (von Freiesleben et al., 2000). 384
385
Chemicals and reagents 386
Deferoxamine mesylate salt (CAS:138-14-7), (±)-6-chloro-PB hydrobromide (S143, CAS:71636-387
61-8), 2,2´-Bipyridyl (CAS:366-18-7), 1,10-Phenanthroline (CAS:66-71-7), Iron (III) chloride 388
hexahydrate (CAS:100025-77-1), Iron (III) nitrate nonahydrate (CAS:7782-61-8) and Iron (II) 389
perchlorate hydrate (CAS:335159-18-7) were all purchased from Sigma-Aldrich. While EDTA 390
disodium salt (CAS:6381-92-6) and DL-Dithiothreitol (CAS:3483-12-3) was purchased from 391
Chemsolute and VWR Life science, respectively. 392
393
hda screen 394
MG1655 hda::cat was grown overnight in minimal poor medium containing 20 µg ml-1 395
chloramphenicol. The overnight culture was diluted to OD600 = 0.0004 (approximately 2x105 cfu 396
ml-1) in 0.9% NaCl. 100 µl of the diluted culture was then plated on minimal poor medium and 397
minimal rich medium plates. Holes were punched in the agar plates using a glass-pipette, and the 398
extracts or compounds to be tested were dispensed into these holes. Following overnight incubation 399
74
at 37oC, the minimal rich medium plates were inspected for growth rescue and the minimal poor 400
medium plates for inhibition zones. 401
402
Multi-copy DARS2 Screen 403
MG1655/pBR322DARS2 and MG1655/pBR322 were grown overnight in minimal poor medium 404
containing 150 µg ml-1 ampicillin. The overnight cultures were then diluted to OD600 = 0.0004 405
(approximately 2x105 cfu ml-1) in 0.9% NaCl. 100 µl of the diluted culture was then spread on 406
minimal rich medium and minimal poor medium plates, containing 150 µg ml-1 ampicillin. A glass-407
pipette was used to punch holes in the agar, and the extracts or compounds to be tested were 408
dispensed into these holes. Following overnight incubation at 37oC, the minimal rich medium plates 409
were inspected for growth rescue and the minimal poor medium plates for inhibition zones. For 410
screening the 400 microbial extracts, 5µl of extract was dispensed in the holes of the plates, and 5µl 411
10% DMSO was used as a negative control. 412
413
Preparation of microbial extracts 414
The 400 microbial extracts were prepared by Naicons srl., Milan, Italy. 10 ml cultures of 415
filamentous actinomycetes were centrifuged at 3000 rpm for 10 minutes to separate the cells from 416
the supernatant. 4 ml ethanol was added to the pellet and incubated for 1h at room temperature with 417
shaking. 0.2 ml aliquots of the ethanolic extracts were distributed in 96-well microtiter plates, dried 418
under vacuum, and stored at 4°C. HP20 resin (Mitsubishi Chemical Co., 1 ml) was added to the 419
supernatant and incubated for 2 hours at room temperature with shaking. The resin was washed with 420
6 ml H2O, and eluted with 5 ml 80% MeOH. 0.25 ml aliquots were distributed in 96-well microtiter 421
plates, dried under vacuum, and stored at 4°C. 422
423
75
HPLC fractionation of extract 18C9 424
Extract, from plate-well 18-C9, was dissolved in 100 l of 80% MeOH. 90 l were fractionated by 425
HPLC on a Shimadzu LC 2010A-HT with the following settings, Column: Merck LiChrosphere 426
RP-18, LiChrocart 5 μm 4.6 x 125mm, phase A: 0.01M HCOONH4 (ammonium formate), phase B: 427
MeCN, flow: 1 ml min-1 at 50°C, UV detection: 230 nm. Linear gradient of phase B: 10 to 95% in 428
18 minutes followed by 5 minutes at 95%. 24 fractions (1 ml each) were collected. 100 l of each 429
fraction were stored for LC/MS analysis while the remaining was dried in a speedvac at 40°C 430
overnight and re-dissolved in 100 µl 10% dmso for the screening. 431
432
Identification of Deferoxamine from extract 18C9 by LC-MS. 433
LC-MS analyses was carried out using a Dionex UltiMate 3000 coupled with an LCQ Fleet mass 434
spectrometer equipped with an electrospray interface (ESI) and a tridimensional ion trap. The 435
following settings were used for liquid chromatography: 1 minute of pre-concentration at 10%, a 7 436
minutes linear gradient from 10 to 95%, followed by an isocratic step at 95% of 2 minutes and 1 437
minute of re-equilibration at 10% of CH3CN with an aqueous phase of 0.05% formic acid. The 438
column was an Atlantis T3 C18 5 μm x 4.6 mm x 50 mm at a flow rate of 0.8 ml min -1. The m/z 439
range (120-2000) and the ESI conditions were as follows: spray voltage of 3500 V, capillary 440
temperature of 275 °C, sheat gas flow rate at 35 and auxiliary gas flow rate at 15. The mass data 441
(.RAW files) from Xcalibur were converted to .mzXML file format, followed by submission to the 442
Global Natural Products Social Molecular Networking (Wang et al., 2016) database for de-443
replication. 444
445
Marker frequency analysis by qPCR 446
76
Cells centrifuged 5 minute 8000x g the supernatant discarded and the cells resuspended in 100 l of 447
cold 10 mM Tris pH7.5. The cells were then fixed by adding 1 ml of 77% ethanol and stored at 4 °C 448
until use. For the qPCR analysis, 100 l of ethanol fixed cells were centrifuged 7 minutes at 17000 449
x g, the supernatant discarded and the samples centrifuged again for 30 seconds at 17000 x g, 450
followed by removal of the remaining ethanol. The cell pellet was resuspended in 1ml cold water 451
and 2 µl was used as template for qPCR analysis. The Quantitative-PCR was performed using a 452
Takara SYBR Premix Ex Taq II (RR820A) in a BioRAD CFX96. All ori/ter ratios were 453
normalized to the ori/ter ratio of MG1655 treated with rifampicin for 2h. The origin and terminus 454
was quantified using primers 5′-TTCGATCACCCCTGCGTACA-3′ and 5′-455
CGCAACAGCATGGCGATAAC-3′ for the origin and 5′-TTGAGCTGCGCCTCATCAAG-3′ and 456
5′-TCAACGTGCGAGCGATGAAT-3′ for the terminus. 457
458
Flow cytometry 459
Preparation of samples for determination of number of origin per cell: 1 ml of cell culture was 460
incubated at 37°C for 2 to 4 hours with 300 µg ml-1 rifampicin and 36 µg ml-1 cephalexin. Cells 461
were fixed in 70% ethanol and stored at 4°C, as described for the marker frequency analysis by 462
qPCR. 463
Preparation of samples for determination of cell size: 1 ml of cell culture was placed on ice and 464
fixed as described for the marker frequency analysis by qPCR. 465
DNA Staining; 100-300 µl of fixed cells were centrifugated at 15,000 x g for 15 min. The 466
supernatant was discarded and the pellet resuspended in 130 µl “Staining solution” (90 µg ml-1 467
mithramycin, 20 µg ml-1 ethidium bromide, 10 mM MgCl2, 10 mM Tris pH 7.5). Samples were 468
then kept on ice for a minimum of 10 min. prior to flow cytometric analysis. Flow cytometry was 469
77
performed using an Apogee A10 Bryte instrument. For each sample, 30 000 to 200 000 cells were 470
analyzed. 471
472
Minimal inhibitory concentration 473
The MIC of DFO was determined by micro-dilution in a 96-well plate. MG1655 was grown to an 474
OD600 of 0.5 in minimal rich medium. The culture was diluted to an OD600 of 0.001 in minimal rich 475
medium. 100 µl diluted culture was added to each well of a 96-well plate containing a dilution 476
series of DFO in minimal rich medium, giving a final concentration range of 512 to 0.5 µg ml-1 of 477
DFO. The 96-well plate was incubated at 37 °C for 24 hours and inspected for visible growth 478
inhibition. 479
480
Minimal rescuing concentration 481
MG1655 hda::cat was grown overnight in minimal poor medium at 37°C. The overnight culture 482
was diluted 100x in minimal rich medium and grown for four hours at 37°C. The culture was 483
diluted to an OD600 of 0.001 in minimal rich medium and 100 µl culture was added to each well of a 484
96-well plate containing a dilution series of DFO in minimal rich medium, giving a final 485
concentration range of 512 to 0.5 µg ml-1 of DFO. The growth at 37 °C during continuous shaking 486
was monitored for sixteen hours, using a Biotek Synergy H1 plate reader. 487
Acknowledgments 488
The authors were funded by grants from the Danish National Research Foundation (DNRF120) 489
through the Center for Bacterial Stress Response and Persistence (BASP) and by the University of 490
Copenhagen Centre for Control of Antibiotic Resistance (UC-Care). 491
78
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81
Fig. 1. Concept of the screen.A) Principle of the screen. In absence of Hda or in presence of multiple copies of DARS2, DNA replication commences too soon and/ or too often resulting in inviability. An anti-DnaA molecule that reduces DnaAactivity reestablishes the initiation frequency to a level that restores viability.Such an anti-DnaA molecule reduces DnaA activity in wild-type cells to a level that no longer sustains viability. B) Schematic representation of the screening method. Hda deficient cells or wild-type cells containing a pBR322-DARS2 plasmid are propagated under permissive growth conditions, i.e. either anaerobic or in minimal poor medium. An estimated twenty thousand cells are spread on two types of agar plates: minimal poor (permissive conditions) and minimal rich (non-permissive conditions) medium. A diffusion assay is performed by punching holes in the agar and introducing 5 ml bioactive extract into each. The plates are incubated aerobically at 37oC for 16h and visually inspected. On the non-permissive conditions plates, positive “hits” are depicted by a small clearing area separating a zone of growth encircling the hole from which the specific extract has been diffusing. The same extract on permissive conditions is depicted by a small clearing area encircling the hole from which the extract has been diffusing.
82
Fig 2. Identification of deferoxamine as a hit.A) Seven extracts rescue the growth hda mutant cells.
Hda deficient cells spread on minimal rich medium plates were tested against seven extracts (19H5, 19C8, 19A6, 18C2, 18H6, 18F7and 18C9). A zone of growth is visible around the holes where the 5 ml of extracts have been introduced.B) Hda deficient cells spread on minimal rich medium plates tested against HPLC separated fractions of extract 18C9. Rescuing activity is seen with fraction 5 and 6.C) LC-MS analysis of fraction 5 identifying deferoxamine as the active compound.D) Hda deficient cells or cells carrying a multi-copy DARS2 plasmid were spread on the indicated plates and tested against varying concentration of deferoxamine. 5 ml of 76, 38, 19, 9.5 and 4.25 mM deferoxamine was dispensed in separated wells.
83
Fig. 3. Deferoxamine does not affect initiation of DNA replication.The indicated cells were grown exponentially at 37 °C in minimal medium supplemented with minimal poor medium (blue) and then diluted into minimal rich medium and incubated for 4 hours at 37 °C in absence of DFO (green) or presence of DFO at a final concentration of 150mM (orange). Cells were treated with rifampicin and cephalexin prior to flow cytometricanalysis. Each panel represents a minimum of 30000 cells. When relevant, the average ori/cell (O/C), ori/mass (O/M) relative to the wild-type (wt) and mass doubling time (t) are shown in the histograms. Inserts show growth of culture where no meaningful doubling time could be obtained. N.D. – Not Determined, N.R. – Not Relevant.
84
Fig. 4. Deferoxamine restores growth hda mutant cells during fast growth.A. Wild-type and Hda deficient cells were grown in LB supplemented with 150 mM DFO. Cells were diluted 5 times in LB without DFO and maintained by dilution in fresh medium for two hours. When indicated Iron II perchlorate was added at a final concentration of 200 mM to titrate DFO. Insert: Growth of hda mutant cells was followed by measuring OD600. Cells were treated with rifampicin and cephalexin prior to flow cytometric analysis. Each panel represents a minimum of 30000 cells. When relevant, the average ori/cell (O/C), ori/mass (O/M) relative to the wild-type (wt) and mass doubling time (t) are shown in the histograms. Orange histograms represent cells grown in the presence of DFO whereas green histograms were derived from cells grown/incubated without DFO for the indicated time. N.D. – Not Determined, N.R. – Not Relevant.B. DFO promotes replication fork progression in Hda deficient cells or wild-type cells containing a pBR322-DARS2 plasmid. The indicated cells were grown exponentially at 37 °C in minimal poor medium (blue), shifted to minimal rich medium and incubated for 4 hours at 37 °C in absence of DFO or presence of DFO at a final concentration of 150 mM. The ori/ter ratios were determined by qPCR analysis. Shown is the mean ± s.d. (n=3).
85
Fig. 5. The effect of iron chelators and reducing agent can be eliminated by addition of excess iron. A. Hda deficient cells were spread on minimal rich medium agar plates containing iron (III) chloride at a final concentration of 3 mM, iron (II) perchlorate at a final concentration of 3 mMor iron (II) perchlorate at a final concentration of 200 mM and tested against metal chelatorsand antioxidant. 5 ml of 10 mM DFO, 10 mM phenanthroline, 10 mM EDTA, 300 mM bipyridilor 650 mM DTT was dispensed in separated wells. B. Hda deficient cells capable of producing SeqA or a cyclic DnaA domain I derived peptide were spread on minimal rich medium agar plates containing 3 mM iron (III) chloride , 3 mMiron (II) perchlorate or 200 mM iron (II) perchlorate. 5 ml of 100 mM IPTG was dispensed in separated wells to induce the overexpression of SeqA or a cyclic DnaA domain I. 5 ml of 10 mM DFO was used as control.
86
Fig. 6. S143 chelates iron.A. Hda mutant cells were plated on minimal rich medium agar plates containing 3 mM iron (III), 3 mM iron (II) perchlorate or 200 mM iron (II) perchlorate were tested against 5 ml of 10 mM S143. B. Wild-type cells were plated on minimal poor medium agar plates containing either 3 mMiron (II) perchlorate or 3 mM or 200 mM iron (II) perchlorate and tested against 5 ml of 10 mMS143. C. Iron binding of S143 and DFO was assayed by monitoring the absorbance at 510nm of the Fe (II)-Phenanthroline complex. Increasing amounts of DFO or S143 was mixed with iron (II) perchlorate (0.015 mM final concentration) and absorbance at 510nm was measured following addition of phenanthroline (1mM final concentration). The absorbance relative to Fe (II)-Phenanthroline is plotted.D. Absorption spectrum of 1mM S143 in ddH2O alone or complexed with 0.2 mM or 0.4 mMiron (III) nitrate.
87
Fig. 7. Model structure for DFO and S143 chelating ironA single DFO molecule forms six bonds with iron (III) while up to three S143 molecules can interact with one iron (III) through chelation by their catechol moieties.
88
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.0 2.0 4.0 6.0 8.0 10.0 12.0 14.0 16.0
32 g DFO ml-1
16 g DFO ml-1
8 g DFO ml-1
4 g DFO ml-1
2 g DFO ml-1
1 g DFO ml-1
0.5 g DFO ml-1
no DFO
Time (hours)
O.D
. 600
nm
Figure S1
Fig. S1 DFO Minimal Recovery Concentration.
Hda cells pre-grown in minimal poor medium were shifted to minimal rich medium at 37 °C in the
presence of DFO at different concentration (see experimental procedures). The growth was moni-
tored by measuring optical density in a microplate reader. Hda deficient cells grown with 32 to 0.5
μg ml-1 of DFO are shown. Cells grown with 8 μg DFO ml-1 and above started growth earlier than
those grown with 4 μg DFO ml-1 and below. The late grown cells may contain mutations suppress-
ing hda.
89
50 60 70 80 90 100 110 120 130 140 150
100 g DFO ml-1
50 g DFO ml-1
25 g DFO ml-1
10 g DFO ml-1
no DFO
O.D
.450
nm
(log
)
time (min)
Figure S2
Fig. S2 DFO Minimal Recovery Concentration.
The effect of DFO on wild type growth. Wild type cells were grown in minimal rich medium and
maintained exponentially growing in absence or in presence of 10, 25, 50 or 100 μg ml-1 DFO.
Growth is monitored by optical density measurement.
90
DFO DMSO
200M Iron(II) perchlorate
glu +casa
3M Iron(III) Chloride
Figure S3
Fig. S3 Excess iron counteracts the effect of DFO in the pBR322-DARS2 screen.
Cells carrying a multi-copy DARS2 plasmid were spread on minimal rich medium agar plates
containing iron (III) chloride at a final concentration of 3 μM or iron (II) perchlorate at a final
concentration of 200 μM and tested against DFO. 5 μl of 10 mM DFO or DMSO was dispensed in
separated wells.
91
0 10 30 50
O.D
. 600
nm
(log
)
time (min)
wt
wt 150M DFO
wt 150M DFO 200 M iron (II) perchlorate
Figure S4
Fig. S4 The effect of DFO on wild type growth is counteracted by iron.
Wild type cells were grown in minimal rich medium and maintained exponentially growing in absence
of DFO, in presence of 150μM DFO or 150μM DFO and excess iron (II). Growth was monitored by
optical density measurement.
92
Abs
orba
nce
(AU
)
10 mM S1431.2 mM S1430.3 mM S143
0
0.02
0.04
0.06
0.08
0.1
0.12
0.14
0.16
0.18
0.2
350 450 550 650
0.05 mM S143No S143
(nm)
Figure S5
Fig. S5 S143 chelates iron.
Absorption spectrum of increasing amounts of S143 in ddH2O mixed with iron (II) perchlorate (0.020
mM final concentration) and phenanthroline (1mM final concentration).
93
Paper III: A Novel Fluorescence Based Screen for
Inhibitors of the Initiation of DNA Replication in
Bacteria.
Currently in review at: Current Drug Discovery Technologies.
94
Send Orders for Reprints to [email protected]
Journal Name, Year, Volume 1
XXX-XXX/14 $58.00+.00 © 2014 Bentham Science Publishers
A Novel Fluorescence Based Screen for Inhibitors of the Initiation of DNA
Replication in Bacteria
Rasmus N. Klitgaarda and Anders Løbner-Olesena*
aDepartment of Biology, Faculty of SCIENCE, University of Copenhagen, Copenhagen, Denmark.
Abstract: Background: One of many strategies to overcome antibiotic resistance is the discovery of
compounds targeting cellular processes, which have not yet been exploited. Methods and materials:
Using various genetic tools, we constructed a novel high throughput, cell based, fluorescence screen for
inhibitors of chromosome replication initiation in bacteria. Results: The screen was validated by
expression of an intra-cellular cyclic peptide interfering with the initiator protein DnaA and by over-
expression of the negative initiation regulator SeqA. We also demonstrate that neither tetracycline nor
ciprofloxacin triggers a false positive result. Finally, 400 extracts isolated mainly from filamentous
actinomycetes were subjected to the screen. Conclusion: We conclude that the presented screen is
applicable for identifying putative inhibitors of DNA replication initiation in a high throughput setup.
Keywords: DNA replication initiation, inhibitors, DnaA, high throughput screen, fluorescence, microbial
extracts.
1. INTRODUCTION
Antibiotic resistance is one of the major health care
problems in the world; therefore, it is important to discover
compounds targeting unexploited processes essential to the
growth or viability of bacteria. One such process is
replication of the bacterial chromosome. Targeting the DNA
replication is attractive for a number of reasons; i) The
replisome number per cell is low, ii) The replication complex
is a multi-protein machinery and therefore contains a large
number of potential targets, iii) Key components of the
replication machinery is well conserved and has low
sequence homology with human replication proteins and iiii)
The DNA replication is an under exploited target, this far
only type-II topoisomerase inhibitors are used in the
clinic[1]. A number of different compounds have been
identified targeting components of the replication machinery
including; DNA ligase (LigA)[2, 3], DNA polymerase III[4,
5], the sliding clamp[6, 7] and single-stranded DNA-binding
proteins[8]. In contrast, only a few efforts have been made to
discover inhibitors of the initiation process of the bacterial
DNA replication [9-11].
In most bacteria, replication of the chromosomal
DNA is initiated from a single origin of replication, termed
oriC. The initiation process is best characterized in
Escherichia coli, where DNA replication is initiated by
binding of the initiator protein DnaA, in its active ATP-
bound form, to the oriC. When sufficient DnaAATP
molecules are bound to the oriC it forms a nucleoprotein
complex responsible for separation of the DNA double
strand[12]. Following duplex opening the nucleoprotein
complex loads the DnaB helicase, with help from the
helicase loader protein DnaC [13, 14]. Loading of DnaB then
triggers the assembly of the remaining parts of the
replication machinery[12]. The initiation process, including
the DnaC assisted loading of the DnaB helicase, is highly
conserved across bacterial species [15], and is therefore an
interesting target for novel antibiotics.
E. coli rnhA mutants, lacking RNase HI activity, are
able to initiate the DNA replication by a protein synthesis
and DnaA/oriC independent pathway called constitutive
Stable DNA Replication (cSDR)[16, 17]. cSDR is initiated
at a number of alternative sites on the chromosome, termed
oriK[18]. It is believed that lack of RNase HI activity leads
to stabilization of nascent RNA transcripts, which anneals to
the DNA template behind the moving RNA polymerase,
creating an R-loop. The RNA is thought to act as a primer
for extension by DNA polymerase I, creating a D-loop like
structure. This structure is then bound by PriA, initiating
assembly of the PriA dependent primosome, loading of the
DnaB helicase and assembly of the replisome [19, 20]. The
DnaA/oriC independency of cSDR makes it a valuable tool
when searching for inhibitors targeting the initiation of DNA
replication.
We here present a high throughput fluorescence based screen, which can be used to specifically screen for inhibitors that targets DNA replication initiation.
*Address correspondence to this author at the Department of Biology, Faculty of SCIENCE, University of Copenhagen, Copenhagen, Denmark; E-mail: [email protected]
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2 Journal Name, 2014, Vol. 0, No. 0 Klitgaard et al.
2. MATERIALS AND METHOD
2.1 Bacterial strains and plasmids
MG1655ΔrnhA::kanR (unpublished, ALO4523) and
MG1655ΔrnhA::KanR, ΔoriC::CamR (unpublished,
ALO4524) were obtained from the laboratories strain
collection. The construction of pMAK7 is described in ref.
[21]. pRNK4 was constructed by PvuI digest of pSC116[7]
followed by re-ligation, thereby removing the
chloramphenicol resistance gene and reconstituting the
ampicillin resistance gene.
2.2 Construction of the mini-chromosome pRNK6
A fragment from the mini-chromosome pRNK3
(unpublished) containing the oriC and cI was PCR
amplified and ligated using MunI and EcoRI into the
OriR6K-dependent vector pSW25T[22], creating pRNK5.
The expression of cI is controlled by a constitutive synthetic
promoter (J23101) from the Anderson promoter
collection[23]. The promoter region was inserted upstream
of cI by PCR amplification of cI from pSB4293[24]. To
stabilize the mini- chromosome a PCR fragment, from
pALO17[25], containing the SopABC partitioning system
was ligated into pRNK5 using EcoRI, creating pRNK6.
2.3 Construction of the screen strain (MG1655ΔrnhA,
ΔoriC::CamR, attB::PR-GFPmut2,KanR)
MG1655ΔrnhA::kanR was the starting point in the
construction of the strain used in the screen. First, the KanR
cassette was flipped out by expression of FLP recombinase
from pCP20[26]. Hereafter, the lambda PR-promoter was
fused to GFPmut2 by PCR amplification from pKEN GFP
mut2[27]. The fragment was then inserted at the lambda
attachment site (attB) on the E. coli chromosome as
described in ref. [28], creating MG1655ΔrnhA, attB::PR-
GFPmut2, KanR. Finally, the oriC was removed by lambda
red recombination [26], deleting a region in the chromosome
from the start codon of viaA to the stop codon of mnmG.
2.4 Primers
Table 1. Primers used for construction of the screen.
Use Sequence
Amplification of cI from pSB4293 and
adding the Anderson promoter J23101
FW
TATAGAGCTCTTTACAGCT
AGCTCAGTCCTAGGTATTAT
GCTAGCGCGGTGATAGATT
TAACGTATGAGCA
Amplification of cI from pSB4293 and
adding the Anderson promoter J23101
RV
GATCGAGCTCTCAGCCAA
ACGTCTCTTCAGG
Amplification of cI/oriC fragment from
pRNK3 (FW)
GATCCAATTGGCCTGACG
GTAGAGCACACGAT
Amplification of cI/oriC fragment from
pRNK3 (RV)
GTATAGAATTCCCGATCAT
GCGTACCATCAAG
Amplification of sopABC fragment from TATAGAATTCTCATGTTTG
pALO17 (FW) ACAGCTTATCATCG
Amplification of sopABC fragment from
pALO17 (RV)
GATCGAATTCCTCGACAG
CGACACACTT
Amplification of GFPmut2 from pKEN
GFP mut2 and adding the PR-promoter
FW
GATCGAATTCGCGTGTTG
ACTATTTTACCTCTGGCGG
TGATAATGGTTGCATGTAC
TAAGGAGGTTGTATGAGT
AAAGGAGAAGAACTTTTC
ACTGGAG
Amplification of GFPmut2 from pKEN
GFP mut2 and adding the PR-promoter
RV
CTTACTCGAGTTATTTGTA
TAGTTCATCCATGCCATGT
GTAATCC
Deletion of oriC FW TTGCCTGGTAAGCGGGTG
CTTACCAGGCATTTTTAAT
GCGGTGTAGGCTGGAGCT
GCTTC
Deletion of oriC RV GCCTACAGGATGTCGGTG
CACAGATTCGCCAGGCAC
AACAATGGGAATTAGCCA
TGGTCC
2.5 Fluorescence screen
Overnight cultures were grown at 370C with the
appropriate antibiotics (40 µg/mL kanamycin, 20 µg/mL
chloramphenicol, 50 µg/mL streptomycin, 150 µg/mL
ampicillin) in AB media supplemented with 0.2% glucose
and 1% casamino acids (ABTG CAA). The overnight
cultures were diluted to OD600 = 0.001 and 100 µL of the
diluted culture was added to the wells of a 96-well plate,
already containing 100 µL ABTG CAA, with the correct
antibiotics and IPTG (final conc. 0.25mM). The plate was
sealed with a Breathe-easy®sealing membrane (Diversified
Biotech) and incubated at 370C for 20 hours while shaking in
a plate shaker (800 RPM). Following incubation, the plates
were centrifuged, the supernatant removed, the cells
resuspended in 200 µL 0.9% NaCl and transferred to clear
bottomed black sided 96-well plate. OD600 and fluorescence
was detected using a BIOTEK synergy H1 plate reader. The
following settings were used for the fluorescence
measurement, excitation: 485nm and emission: 528nm. The
treatment with tetracycline and ciprofloxacin was carried out
as minimal inhibitory concentration assay by micro dilution
in a 96-well plate, the plate was otherwise treated as stated
above. The MIC for tetracycline and ciprofloxacin was 1 and
0,015 µg/ml, respectively.
2.6 Preparation of microbial extracts
The extracts subjected to the screen were prepared
by Naicons srl., Milan, Italy. Cultures of filamentous
actinomycetes (10 mL) were centrifuged (3000 rpm / 10
minutes) to separate the cells from the supernatant. Ethanol
(4 mL) was added to the pellet and incubated for 1h at room
temperature with shaking: 0.2 ml aliquots of the ethanolic
extract were distributed in 96-well microtiter plates, dried
under vacuum, and stored at 4°C. HP20 resin (Mitsubishi
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Short Running Title of the Article Journal Name, 2014, Vol. 0, No. 0 3
Chemical Co., 1 mL) was added to the supernatant and
incubated for 2 hours at room temperature with shaking; the
resin was washed with 6 mL H2O, and eluted with 5 mL
MeOH: 0.25 mL aliquots were distributed in 96-well
microtiter plates, dried under vacuum, and stored at 4°C.
2.7 Extract screening
Dried extracts were dissolved in 99.6% DMSO and
diluted with water to a final concentration of 10% DMSO.
10 µL of each extract was suspended into the wells of a 96-
well polypropylene plate. An overnight culture of the screen
strain was grown and diluted as described above and 190µL
was added to each well containing the extracts. 190µL of the
following control strains were added to wells with 10µL
10% DMSO; i) MG1655ΔrnhA, ΔoriC::CamR, attB::PR-
GFPmut2,KanR, ii) MG1655ΔrnhA, ΔoriC::CamR, attB::PR-
GFPmut2,KanR/pRNK6, iii) MG1655ΔrnhA::KanR,
ΔoriC::CamR. The incubation and measurements were
performed as described in the section above. To assess the
auto-fluorescence of the extracts, the screen was performed
in parallel using the non-fluorescing strain;
MG1655ΔrnhA::KanR, ΔoriC::CamR.
2.8 Analysis of fluorescence data
The fluorescence data obtained from each well in
the 96-well plate was first adjusted for background
fluorescence. This was done by subtracting the fluorescence
of the non-GFP strain, MG1655ΔrnhA::KanR, ΔoriC::CamR.
The fluorescence from each well was compared to the
fluorescence of MG1655ΔrnhA, ΔoriC::CamR, attB::PR-
GFPmut2,KanR/pRNK6, grown in ABTG CAA with 0.5%
DMSO.
3. RESULTS
3.1 Construction and validation of the screen
The screen utilizes an E. coli rnhA, oriC mutant that
replicates its chromosomal DNA by cSDR. As the strain
does not initiate from oriC it is insensitive to putative
inhibitors of the oriC dependent initiation process. The
fluorescence reporter system consists of GFPmut2[27]
expressed from a lambda phage PR-promoter inserted into
the attB site on the chromosome. Transcription from the PR-
promoter is repressed by the lambda phage cI repressor,
which is constitutively expressed on a plasmid (pRNK6) that
only replicates from oriC, also known as a mini-
chromosome. The rationale behind the screen is that
Inhibition of mini-chromosome replication will lead to loss
of cI expression and to expression of GFP, which can be
detected as an increase in fluorescence in a high-throughput
setup. (Figure 1).
Fig. 1. Graphical representation of the screen. A) cI
expressed from the mini-chromosome pRNK6, represses the
expression of GFPmut2 from the PR-promoter. B) If the
initiation of replication by DnaA is blocked, pRNK6 is lost
overtime. Hence, the repression of the PR-promoter is
released and GFP is expressed.
Following construction of the strain,
MG1655ΔrnhA, ΔoriC attB::PR-GFPmut2, it was verified by
microscopy that the cells were fluorescing, confirming that
GFP was expressed from the PR-promoter (Figure 2AB).
Hereafter it was assessed if cI expressed from pRNK6
reduced expression of GFPmut2 from the lambda PR-
promoter. This was done by quantifying the fluorescence,
following 20 hour incubation at 370C, for MG1655ΔrnhA,
ΔoriC attB::PR-GFPmut2 with and without pRNK6. The
results show that the presence of pRNK6 reduced
fluorescence by more than 50% relative to the strain without
pRNK6 (Figure 2C). Indicating that cI is repressing the
expression of GFP as expected. MG1655ΔrnhA, ΔoriC
attB::PR-GFPmut2/pRNK6 is from now on referred to as the
screen strain.
To validate the screen further, we introduced a
plasmid, pRNK4, that encodes a cyclic peptide previously
shown to inhibit DnaA function and hence replication
initiation [7]. In the screen strain expression of the DnaA
inhibitor by IPTG induction, led to a 50% increase in
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4 Journal Name, 2014, Vol. 0, No. 0 Klitgaard et al.
fluorescence relative to the un-induced control (Figure 2D).
The screen was further verified by over-expression of SeqA
protein from the plasmid pMAK7. SeqA regulates the
replication initiation by binding to hemi-methylated GATC
sequences in oriC, thereby sterically hindering initiation by
DnaA[29]. Over-expression of SeqA, by IPTG induction,
increased the fluorescence by 30% relative to the un-induced
control (Figure 2E). Overall these results verifies that if
replication of the oriC dependent mini-chromosome is
inhibited, repression of the PR-promoter is released and GFP
is expressed, leading to an increase in the fluorescence
signal.
Fig. 2. Validation of the screen. A) Fluorescence
microscopy of the strain, MG1655ΔrnhA, ΔoriC attB::PR-
GFPmut2. B) Phase contrast of the same cells as in picture
A. C) Verification that cI, expressed from pRNK6, reduces
expression of GFP by repression of the lambda PR-promoter.
D) Verification of the screen, by expression of a DnaA
inhibitor (pRNK4) and E) over-expression of seqA
(pMAK7) in the screen strain.
3.2 Impact of sub-inhibitory concentrations of antibiotics
on the screen.
Natural extracts from plants and microbes are
frequently used when screening for novel antibiotics, these
extracts are usually complex and often contain several
compounds with antibacterial properties. It is therefore
important to asses if sub-inhibitory concentrations of
antibiotics, would lead to false-positives in the screen.
Specifically, we were interested in the effect of inhibiting
translation or DNA replication elongation. The screen was
therefore subjected to sub-inhibitory levels (0.5xMIC and
0.25xMIC) of either tetracycline, an inhibitor of translation,
or the DNA replication elongation inhibitor, ciprofloxacin.
This resulted in fluorescence signals that were lower relative
to the untreated sample (Figure 3), indicating that sub-
inhibitory concentrations of antibiotics, targeting translation
or DNA replication elongation, does not lead to false
positives.
Fig. 3. Relative fluorescence of the screen strain treated with
0.25x and 0.5xMIC of tetracycline or ciprofloxacin.
3.3 Screening microbial extracts
In an initial attempt to discover a novel inhibitor of
the initiation of DNA replication, we screened 400 microbial
extracts, mainly derived from filamentous actinomycetes.
None of the extracts gave a positive hit in the screen,
suggesting that initiation inhibitors are rare and that a high
number of extracts needs to be screened in order to get a
positive hit.
4. DISCUSSION
In the battle against antibiotic resistance, it is
important to discover and develop novel compounds
targeting essential cellular processes. The screen presented
here could become a valuable tool in identifying inhibitors of
DNA replication initiation in bacteria. We expected that
expression of cI from pRNK6 would repress the expression
of GFP from the PR-promoter. Quantifying the relative
fluorescence with and without pRNK6, showed a partial
reduction in the fluorescence signal of around 50%. This
could indicate that the PR-promoter is leaky, or more likely it
is a result of instability and loss of the mini-chromosome, as
mini-chromosomes become more unstable at slow growth
[30]. As there are no known replication initiation inhibitors
that can cross the E. coli cell membrane, the screen was
validated by expression of a cyclic peptide inhibiting the
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Short Running Title of the Article Journal Name, 2014, Vol. 0, No. 0 5
function of DnaA and over-expression of the negative
initiation regulator SeqA.
We wondered whether non-lethal inhibition of
translation or DNA replication elongation could affect
segregation of the mini-chromosome or the fluorescence
reporter system, in a manner that would lead to false
positives. Subjecting the screen to sub-inhibitory
concentrations of the translation inhibitor, tetracycline, or the
DNA replication inhibitor, ciprofloxacin, confirmed that this
was not the case. Overall, these results show that the screen
is applicable for identifying novel inhibitors of the initiation
of DNA replication. However, it should be noted that other
classes of inhibitors might influence the segregation of the
mini-chromosome, without inhibiting the growth of the
bacteria, leading to an increase in fluorescence. Especially
putative inhibitors of the SopABC segregation system could
lead to false positives. However, one could differentiate
between a replication initiation inhibitor and a SopABC
inhibitor, as the later would most likely not have an effect on
the growth of a wild-type E. coli strain. Furthermore, it is
well known that transcription from the mioC gene promoter,
into the oriC, is important to maintain mini-chromosome
stability and copy number, but that it does not stimulate the
initiation of chromosomal replication[31]. Inhibiting
transcription from the mioC promoter could therefore
potentially lead to a false positive result in the screen.
None of the 400 extracts that were screened gave a
positive hit. However, we gained valuable insight about the
practical procedure for the screen. Furthermore, it indicates
that the screen is specific and that a larger number extracts or
compounds needs to be screened in order to get a positive
hit. The specificity of the screen was not unexpected, as the
main difference between replication by cSDR and replication
from the oriC is the initiation by DnaA[20]. Even the DnaB
helicase and its chaperone, DnaC, have been shown to be
required for cSDR[17]. As the initiation by DnaA is a multi-
step process, direct inhibition hereof can potentially happen
at multiple points. Early in the process, the initial binding of
DnaA to the oriC could be hindered by blocking the
interaction between the HTH DNA-binding motif in domain
IV of DnaA and the oriC[32]. Furthermore, oligomerization
of DnaA at the oriC and formation of the nucleoprotein
complex, could be inhibited by targeting either domain I or
III of DnaA, that have been shown to mediate the
oligomerization [12, 15]. Finally, the process of loading the
DnaB helicase, mediated by interactions between regions of
domain I and III of DnaA and the DnaB helicase [33], might
also serve as a target for putative inhibitors of the initiation
process that would be identified by the screen presented
here. In addition to direct inhibition of DnaA, the initiation
process might also be subdued by putative inhibitors
targeting factors stimulating the initiation process, including;
DiaA[34], H-NS[35], Fis[36], IHF[37] and HU[38]. Though
it should be noted that only DiaA, FIS and HU mutants are
unable to stably maintain mini-chromosomes[34, 38, 39],
hence inhibition of IHF or H-NS may not lead to a positive
hit in the screen[37, 40].
CONFLICT OF INTEREST
The authors declare no conflict of interest.
ACKNOWLEDGEMENTS
We acknowledge financial support from the University of
Copenhagen Centre for Control of Antibiotic Resistance
(UC-Care) and by the Center for Bacterial Stress Response
and Persistence (BASP) funded by a grant from the Danish
National Research Foundation (DNRF120). Finally, we
would like to thank NAICONS srl. for preparing and sharing
their microbial extracts.
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Received: March 20, 2014 Revised: April 16, 2014 Accepted: April 20, 2014
101
Discussion
Antibiotic resistance has become an urgent problem, which not only threatens the future of health care,
as we know it today, but also might turnout out to be a heavy burden for the global economy (12, 13).
An effective strategy to overcome antibiotics resistance will need to be multidimensional. Thus,
facilitate a sustainable use of antibiotics in the clinic as well as in the industry, diminish the spread of
infectious disease, and encourage development of novel drugs and preservation of existing antibiotics.
In the last 10 years over 50 national and international initiatives have been founded with the goal of
encouraging research and development of antibiotics (258). Though action has already been taken;
evidence and experience show that the current market for antibiotics does not foster major investments
from the large pharmaceutical companies, who have set their course for more profitable markets (12).
We have contributed to the fight against antibiotic resistance, by searching for potential helper drug
targets to reverse quinolone resistance and thereby preserve the efficacy of one of the most important
classes of antibiotics. Furthermore, we have developed and verified two distinct strategies for the
discovery of novel classes of antibiotics targeting the initiation of chromosomal DNA replication in
bacteria.
Potentiation of the quinolones
In paper I, we sought to identify targets for potentiation of ciprofloxacin by introduction of more than
twenty separate single gene deletions in a high-level ciprofloxacin resistant strain. None of the tested
gene deletions rendered the high-level resistant strain clinically susceptible. However, deletion of acrA,
tolC, recA or recC decreased the MIC of a low-level ciprofloxacin resistant strain beneath the clinical
break point. Indicating that inhibition of the AcrAB-tolC efflux-pump or HR repair of DNA DSBs, via RecA
or RecC inhibition, is a plausible strategy for reversal of low-level ciprofloxacin resistance. These findings
are in agreement with the observations made by Tran et al. and Recacha et al.(129, 130).
The discovery and development of putative inhibitors of RecA, and thereby the SOS
response, is attractive for several reasons. Most bactericidal antibiotics are inducers of the SOS response
(55), thus inhibition of RecA might not only potentiate the quinolones but also other classes of
bactericidal antibiotics. The SOS response also plays an important role in the evolution of antibiotic
resistance, by inducing horizontal gene transfer (HGT) of antibiotic resistance genes (259) and by
promoting mutagenesis via the error prone DNA polymerases IV and V (86). In addition, induction of the
SOS response has been shown to promote HGT of pathogenicity associated genes in E. coli and S.
102
aureus. Thus, RecA inhibitors could potentially exert a dual mode of action in increasing antibiotic
susceptibility and decreasing evolution of antibiotic resistance and pathogenicity.
As described earlier, multiple efforts have been made to identify putative inhibitors of
RecA, leading to the discovery of several compounds that blocks the ATPase activity of RecA in vitro
(121, 122, 124). It is unknown if any these compounds are still under development. Copper
phtalocyanine-3, 4´, 4´´,4´´´-tetrasulfonic acid (CuPTA) and suramin are the only compounds that
evidently inhibits RecA in vivo (119, 125). In paper I, we report that CuPTA does not change the
ciprofloxacin MIC for a high- or low-level ciprofloxacin resistant strain. Indicating that either CuPTA is a
weak RecA inhibitor or that a given inhibitor needs to completely inactivate RecA for potentiating
ciprofloxacin. Hence, development of an effective RecA inhibitor in regards to potentiation of the
quinolones and other bactericidal antibiotics might prove difficult. In contrast to RecA, there only exist a
single report of identification of putative inhibitors of the RecBCD complex, however the potential of the
identified compounds to potentiate the quinolones has not been assessed (260).
Inhibition of efflux pumps, like AcrAB-TolC, is a well investigated mean of potentiating
antibiotics. Similar to inhibition of RecA, efflux pump inhibition generates some desirable side effects in
addition to decreasing the MIC. In E. coli, deletion of acrAB delays the emergence of levofloxacin
resistance (261), while the virulence of Salmonella enterica to some extend relies on the functionality of
its drug efflux systems (262). A described earlier several EPIs that targets the AcrAB-TolC efflux pump
have been identified. However, not a single EPI has made it through clinical trials and into the clinic
(263, 264). One of the major challenges in EPI development is the broad compound specificity exerted
by most efflux pumps. Consequently, it is challenging to setup guidelines for discriminating between
efflux pump substrates and inhibitors, making it difficult to pick suitable compound libraries for
screening (264). Current challenges in clinical development of EPIs are; toxicity, pharmacokinetics,
potency and spectrum of activity (265). Hence, further insight into the structure and function of efflux
pumps is essential for successful discovery and development of EPIs in the future.
Tran et al. showed that combinatorial disruption of the AcrAB-TolC efflux pump and the
SOS response rendered a high-level ciprofloxacin resistant strain clinically susceptible (129). However,
deployment of this observation in the clinic would require a three drug combinatorial treatment.
Experience from combinatorial drug therapy of cancer, has shown that treatment with multiple drugs is
challenging due to overlapping toxicities and differences in pharmacological profiles (266). Indicating,
that it would be difficult to develop such a treatment, though there is no doubt that combinatorial
inhibition of drug efflux and the SOS response, would be a powerful weapon in overcoming antibiotic
resistance.
103
Based on the findings presented in paper I, it seems that reversing high-level ciprofloxacin
resistance with a single helper drug is not possible. However, during the gene deletion analysis, we were
not able to delete priA in the high-level resistant strain, LM693. A similar observation was reported by
Cirz et al., who were unable to construct an E. coli gyrA(S83L) ΔpriA mutant (72). PriA is the initiator
protein of replication restart, a housekeeping process that facilitates the restart of stalled replication
forks during normal growth (267). E. coli priA- strains suffers from severe growth retardation and are
hyper-susceptible to ciprofloxacin with a MIC below 1 ng/ml (72). The above observations indicates that
PriA inhibition is possibly lethal to bacterial strains with gyrA mutations conferring ciprofloxacin
resistance. Thus, combinatorial treatment of a PriA inhibitor with ciprofloxacin, or cycling between the
two, could be an effective mean of treating infections caused by ciprofloxacin resistant bacteria.
Targeting the commencement of DNA replication in bacteria
Duplication of the chromosome is an essential part of the bacterial cell cycle and its initiation could
potentially serve as a novel antibiotic target. We have presented two novel cell based strategies for
identifying DNA replication initiation inhibitors. The replication initiation process and its regulation is
complex and can therefore, potentially be inhibited via multiple different targets, of which directly
targeting DnaA is the most apparent one. However, the negative or positive regulation of the initiation,
might also serve as potential targets. Emphasized by the fact that deletion of either datA, DARS1 or
DARS2 in E. coli, results in a reduced ability to colonize the large intestine in mice (183).
DnaA binding to the DnaA boxes in the oriC is a plausible target for putative inhibitors of
the replication initiation. Specifically, targeting the HTH motif of DnaA domain IV could in principal block
the binding of DnaA to both the high and low affinity DnaA boxes in the oriC. Interference with the
assembly of the DnaAATP-OriC nucleoprotein complex is attainable in multiple ways. Binding of an
inhibitor in the nucleotide-binding pocket of the AAA+ module of domain III, could block ATP binding
and lock the structural conformation of DnaA in an apo-DnaA or DnaAADP-like state that is inactive in
DnaA oligmerization. In addition, the cooperative binding of DnaAATP could also be inhibited by
interfering with the DnaA-DnaA interactions mediated by specific residues of DnaA domain I and III.
However, DnaA boxes in the oriC are not arranged similarly across bacterial species, indicating that
there are many different ways of assembling the nucleoprotein complex. Thus, it is not necessarily the
same residues that mediates the DnaA-DnaA interactions in different bacterial species (188). Due to
their stimulatory role in the replication initiation process, factors like; IHF, DiaA, HU, and H-NS, might
also serve as targets for inhibiting replication initiation. Although none of these factors are essential for
replication initiation, their deletion does lead to under-initiation (163, 268-270), though it remains to be
assessed if it has a lethal effect in a hostile environment like the human body.
104
Loading of the DnaB helicase by DnaA is an essential step in initiating the replication
process and is therefore an obvious target. Inhibiting the binding of DnaC would likely block the loading
of DnaB onto the ssDUE. As DnaB would not be locked into to the open conformation that is essential
for its loading onto the ssDUE (193). In addition, hindering the interaction between DnaB and DnaA
domain I should also block the loading of the DnaB helicase.
As mentioned above, the processes that regulates the DnaAATP/DnaAADP ratio is also a
potential target for interfering with the replication initiation. Interestingly, over-initiation seems to be
more lethal than decreasing the initiation frequency (237). Thus, inhibiting the conversion of DnaAATP to
DnaAADP by RIDA or DDAH, or the sequestration of the oriC by SeqA, is likely more favorable than
targeting the rejuvenation of DnaAATP. Obstructing SeqA binding to the GATC in the oriC and the DnaA
promoter region, would likely lead to over-initiation and increased levels of DnaAATP. However, the
increase in initiation observed when seqA is deleted is not detrimental to the cell (271). DDAH is less
efficient in stimulating DnaAATP hydrolysis than RIDA (170), indicating that RIDA is the favorable target.
RIDA inactivation could be achieved by; i) hindering binding of ADP to Hda, ii) blocking the ATPase
stimulatory interaction of the Hda Arg-finger with the DnaAATP ATPase in domain III, iii) inhibiting the
stabilizing interactions between the C-terminal of Hda and DnaAATP domain I. Though the idea of
inducing over-initiation is intriguing, the fact that secondary mutations arise quickly in cells that are
over-initiating (240), designates that resistance would quickly develop. Furthermore, the ROS
dependency of the lethal action of hyper-replication suggests, that the strategy of inducing over-
initiation is not plausible at low ROS conditions i.e. anaerobsis or when free iron availability is limited.
Inhibition of the regeneration of DnaAATP via DARS1 or DARS2, is likely achievable by
blocking the DnaA-DnaA interactions in the DnaA box core region, or by hindering the binding of IHF or
Fis to the regulatory region of DARS2. The mechanisms that lead to DnaAATP regeneration mediated by
phospholipids are unknown. Consequently, inhibition of the phospholipid synthesis is the only known
mean by which this process could be inhibited. Targeting the regulatory mechanisms of the replication
initiation has a downside, as it currently not known how many bacterial species that uses the DnaAATP
level to regulate replication initiation. Hda homologs have been identified in Caulobacter and most
enterobacteria, but not in Bacillus, Staphylococcus and H. pylori (131, 272). DARS1 and DARS2 are likely
conserved in proteobacteria that are closely related to E. coli, while more distantly related
proteobacteria have DARS-like DnaA box clusters in other intergenic regions. Non-proteobacterial
species like S. aureus, Mycobacterium tuberculosis and Bacillus subtilis have DnaA box clusters near
dnaA, though with a different arrangement of the DnaA boxes than the one in E. coli (148).
Consequently, compounds that target the regulation of replication initiation will likely not have a broad
bacterial spectrum.
105
Despite all of the above mentioned potential targets, it is curios that not a single
compound that targets the replication initiation process has been identified. This fact could indicate that
the replication initiation is not an ideal target for development of novel antibiotics. Furthermore, the
complexity of its regulation may enhance the occurrence of secondary compensatory mutations that
counteract the inhibitory action of a given compound. Yet, it is important to keep in mind that direct
efforts at identifying replication initiation inhibiters has so far been scarce.
The screens presented in paper II and III have some distinct differences concerning their
specificity and practical applicability. The fluorescence-based screen relies on replication by cSDR and is
therefore not able to identify putative inhibitors of the DnaB helicase loading; as such compounds would
kill the cells. Conversely, the screens based on over-initiating cells cannot be used to identify inhibitors
that induce over-initiation. Whereas, over-initiation of the mini-chromosome in the fluorescence based
screen, could potentially lead to instability and loss of the mini-chromosome and thereby expression of
the green fluorescent protein (GFP). Concerning the practical applicability, the fluorescence based
screen is performed in 96-well plates in a high throughput manner. In contrast, the agar plate based
platform of the hda and DARS2 screens needs to undergo further optimization for use in a high
throughput setup. The agar plate based platform has a significant advantage, as a concentration
gradient is created when the extract or compound diffuse from the well into the agar. Thus, several drug
concentrations are tested at once, in contrast to the fluorescence based screen, where only a single
concentration is tested.
Why is severe over-initiation of the DNA replication lethal?
In paper II, we show that the iron chelator deferoxamine rescues the growth of over-initiating cells by
promoting the processivity of replication forks. Deferoxamine has been shown to inhibit the Fenton
reaction and thereby the production of ROS both in vivo and in vitro. Our findings therefore support the
proposed model that the lethal action of over-initiating the chromosomal DNA replication is caused by
formation of DSBs in the DNA, when replication forks encounters 8-oxo-dGTP lesions that are under
repair by the GO system (235). Since overproduction of ribonucleotide reductase (RNR), restores the
growth of over-initiating cells, it has been suggested that during replication over-initiation the cells are
starved for dNTPs because of the increased number of replication forks (238, 239, 241). The dNTP
starvation is proposedly responsible for the severe growth retardation of cells deficient in Hda (239).
However, several observations contradict this model; i) the reduction in the dNTP pool of the hda
mutant is not significant, ii) the inviability of an hda mutant does not resemble the inviability observed
for cells starved in dNTPs, as dNTP starvation leads to obliteration of the oriC, in contrast to the
observed increase in oriC copy number for hda mutants, iii) an increase in all four dNTPs is not observed
106
when RNR is overexpressed, specifically the dGTP level remains more or less unchanged, thus a hda
mutant overexpressing RNR is still starved in dGTP (271). Conversely, the observed suppression of the
hda phenotype by overproduction of RNR is more likely caused by a reduction in replication initiation, as
hda mutants overproducing RNR has an origin concentration resembling that of a wild-type (241, 271).
Hence, over-expression of RNR in an hda mutant lowers the initiation frequency, giving time for efficient
repair of 8-oxo-dGTP lesions.
Conclusions
Utilizing genetic screens and differential gene-expression analysis, we have shown that reversing
ciprofloxacin resistance in a high-level ciprofloxacin resistant E. coli strain is likely not possible. However,
our genetic screen revealed the AcrAB-tolC efflux pump, and the SOS response proteins RecA and RecC,
as plausible targets for ciprofloxacin helper drugs in E. coli strains with a MIC just above clinical
breakpoint.
We have constructed and verified three screens for identifying inhibitors of the initiation
of chromosomal DNA replication in bacteria. One screen relies on replication inhibition of an OriC
dependent mini-chromosome, leading to expression of GFP and thereby a detectable increase in
fluorescence. The two other screens are based on growth rescue of cells that exerts lethal over-initiation
of the DNA replication, by either harboring multiple copies of DARS2 or being deficient in Hda.
As a pilot screen for inhibitors of the replication initiation process, we subjected our novel
screens to a library of 400 actinomycetes extracts. Even though we did not identify any initiation
inhibitors, the iron chelator deferoxamine, a known inhibitor of the Fenton reaction (245), was
identified as a compound that rescues the growth of over-initiating cells. Corroborating the model that
the lethality of over-initiating the chromosomal DNA replication is caused by formation of DSBs in the
DNA, when replication forks encounters 8-oxo-dGTP lesions that are under repair by the GO system
(235). In addition, we also showed that the growth rescue of over-initiating cells exerted by the
suggested gyrase inhibitor (±)-6-Chloro-PB hydrobromide (S143) is, at least in part, due to its ability to
chelate iron.
Future perspectives
Antibiotic resistance will be a remaining threat to global health care. However, by heavily investing in
antibiotic drug development and regulating the use of antibiotics globally. It may be possible to halt or
slow the current negative development. Based on our findings that the AcrAB-TolC efflux pump and the
107
SOS response genes RecA and RecC, might serve as ciprofloxacin helper drug targets, in treating low-
level resistant strains. In addition to the fact that such helper drugs have the potential to potentiate
other known antibiotics and decrease the evolution of antibiotic resistance, it would be interesting to
setup screens for identifying inhibitors of these three targets. Moreover, it would be attractive to assess
if PriA deletion is truly lethal for bacterial strains carrying gyrA mutations that confer ciprofloxacin
resistance.
As we now have two distinct strategies for identifying replication inhibitors, the next
logical step is to obtain several chemical or natural extract libraries that could be subjected to the
screens. The 400 extracts that were screened in paper II and III, are part of a large library of more than
4000 microbial extracts, owned by our collaborator Naicons srl., which has proven to be a source of
novel antibiotic compounds (273, 274). Hopefully, future funding will give us the opportunity to screen
the remainder of this vast library of bioactive natural extracts.
108
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