Regulation of DNA replication and cellular responses to
perturbations in replication in the bacterium Bacillus subtilis
by
Alexi I. Goranov
Bachelor of Science in Biology
University of Texas at Dallas, 2001
Submitted to the Department of Biology in Partial Fulfillment of the Requirements for
the Degree of Doctor of Philosophy in Biology
at the Massachusetts Institute of Technology
September, 2006
© Massachusetts Institute of Technology
All rights reserved
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Department of Biology
September 1, 2006
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SAccepted
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A ccepted by ..........
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.................................
Stephen P. Bell
Professor of Biology
Chair, Biology Graduate Committee
IN-W E
MASSACH-USElS
OF TECHNOLOGY
SEP 13 2006
LIBRARIES
.... ..........
Alan D. Grossman
Professor of Biology
Thesis Advisor
ARCHIVES
Regulation of DNA replication and cellular responses to perturbations in replication in
the bacterium Bacillus subtilis
by
Alexi I. Goranov
Abstract
When a cell grows and divides to give rise to genetically identical cells, the genome of
the cell is duplicated prior to cell-division. The process of genomic duplication is called
DNA replication, and is closely coordinated with other processes in the cell, such as
growth rate, and cell division. The mechanisms that regulate when DNA replication
initiates and how cells respond to perturbations in replication are not well understood. I
used the gram-positive bacterium Bacillus subtilis to address these questions. My
research showed that a conserved component of the DNA replication machinery,
processivity P-clamp, regulates the initiation of replication. This regulation appears to
affect the loading of helicase, a replication component that generates the single-strand
DNA template for replication. My results indicate that the replication initiation protein
DnaA is the likely target of P-clamp regulation. I also observed that in vivo, in B. subtilis,
most of the DNA replication machinery, including P-clamp, can associate with the origin
of replication before helicase. This is in stark contrast to in vitro studies in other bacteria.
I also addressed the question of how B. subtilis responds to perturbations in DNA
replication and DNA damage. My results demonstrate that the conserved recombination
protein, RecA, mediates most of the transcriptional response under the tested conditions.
More than 75% of the RecA-mediated transcriptional response is due to the expression of
phage and mobile element genes and their indirect effects. Under conditions of
replication elongation arrest, there is still a significant recA-independent response, at least
part of which is mediated by the replication protein DnaA. The DnaA-mediated response
appears to be conserved in other bacteria, as homologues if the affected genes also have
DnaA binding sites in their promoter regions. Previously, one of the DnaA regulated
genes, sda, has been shown to affect cell viability after perturbations in replication. Here
I showed that another DnaA-regulated gene,ftsL, also affects cell survival after
replication arrest by coordinating replication and cell-division.
I believe that my results have furthered our understanding of how replication is
coordinated with other cell-cycle processes, and have raised interesting questions for
future investigation.
Thesis Advisor: Alan D. Grossman
Title: Professor of Biology
Acknowledgements
I wish to thank my advisor, Dr Alan Grossman, for his dedication, support,
encouragement, and intellectual contributions to this work. It has been a terrific
experience working and learning from him. His emphasis on student education has
helped me form a better understanding of how to think and communicate about science.
I also thank past and present members of the Grossman lab who made my stay there
enjoyable and productive. I thank them for their support and critical and stimulating
discussions of my work and science in general. In particular I thank C. Lee for reading
and commenting on virtually everything I have written, for technical help, and for
making life in the lab easy. I thank M. Berkmen for the numerable constructs and strains
she has made that have allowed me to progress in my studies faster and more efficiently,
and for her discussions, comments, and help. I also thank J. Auchtung, J. Wang, E.
Kuester-Schoeck, and A. Breier for much technical help and for reading and commenting
on my manuscripts. I also thank M. Rokop and W. Burkholder for introducing me to
DNA replication and for helping me start my project.
I also thank my thesis committee members, T. Baker, S. Bell, F. Solomon, and A. Wright,
for their stimulating discussions and helpful comments on my work in general, as well as
their comments on my manuscripts.
I also thank members of the Baker, Bell, Endy, Kaiser, Sauer, and Walker labs for help,
reagents, and for allowing me to use their equipment.
I also thank my family for their support, and especially my wife for her love.
Table of contents
Abstract.............................................................................
page
........................................... 2
A cknow ledgem ents................................................
......................................................
3
Table of Contents...................................................
......................................................
4
L ist of T ables.......................................................
.........................................................
5
L ist of Figures.....................................................
..........................................................
6
Chapter 1:
Chapter 2:
Chapter 3:
Chapter 4:
Introduction.........................................
................................................
8
A transcriptional response to replication status mediated by the
conserved bacterial replication protein DnaA ....................................
49
Characterization of the global transcriptional responses to
different types of DNA damage and disruption of replication
in Bacillus subtilis .............................................
........
94
Early association of replisome components with the origin
of replication provides a mechanism for regulating the initiation
of DNA replication .....................................
140
Appendix A:
Supplementary Materials and Methods for Chapter 4......................... 161
Chapter 5:
P-clamp and YabA regulate the initiation of DNA replication
in Bacillus subtilis .....................................
C hapter 6:
D iscussion.................................................
172
........................................ 203
List of Tables
page
Chapter 1:
Table 1.
Components of the replication machinery in B. subtilis.... 12
Chapter 2:
Table 1.
B. subtilis strains used ......................................
Table 2.
Effects of inhibiting DNA replication on gene
expression .................................... ............
Chapter 3:
.... 54
66
Table 3.
Conservation of DnaA binding sites in other bacteria........84
Table 1.
B. subtilis strains used...................................
Table 2.
DNA damage and perturbations in replication cause
changes in expression of many genes............................ 108
Table 3.
MMC and UV treatments affect a few of the genes
proposed to be regulated by DnaA.....................................13
Table 4.
LexA appears to repress approximately 63 genes in 26
operons ............................ ..........
.........
Table 5.
..... 100
117
Phage induction affects the expression of many
non-phage genes......................................... 121
Appendix A: Table 1.
B. subtilis strains used.................................
...... 169
Chapter 5:
B. subtilis strains used.................................
180
Table 1.
List of Figures
Chapter 1
Chapter 2
Figure
Replisome structure in bacteria..............................
14
Figure
Helicase loading in B. subtilis ....................................
18
Figure
Origin structure in Gram + and Gram- bacteria...........21
Figure
Initiation of replication in B. subtilis ...........................
Figure
DnaA inactivation by Hda and P-clamp in E. coli..........28
Figure
DnaA-ATP fluctuation during the cell cycle
affects the ability of DnaA to regulate replication
in E.coli ...............................................
.......................... 31
Figure 1
Changes in mRNA levels in response to inhibition of
replication......................... .................
Chapter 4
25
62
Figure
DnaA appears to directly regulate at least 20 operons......73
Figure
Regulation offtsL contributes to the inhibition of cell
division...............................................
77
Conservation of potential DnaA binding sites in other
bacteria ...............................................
80
Figure 4
Chapter 3
page
Figure 1
DNA damage and replication arrest affect the
expression of large number of genes in a
recA-dependent manner................................................105
Figure 2
MMC treatment causes a relative increase in origin
proximal gene expression and gene dosage..................129
Figure
Effects of MMC and HPUra on DNA replication............32
Figure
Use of Chromatin Immunoprecipitation (ChlIP) to
assay the association of replication proteins with
the origin of replication..............................144
Figure 2
Association of replisome components with the
origin does not require helicase...............................
Figure 3
Overproduction of P-clamp (DnaN) stimulates DNA
replication in an oriC-dependent manner and affects the
association of helicase and helicase loader (DnaB) with
148
the origin, but not the association of DnaA..................1.... 52
Figure 4
Appendix A: Figure 1
Chapter 5:
Figure 1
Figure 2
Model for the order of association of replisome
components to the origin of replication in B.
subtilis..............................................156
Association of clamp loader/z-subunit (DnaX) and
helicase (DnaC) with the origin of initiation in
dnaD23ts cells..............................................................
Effects of P-clamp on the initiation of DNA
replication are likely through YabA ...............................
DnaA and P-clamp overexpression have additive
effects on DNA replication.........................
167
83
186
Figure 3
GFP-YabA focus formation does not depend on
DnaA or DnaA-dependent replication and is
disturbed replication arrest ............................................. 89
Figure 4
yabA and dnaNdo not affect the expression of
DnaA-regulated genes................................
..... 192
Chapter 1
Introduction
During each cell division cycle, cells grow, replicate their genome, and divide to
make two cells. It is a fundamental biological question as to how these events are
coordinated with each other. DNA replication, the process through which the genome is
duplicated, is central to cellular propagation, and its proper regulation is essential for cell
cycle coordination. Replication is affected by growth rate, such that the faster cells grow,
the more often they replicate their genome. In prokaryotic as well as eukaryotic cells,
replication initiates when cells have reached a defined size, which varies with growth
condition (44, 51, 52, 83, 109). How growth rate and other external stimuli regulate
replication initiation is not well understood. What is becoming increasingly apparent,
however, is that if the rate of replication initiation is mis-regulated, cells lose viability
and produce progeny with incomplete genomes, thus indicating that proper regulation of
replication is essential to maintaining genome stability (24, 127, 143).
DNA replication can be separated into three temporal steps: initiation, elongation,
and termination. During initiation, specific regions on the chromosome(s) are recognized
by specialized proteins (99), which load the replication machinery onto the DNA
template. During elongation, the most time consuming step of replication, the genome is
duplicated by the replication machinery (3, 50, 66). The last step is termination when the
opposing replication forks meet and are resolved, resulting in two completed and
unlinked chromosomes (138-140).
Although much is known about the different biochemical steps required for the
initiation, elongation and termination of replication, the regulation of replication is not
well understood. In the gram-positive bacterium Bacillus subtilis, essential proteins
involved in DNA replication were identified through classical genetic methods and as
homologues of replication proteins from other bacteria (53, 69, 102). I used B. subtilis as
a model organism to gain further understanding of how the initiation of DNA replication
is regulated, and how replication regulates other cellular processes. A summary of
replication events relevant to this thesis are introduced below, followed by a brief
summary of the findings presented in my thesis.
Replication machinery in B. subtilis. The genome is duplicated during the
elongation step of DNA replication, which is mediated by a complex of proteins
generally called the replisome (50, 75). In B. subtilis, the replisome is comprised of about
10 different proteins all of which are essential (Table 1, Figure 1) (63, 77). The enzymatic
activities of the replisome are conserved among divergent bacterial species (3, 50, 66).
The replisome contains the DNA polymerase holoenzyme, the replicative helicase,
primase, and SSB.
In B. subtilis and some other gram positive bacteria, the DNA polymerase
holoenzyme includes a major DNA polymerase (PolC), another essential DNA
polymerase (DnaE), dimerization c-subunit and y-clamp loader (DnaX), processivity f3clamp (DnaN), 6-clamp loader (HolA), and 6'-clamp loader (HolB) (13, 29, 63, 77, 102).
B. subtilis does not encode X-and xy-subunits (part of the clamp loading complex in E.
coli) or the e-and 6-subunits (3' to 5' exo-nucleolytic activity in E. coli) (69). In B.
subtilis, PolC encodes 3' to 5' exo-nucleolytic activity (77, 123, 124). The clamp loader 8
subunit is also very different from that in E. coli, either containing or no longer requiring
the functions of the E. coli X-and V-subunits (13, 102). B. subtilis like most other bacteria,
and unlike E. coli, encodes two different polymerases of the DnaE family, PolC and
DnaE (12, 29, 69, 72, 116). Although DnaE is essential for viability, its function is
unclear. Initial evidence suggested that DnaE may be the lagging strand DNA polymerase
(29), but biochemical studies with purified DnaE from B. subtilis and Streptococcus
pyogenes showed that DnaE is relatively slow in DNA synthesis (-60 nt/ sec as compared
to > 800nt/sec for PolC) (12), and introduces mismatches at a very high rate (12, 29, 72),
suggesting that DnaE may not be a major DNA polymerase, and that PolC replicates the
bulk of the chromosome.
DNA polymerase stays on the DNA template without dissociating for thousands of
base pairs through its interaction with the P-clamp, which is topologically bound to the
DNA (3, 27, 50, 66). P-clamp is a dimer in bacteria and forms a doughnut shaped
structure which encircles the DNA (46, 50, 64, 105). DNA polymerase interacts with the
P-clamp and thus remains tethered to the DNA template (27, 79, 84, 85). The P-clamp
loading machinery (8-, 8'-, and y -subunits) loads P-clamp onto the DNA template (28,
50, 110). Structural analysis of P-clamp and clamp loader from E. coli and eukaryotes
have provided a model where the clamp loader catalyzes the loading of P-clamp at the
junction of ssDNA and an RNA primer synthesized by primase (see below) (1, 9, 50). In
B. subtilis, the dnaX gene makes a single protein, t-subunit (13, 77), unlike dnaX in E.
coli, where it generates the t-subunit and the shorter y -subunit (66). Thus in B. subtilis,
S. pyogenes, and likely many other bacteria, the T-subunit of the replisome functions as a
component of the clamp loader (13, 77). Another essential function of the --subunit,
thought to be shared between T-subunits of all bacterial DNA polymerases, is that it
serves to keep together the leading and lagging strand DNA polymerases (3, 50) (Figure
1). The DNA polymerase holoenzyme has been reconstituted in vitro from only one
gram-positive bacterium, S. pyogenes. The stoichiometry of monomeric subunits at a
Table 1. Components of the replication machinery in B. subtilis
Gene
dnaA
dnaB
dnaD
dnal
dnaC
dnaG
ssb
polC
dnaE
dnaN
dnaX
Encoded Protein Function
Initiation factor. Recognition and melting of origin of replication.
AAA+ protein. Transcription factor
Initiation factor. Helicase loading. Membrane association f origin
Initiation factor. Helicase loading
Initiation factor. Helicase loading. AAA+ protein
Helicase
Primase
Single-stranded DNA binding protein
DNA polymerase III. DNA polymerase catalytic a-subunit. Includes 5'3' exo activity
DNA polymerase III. Essential DNA polymerase. Unclear function
P-processivity clamp
r-subunit. p-clamp loading. Coupling of leading and lagging strand.
AAA+ protein
holA
holB
6-subunit. p-clamp loading
6'-subunit. P-clamp loading
replication fork was determined to be PolC 2 14 61 6'1
4 (13).
The structure and relevant
interactions of bacterial replication components are shown in Figure 1.
The replisome also contains helicase and primase. The function of primase (DnaG
in B. subtilis) is to synthesize a short (-6nt) RNA primer from which DNA polymerase
begins to replicate the chromosome (3, 21, 50, 66). Primase is needed repeatedly at the
lagging strand, where DNA polymerase needs to synthesize a new fragment of DNA
every -1000bp (3, 66, 73). PolC extends the RNA primer in E. coli (66), however the
ability of PoIC to begin synthesis from an RNA primer has not been demonstrated in B.
subtilis. Helicase (DnaC in B. subtilis) is a hexameric, ATP-driven machine, which
unwinds dsDNA to expose ssDNA that serves as a template for replication. Helicase in B.
subtilis moves 5' to 3' on the lagging DNA strand to cause DNA unwinding (137).
The activity of primase is modulated by its interactions with helicase (21). Primase
initiates primer synthesis more often and synthesizes shorter primers when interacting
with helicase (21). Helicase function, on the other hand, is affected by its interactions
with the DNA polymerase holoenzyme, the t-subunit in particular. In E. coli, when
helicase interacts with t, it unwinds DNA at a higher speed (3, 61).
Interestingly, the replisome forms a "factory", a conglomerate of many replication
proteins carrying out DNA synthesis, at a specific position in the cell (5, 49, 75, 76, 78).
In B. subtilis, when replisome components are fused to GFP under slow growing
conditions, they form foci that localize to the middle 25% of cell length (75). These
observations indicate that during replication the replicated DNA is moved through the
replisome, as opposed to the replisome moving on the DNA to replicate it. This
observation of visible foci also suggested that there are many (> 40 molecules) of the
helicase
'/
primase
y/'t
complex
¥
~
•
\
PoilU core
~)
PoiClDneE?
I
Figure 1
14
Figure 1. Replisome structure in bacteria. The components of one of the replication
forks are shown. The replisome is thought to contain two such complexes to form the
clock-wise and counter-clock-wise replication forks. The hexameric helicase (DnaC in B.
subtilis) is shown in purple, moving in the 5' to 3' direction. Helicase interacts with
primase (DnaG), shown in blue, and T-subunit of DNA polymerase shown in green.
Primase is shown synthesizing a short RNA primer, in pink, on the lagging strand. One
DNA polymerase (PolC or DnaE) is shown, yellow and red, on leading and lagging
strand. The polymerases are tethered to the DNA through interactions with the P-clamp,
red. P-clamp is loaded by the clamp loader (green) containing the --, 6-, and 6'- subunits.
Although the figure shows only two T-subunits, interacting with DNA polymerase and
helicase, there are estimated four T-subunits per two PolC DNA polymerases. T-subunits
in B. subtilis also function as the y-components of the clamp loader. Although in E. coli,
y- and '-subunits are derived form the same gene, dnaX, through translational slippage,
such that the '-subunit contains an extra C-terminal region, in B. subtilis and other
bacteria, dnaXappears to produce a single peptide which functions as both the y- and zsubunit (see text for details). SSB tetramers shown in cyan cover the ssDNA exposed
between Okazaki fragments on the lagging strand. Adapted from Johnson and O'Donnell.
Ann. Rev. Biochem. 2005.
replisome components present in the "factory" during replication, which is many more
than expected for a single DNA polymerase holoenzyme at each replication fork. The
significance of this "repository" for replication components is not yet understood,
although the foci only form once replication is initiated (75).
The eukaryotic replisome is comprised of more protein subunits than the bacterial
replisome, but many enzymatic properties are common in both bacteria and eukaryotes
(3, 8, 50). The processivity P-clamp is encoded by PCNA. Although P-clamp is a trimer
in eukaryotes, the domain structure of the assembled trimer is the same as in the bacterial
dimer (46, 59). The RFC complex functions analogously to the z-complex in bacteria and
acts as a clamp loader, and its subunits function analogously to the bacterial clamp loader
(28, 50). The eukaryotic helicase is composed of at least six peptides (MCM2-7) and
likely other associated components (8). The primase, Pola, contains both RNA and DNA
synthesis activities (3, 8). Since eukaryotic polymerase is unable to start synthesis from
an RNA primer, Pola first synthesizes a short RNA primer and then synthesizes a short
DNA primer (50). There are two Pol III type DNA polymerases (major DNA
polymerases) in eukaryotes, Pol8 and Pole (8, 34). Po16 is thought to be the DNA
polymerase at the leading strand and Pole is thought to be the lagging strand DNA
polymerase (8). There is however some controversy regarding the function of Pole.
Although point mutations in Pole can be lethal (100), at least in Saccharomyces
cerevisiae, strains carrying a deletion of Pole polymerase are viable (8), thus indicating
that, much like DnaE in B. subtilis, Pole may not be a major DNA polymerase.
Eukaryotes also have a tetrameric single strand DNA binding protein, RPA (8, 50).
Helicase loading. Helicase in B. subtilis (DnaC) is a monomer in solution and
forms hexamers when loaded onto DNA (137). Helicase loading occurs either during the
initiation of DNA replication at the origin, or after DNA damage repair, which could
happen anywhere on the chromosome (10, 35, 66, 90, 113, 117, 129). In both cases
helicase is loaded by three proteins: DnaI, DnaD, and DnaB (10, 35, 90, 113, 117). DnaB
and DnaD are conserved among low-GC gram-positive bacteria, while DnaI is
homologous to the E. coli helicase loader DnaC (117, 129). These three proteins alone
cannot load helicase onto DNA, as they need to be recruited to the origin, or to the
repaired replication fork, by DnaA or PriA, respectively (10, 48, 90). PriA is needed for
replication restart after replication forks encounter blocks (23, 24, 89, 122), and DnaA is
the replication initiation protein in bacteria (93), as is discussed below. The order of
assembly is delineated from genetic experiments showing that gain-of-function mutations
in DnaB suppress the deletion of PriA (117), that DnaA and DnaD interact directly (48),
and that mutations in DnaB and DnaD prevent helicase association with the origin of
replication in vivo (117). Furthermore, in vitro studies have shown that PriA stimulates
the association of DnaD to a forked template, and that DnaB and DnaD mediate helicase
loading (11, 90). Our current understanding is that DnaA or PriA bind a specific DNA
structure and recruit DnaD, which then recruits DnaB and Dnal, which then load helicase
(Figure 2).
DnaB and DnaD alone posses weak ssDNA and dsDNA binding activity (11, 129).
Although binding of DnaB and DnaD to forked DNA is stimulated by PriA (90),
mutations in dnaB, such as dnaB371, that suppress the lack of PriA (117), increase the
affinity of DnaB for ssDNA (11). Such DnaB mutations also recruit DnaD to the ssDNA
1
Forked DNA
template
Helicase (DnaC)
Figure 2
18
Figure. 2 Helicase loading in B. subtilis. In B. subtilis, helicase (DnaC) is loaded at a
forked DNA structure by the action of three proteins, DnaI, DnaD, and DnaB. These
three proteins are recruited to the proper DNA structure by the actions of PriA (blue), and
likely DnaA. PriA (or DnaA) recruit DnaD (yellow) to the forked structure by a direct
interaction, and DnaD recruits DnaB (green). DnaB and DnaI (purple) load the hexameric
helicase (red) from DnaC monomers present in solution. In E. coli, there are no DnaD
and DnaB homologues, but there is a DnaI homologue, called DnaC in E. coli.
even without PriA present (11, 117). These observations led to the hypothesis that these
gain-of-function mutations of DnaB can bind ssDNA at repaired replication forks and
mediate helicase loading without PriA. DnaD is still required in such gain-of function
mutants of DnaB. These DnaB mutants can also suppress defects in DnaD function (117).
In E. coli, which has no DnaD and DnaB homologues, DnaA interacts directly with
helicase (33, 91, 135). DnaA mutants have been isolated that are unable to interact with
helicase, while maintaining all other tested biochemical activities (33). Such mutants
were unable to initiate DNA replication, indicating that DnaA plays an important role in
helicase loading (33).
Initiation of DNA replication in B. subtilis. The genome of B. subtilis consists of a
single circular chromosome, approximately 4.2 Mb in size (69). The chromosome is
replicated from a single origin of replication, called oriC, located at 00 (77, 98, 99). The
origin of replication was isolated by classical genetic methods as a piece of chromosomal
DNA that allowed an E. coli plasmid to replicate in B. subtilis (98). The minimal origin
of replication spans - 2 Kb and includes 500bp upstream of the dnaA locus, the dnaA
ORF, and the 200 bp dnaA-dnaN intergenic region (77, 98, 99, 106). The location of the
origin of replication proximal to the dnaA locus is a characteristic conserved among
many bacteria, although there are exceptions, E. coli being the most well studied example
(Figure 3). The proximity of the origin of replication and the dnaA locus is consistent
with their co-evolution, as DnaA is the replication initiation protein that recognizes and
acts upon the origin to initiate replication. DnaA is a highly conserved protein that
belongs to the AAA+ class of enzymes (28, 93, 94, 128). DnaA binds DNA in a sequence
<Gram-positive bacteria>
Bacillus subtilis
rpmH
DnaA boxes
p
jet1
-4
.... bubd4[4
DnaA boxes
dnaN
dnaA
Mi
d&---hI
b'
I
I
'...io '
oriC
te
Initiation site
dnaNV
idbk'
bd~h dnaA
Spiroplasmacitri
r---\I
I
oriC
Streptomyces lividans
dnaA
rpmH
,I'--1
---
dnaN
b Ir--bbdbb
... ------
r~----mmk -
)
I
b
-
---
1- .
bbd d
--r------~r---------···-
-···············-············--- r .
1-
oriC
Mycobacterium smegmatis
rpmH
dnaN
dnaA
WM
))
.
I
i
oriC
<Gram-negative bacteria>
Escherichia coli
Pg~dA
L
I
oric
rpmH
-
c....
Ip
---- ---
•
---
---
rpmH P
I
Caulobactercrescentus
hemE,
•
dnaA
• t, r-dnaN
-
ca 40kbp
Pseudomonas
putida
•
rP
•
oriC
•
•
dnaA dnaN
P
I1
.. 4bb*r
ora"c
oriC
I
m
dnaoA
•
Jr_• __]LY
dnapN
•
ca 2kbp
Figure 3
ca 200kbp
Figure 3. Origin structure in Gram + and Gram- bacteria. A schematic representation
of the origin of DNA replication of several bacterial chromosomes is shown. Genes are
labeled on top and direction of transcription of a gene is indicated by the direction of the
arrow representing the gene. DnaA is shown in black. DnaA binding sites are shown as
arrowheads with filled arrowheads being consensus DnaA binding sites. The minimum
origin region is underlined. A thin arrow followed by "P" indicates mapped promoters
near the origin. Discontinuities in the diagram are indicated by a dotted line and the
distance skipped is indicated underneath.
The one underlining feature is the presence of DnaA binding sites near the origin of
replication, and, in Gram+ bacteria, the proximity of DnaA to the origin. In most
bacteria, dnaA and dnaN form an operon. Adapted from Moriya, Imai, et al. Plasmid.
1999.
specific manner, serving to initiate replication and to also regulate gene expression at
many loci (14, 95, 98).
There are DnaA binding sites (consensus TTa/tTnCACA, n=A, G, T, or C)
upstream and downstream the dnaA ORF (98). Estimates from E. coli suggest that
between 6 and 20 DnaA molecules bind at the origin (18, 66), but it is not known how
many DnaA molecules bind to the origin in B. subtilis. Within the dnaA-dnaN intergenic
region, close to the downstream DnaA binding sites, there is a 27 bp A-T rich region. The
DnaA molecules at both upstream and downstream binding sites are thought to interact
with each other, causing a bend in the DNA and also likely creating torsion, which melts
the A-T rich region and exposing single-strand DNA (ssDNA) (Figure 4) (67, 68, 77, 99).
Melting of the A-T rich region, but not DnaA binding, requires that the DNA molecule be
supercoiled (38, 39, 67, 68). This requirement may explain why inhibition of
transcription, which affects DNA supercoiling, abolishes the initiation of DNA
replication (6, 71). In vitro, the region melted by DnaA, in both E. coli and B. subtilis, is
stabilized and extended by single stranded DNA binding protein (SSB) to about 35bp (4,
41, 67, 68). In vivo experiments in E. coli, utilizing a plasmid carrying the origin of
replication, reveal that about 35bp are melted as well (41). It is worth noting, however,
that the initiation of DNA replication from the same origin differs in requirements
depending on whether the origin is on a plasmid or on the chromosome. It is still
unknown how much of the origin is melted during initiation within the context of the
chromosome. The region melted by DnaA serves to recruit other replication proteins
needed to begin the elongation step of replication (discussed in next section).
Although, in vitro, DnaA alone can melt the origin, experiments from E. coli have
demonstrated that many other proteins aid DnaA in origin melting. The sequence specific
DNA binding proteins Fis and IHF, and the non-specific DNA binding proteins HU and
SSB, and near-by promoter activity have all been demonstrated to affect origin melting
by DnaA in E. coli (66-68, 120). Homologues of Fis and IHF are not present in B.
subtilis, but HU and SSB are thought to be important during initiation (68). The role of
near-by promoter activity in regulating replication initiation in B. subtilis remains
unexplored.
Regulation of the initiation of DNA replication in bacteria: Lessons from E. coli. It
is of major interest to understand how DnaA is regulated so that replication initiates only
at the appropriate time during the cell cycle and so that the rate of replication initiation is
coordinated with growth rate. Many studies in E. coli have demonstrated that DnaA is
regulated at the level of expression, function, and protein sequestration. Inactivation of
any of the layers of regulation results in perturbations in the timing of initiation. The
expression of dnaA is auto-regulated such that DnaA represses its own promoter. In
addition, the expression of dnaA is repressed during parts of the cell-cycle by occlusion
of the dnaA locus by SeqA, which binds methylated and hemi-methylated DNA (104).
DnaA molecules are also sequestered by the datA locus on the chromosome which has
the capacity to bind many DnaA molecules, although there are only two recognizable
DnaA binding sites at that locus (30, 97, 108).
In addition to the regulation of DnaA abundance, DnaA function is regulated by its
nucleotide bound state. DnaA binds ATP or ADP and can hydrolyze, albeit slowly, ATP
to ADP (126). The ATP-bound form of DnaA is functional for the initiation of DNA
DnaA boxes AT
DnaA boxes
o
o
o
0 0
000
oDnaB
DnaI complex
ODnaA
IOnaD
AT
DnaC helicase-
o
ftA
UV
Figure 4
25
Figure 4. Initiation of replication in B. subtilis. Regions with DnaA boxes are
presented as gray boxes surrounding the dnaA ORF, a white rectangle, and the AT-rich
region is shown as a black box. DnaA, white circles, binds to the DnaA boxes flanking
the dnaA ORF and causes a bending of the DNA such that DnaA molecules on each side
of the dnaA ORF can interact with each other. Possibly with the help of other proteins
and transcription, DnaA melts the AT-rich region, exposing ssDNA. The exposed ssDNA
serves as a landing pad for other replication proteins. Adapted from Moriya, Imai, et al.
Plasmid. 1999.
replication and also for transcriptional control (43, 92, 126, 130). In vitro experiments
have suggested that ATP hydrolysis by DnaA is not required for initiation of DNA
replication (126). The ATPase function of DnaA is regulated and the ATP/ADP bound
form of DnaA varies during the cell cycle. The ratio of ATP/ADP bound DnaA is 0.8
before replication initiates, and decreases to 0.2 after replication initiates (70). The
ATPase function of DnaA is activated by a protein homologous to DnaA (Hda) in
complex with P-clamp loaded onto DNA (Figure 5) (55, 56, 58, 101, 111, 131, 132). The
evidence suggests that during the elongation step of DNA replication, Hda interacts with
the P-clamp and inactivates DnaA by stimulating its ATPase activity (15, 16, 56, 58, 101,
111, 131, 132). In this context P--clamp is also a negative regulator of replication as it
stimulates Hda, although overexpression of 3-clamp does not affect replication in E. coli
(Lyle Simmons, personal communication).
Recent studies have shown that while both DnaA-ATP and DnaA-ADP bind the
DnaA consensus box with high affinity, the DnaA-ATP form can bind to additional DNA
sequences, which differ slightly from the consensus (92, 130). The binding to these extra
DnaA binding sites is thought to allow DnaA to form a larger or more extensive nucleoprotein complex that is needed to melt the origin region (93, 94). DnaA mutants that are
unable to hydrolyze ATP, but still bind ATP or ADP, are hyper-active for replication
initiation (101). Other DnaA mutants, such as DnaAcos, can be hyper-active for DNA
replication initiation, even though they have lost their ability to bind nucleotides (54,
127). The sum of all data suggests that ATP binding alters the structure of the protein to
allow it to bind the origin of replication in a specific manner, which leads to origin
melting. Although, the de novo synthesis of DnaA is generating DnaA-ATP, acidic
~-clamp
ATP~
ADP~
ATP~~
~-clamp on DNA
Hda --I
--
Figure 5
28
DnaA
Figure 5. DnaA inactivation by Hda and P-clamp in E. coli. The regulation of DnaA
in E. coli is diagramed. p-clamp associated with DNA enables Hda to interact with DnaA
and to stimulate the intrinsic ATPase activity of DnaA, thus converting ATP-DnaA into
the inactive ADP-DnaA. DnaA is also thought to interact with the DNA molecule. The
genetic formalism of the regulation of DnaA is shown below the diagram.
phospholipids in the cellular membrane have been implicated in aiding the exchange of
ADP for ATP by DnaA (19, 25, 26, 40, 62, 80, 142). These conclusions were supported
by in vitro studies, as well as by the observations that cells unable to make acidic
phospholipids were sick and that the sickness was suppressed by mutations in DnaA (80,
142). The role of phospholipids remains controversial, as cells unable to make
phospholipids exhibit multiple phenotypes, which appear to be primarily due to the
accumulation of toxic metabolic intermediates, and not due to perturbations in replication
(60).
A simplified model of the E. coli cell cycle suggests that DnaA is synthesized in
the ATP bound form due to the lower Kd for ATP and the high ATP/ADP nucleotide
ratio inside the cell (Figure 6) (30). Once sufficient amounts of DnaA-ATP accumulate,
the expression of dnaA is repressed by DnaA-ATP, and the initiation of DNA replication
proceeds. Once replication starts, the ATPase function of DnaA is activated by n--clamp
and Hda, and DnaA is converted to the inactive DnaA-ADP, which is inert for initiating
replication. Although the conversion of DnaA-ATP to DnaA-ADP relieves the autorepression by DnaA, the expression of dnaA is further limited by the action of SeqA such
that dnaA is expressed only later during the cell cycle. These levels of regulation ensure
that the cells do not initiate replication too soon after the last round of replication
initiation (15). These modes of regulation result in cyclical peaks of DnaA expression
and of DnaA-ATP levels, which is lead to cyclic initiation of replication and expression
of DnaA regulated genes (43, 107, 133, 134, 136).
Regulation of the initiation of DNA replication in bacteria: Regulation of DnaA in
B. subtilis. Like in E. coli, DnaA is a positive regulator of DNA replication in B. subtilis,
I
T•
TD
1I i .
....
.. T
T
Figure 6
Figure 6. DnaA-ATP fluctuation during the cell cycle affects the ability of DnaA to
regulate replication in E.coli. The following example is for slow growth conditions
when replication does not start again until cell division (D) is completed. At the
beginning of the replication cycle (I), DnaA is thought to be primarily in the ATP bound
form, thus stimulating origin melting. Once replication starts, the ATPase function of
DnaA is stimulated by Hda and replisome, converting most of DnaA into the inactive
ADP bound form. Since the expression of dnaA is repressed by SeqA (Seq boxes) and no
new DnaA-ATP is made while the existing DnaA-ATP is converted to DnaA-ADP, the
amount of DnaA-ATP decreases to about 20% of total DnaA. As dnaA begins being
expressed, new DnaA-ATP molecules are being made, most of which are still converted
to DnaA-ADP by replication, but DnaA-ATP levels reach a steady state as they are made
and converted at the same time. Once replication is terminated (T), for the rest of the cell
cycle DnaA-ATP is generated by de novo DnaA synthesis and maybe through
exchanging ADP for ATP in existing DnaA molecules. Once sufficient DnaA-ATP has
been accumulated, a new replication cycle begins. Adapted from Kurokawa, Nishida, et
al. EMBO 1999.
and its expression is auto-regulated (109). However, in B. subtilis the expression of dnaN,
which encodes n-clamp and is in operon with dnaA, is tightly linked to that of dnaA
(109). Overexpression of dnaA from an ectopic locus in B. subtilis causes a decrease in
replication due to the repression of the endogenous dnaANoperon and concomitant
depletion of dnaN, which is required for replication. Overexpression of dnaA in cells
where dnaN is not depleted results in an increased rate of replication, as is observed in E.
coli (109).
B. subtilis DnaA binds ATP and ADP (37), but it is not demonstrated whether
ATP-DnaA is required for origin melting and transcriptional regulation, nor is it known
whether the ratio of ATP/ADP bound DnaA cycles as in E. coli. The adaptor protein that
stimulates the ATPase activity of DnaA in E. coli, Hda, is a member of the AAA+ class
of proteins, but it lacks an obvious homologue in B. subtilis, other than DnaA.
Recently a new negative regulator of the initiation of DNA replication, YabA, was
identified in B. subtilis (102). YabA interacts with DnaA, and a YabA-GFP fusion
localizes with the replication machinery (45, 102, 103). These results led to the
speculation that YabA sequesters DnaA away from the origin of replication, analogously
to the datA locus in E. coli (103). I will discus the function of YabA in more detail in
Chapter 5.
In E. coli peak levels of DnaA are reached at the time of replication initiation (17,
136). The replication initiation models therefore generally propose that the rate-limiting
step in replication initiation is the accumulation of sufficient amounts of DnaA-ATP.
However, in B. subtilis DnaA stops accumulating -15 min prior to the initiation of DNA
replication (99). Based on observations of plasmid replication (65, 112), it is possible that
protein chaperones, ClpX and CIpA, alter the interactions the DnaA molecules at the
origin, such that initiation can proceed without the need for more DnaA molecules.
However, it is also possible that events after DnaA binding to the origin and origin
melting are also regulated, and contribute to the timing of replication initiation.
Unified view of initiation. As discussed above, during the initiation of DNA
replication several events happen: DnaA recognizes and melts the origin of replication,
DNA polymerase holoenzyme, and helicase are loaded at the origin of replication. A
series of in vitro experiments with purified E. coli replication components have shown
that once DnaA melts the origin helicase can be loaded, while DnaA melting of the origin
is insufficient for DNA polymerase to stably associate with the origin (32). Helicase
action to unwind more DNA was required for primase and DNA polymerase to associate
with the origin of replication in vitro (32). The observations lead to a model where DnaA
melts the origin, stimulates helicase loading, and the actions of helicase and primase
recruit the DNA polymerase holoenzyme to the origin (3, 32, 50, 66). In this model, the
only regulated step during initiation is the melting of the origin by DnaA, and once the
origin is melted the following steps happen spontaneously. The order of assembly of the
replication components has not been extensively investigated in vivo, in either B. subtilis
or other bacteria. Understanding the order of events during initiation will likely reveal
additional regulatory mechanisms and offer more clues as to how growth regulates the
initiation of DNA replication. In Chapter 3 I discuss my findings regarding the assembly
of replication components with the origin of replication in vivo in B. subtilis.
Coupling DNA replication and the cell cycle. Cells are exposed to a variety
damaging agents, such as oxygen radicals, ultraviolet and gamma radiation, and
antibiotics, which can damage DNA and perturb replication (36). If the rest of the cell
cycle and cell division proceeded under these conditions, the resulting cells would have
incomplete genomes and likely die (36, 143). To increase their chances of survival, cells
have evolved mechanisms of detecting and repairing DNA damage, as well as delaying
progression through the cell cycle (14, 24, 119, 143). One of the earliest identified such
mechanism in bacteria is the SOS response, which is conserved among many bacteria (2,
7, 22, 36, 114, 115, 141). The SOS response consists of the induction of many genes and
is triggered by perturbations in the elongation step of DNA replication (36). Problems
during replication generate ssDNA, which is not replicated rapidly due to the uncoupling
of helicase and the stalled DNA polymerase, or due to the recession of double stranded
DNA breaks (DSB) by specialized proteins (36). Such exposed ssDNA is bound by the
recombination protein RecA, which in complex with ssDNA inactivates the
transcriptional repressor LexA (81, 88, 96, 125). In B. subtilis, the SOS response is
caused by the induction of LexA-repressed genes, including those with functions in UV
DNA damage repair (uvrAB), recombination (recA), transcriptional regulation (recA,
lexA), DNA lesion bypass (polYl), and cell-division inhibition (yneA) (2, 20, 31, 57, 86).
I discuss the detailed composition of the SOS response in B. subtilis in Chapter 3.
Interestingly, mutants defective in the SOS response can still delay cell division or
sporulation (development) in response to DNA damage (47, 57, 82, 87). These
observations suggest that additional mechanisms operate to detect replication
perturbations and delay cell cycle processes.
The replication initiation protein DnaA is also a transcription factor in all tested
bacteria, including B. subtilis and E. coli (14, 95, 109, 121). In B. subtilis, DnaA has been
linked to coordinating replication with development through delaying sporulation under
conditions where DNA replication is inhibited (14, 119). DnaA directly induces the
expression of a small protein, Sda, which inhibits the early steps of sporulation (14, 118,
119). In B. subtilis, DnaA is a positive regulator of some genes, such as sda, and a
negative regulator of others, such as dnaA-N (14, 109, 118). Studies from E. coli have
indicated that DnaA requires at least one consensus and one more degenerate DnaA
binding site to affect transcription, although the mechanism by which DnaA affects gene
expression is not well understood (42). Studies from E. coli have proposed that DnaA
negatively regulates its own expression by occluding the dnaA promoter away from RNA
polymerase (74). The ability of DnaA to positively regulate transcription is not yet
explained, as there is no known interaction between DnaA and RNA polymerase (128).
One hypothesis is that DnaA alters the topology (supercoiling) of the promoter, which
stimulates RNA polymerase binding to the promoter. Alternatively, DnaA could be
removing or inhibiting the action of negative regulators of transcription, or DnaA could
be interacting with RNA polymerase directly.
The ability of DnaA to function as a transcriptional regulator has been linked to its
nucleotide bound state (43, 130). In E. coli, DnaA-ATP has been shown to inhibit
transcription from the dnaA promoter in vitro (130), and more recent evidence suggests
that other DnaA regulated genes might be affected by the deletion of Hda, which
regulates that ATPase activity of DnaA (43). In B. subtilis, there is little known about the
importance of nucleotide bound form of DnaA with respect to transcriptional regulation.
A better understanding of what genes are regulated by DnaA and their function will
provide more insight into how cells deal with DNA damage. I discuss the effects of
perturbations in DNA replication on gene expression and the involvement of DnaA in
mediation this response in Chapter 2.
Thesis summary. My work has concentrated on understanding how B. subtilis
regulates the initiation of DNA replication, and how replication regulates other cellular
processes. Chapters 2 and 3 focus on the transcriptional responses to perturbations in
DNA replication. Chapter 2 focuses on the recA independent transcriptional responses to
inhibiting initiation or elongation of replication. My data demonstrate that -20 operons
that each contain at least 2 recognizable DnaA binding sites within their promoter
regions, including the two known DnaA regulated operons, are affected under conditions
of perturbed replication. I showed that DnaA binds directly to the promoters of some of
these operons, and that the regulation of one of them, containing essential cell division
genes, is needed for delaying cell division after replication arrest. In collaboration with
Luba Katz and Chris Burge, we showed that many of the gene affected by DnaA in B.
subtilis have homologues in other bacteria, and that the DnaA binding sites are often
conserved as well. These data indicate that DnaA is likely mediating a conserved
response to replication perturbations.
Chapter 3 focuses on comparing the transcriptional effects of different DNA
damaging reagents and replication arrest. The data indicate that RecA is a major but not
the only transcriptional regulator after DNA damage, affecting >450 genes of total of600 affected genes. The effects of RecA are mediated in part by inactivating LexA (- 60
genes) and in part by inducing phage and mobile element genes (-200 genes), and their
indirect effects (-200 genes). Our results also demonstrate that the transcriptional effects
of different DNA damaging reagents are only partially overlapping and that the effects on
replication elongation by these agents are different. Our results suggest that cells respond
differently to different treatments, likely to optimize their survival under the different
conditions.
Chapters 4 and 5 reveal the complexity of the regulation of DNA replication in B.
subtilis. The goal of these experiments was to analyze the order of association of
replication components with the origin of replication, and elucidate mechanisms that
regulate these events. I showed that, contrary to current models, in B. subtilis the DNA
polymerase holoenzyme does not require helicase for association with the origin of
replication. I also show that one of the components that associates before helicase, 3clamp, regulates the initiation of DNA replication by affecting helicase loading.
In Chapter 5 I explored the effects of P-clamp on DNA replication and show that Pclamp and YabA are a part of the same regulatory pathway. The regulated target of this
pathway is likely DnaA. I show that P-clamp and YabA do not affect the expression of
DnaA regulated genes, thus imposing limits on models of how YabA and P-clamp may
affect DnaA.
Acknowledgements
I wish to thank A. Grossman, Catherine Lee, T. Baker, S. Bell, and A. Wright, for helpful
suggestions and comments.
References
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
Ason, B., R. Handayani, C. R. Williams, J. G. Bertram, M. M. Hingorani, M.
O'Donnell, M. F. Goodman, and L. B. Bloom. 2003. Mechanism of loading the
Escherichia coli DNA polymerase III beta sliding clamp on DNA. Bona fide
primer/templates preferentially trigger the gamma complex to hydrolyze ATP and
load the clamp. J Biol Chem 278:10033-40.
Au, N., E. Kuester-Schoeck, V. Mandava, L. E. Bothwell, S. P. Canny, K.
Chachu, S. A. Colavito, S. N. Fuller, E. S. Groban, L. A. Hensley, T. C.
O'Brien, A. Shah, J. T. Tierney, L. L. Tomm, T. M. O'Gara, A. I. Goranov,
A. D. Grossman, and C. M. Lovett. 2005. Genetic composition of the Bacillus
subtilis SOS system. J Bacteriol 187:7655-66.
Baker, T. A., and S. P. Bell. 1998. Polymerases and the replisome: machines
within machines. Cell 92:295-305.
Baker, T. A., K. Sekimizu, B. E. Funnell, and A. Kornberg. 1986. Extensive
unwinding of the plasmid template during staged enzymatic initiation of DNA
replication from the origin of the Escherichia coli chromosome. Cell 45:53-64.
Bates, D., and N. Kleckner. 2005. Chromosome and replisome dynamics in E.
coli: loss of sister cohesion triggers global chromosome movement and mediates
chromosome segregation. Cell 121:899-911.
Bates, D. B., E. Boye, T. Asai, and T. Kogoma. 1997. The absence of effect of
gid or mioC transcription on the initiation of chromosomal replication in
Escherichia coli. Proc Natl Acad Sci U S A 94:12497-502.
Beaber, J. W., B. Hochhut, and M. K. Waldor. 2004. SOS response promotes
horizontal dissemination of antibiotic resistance genes. Nature 427:72-4.
Bell, S. P., and A. Dutta. 2002. DNA replication in eukaryotic cells. Annu Rev
Biochem 71:333-74.
Bowman, G. D., M. O'Donnell, and J. Kuriyan. 2004. Structural analysis of a
eukaryotic sliding DNA clamp-clamp loader complex. Nature 429:724-30.
Bruand, C., M. Farache, S. McGovern, S. D. Ehrlich, and P. Polard. 2001.
DnaB, DnaD and Dnal proteins are components of the Bacillus subtilis replication
restart primosome. Mol Microbiol 42:245-55.
Bruand, C., M. Velten, S. McGovern, S. Marsin, C. Serena, S. D. Ehrlich,
and P. Polard. 2005. Functional interplay between the Bacillus subtilis DnaD and
DnaB proteins essential for initiation and re-initiation of DNA replication. Mol
Microbiol 55:1138-50.
Bruck, I., M. F. Goodman, and M. O'Donnell. 2003. The essential C family
DnaE polymerase is error-prone and efficient at lesion bypass. J Biol Chem
278:44361-8.
Bruck, I., and M. O'Donnell. 2000. The DNA replication machine of a grampositive organism. J Biol Chem 275:28971-83.
Burkholder, W. F., I. Kurtser, and A. D. Grossman. 2001. Replication
initiation proteins regulate a developmental checkpoint in Bacillus subtilis. Cell
104:269-79.
Camara, J. E., A. M. Breier, T. Brendler, S. Austin, N. R. Cozzarelli, and E.
Crooke. 2005. Hda inactivation of DnaA is the predominant mechanism
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
preventing hyperinitiation of Escherichia coli DNA replication. EMBO Rep
6:736-41.
Camara, J. E., K. Skarstad, and E. Crooke. 2003. Controlled initiation of
chromosomal replication in Escherichia coli requires functional Hda protein. J
Bacteriol 185:3244-8.
Campbell, J. L., and N. Kleckner. 1990. E. coli oriC and the dnaA gene
promoter are sequestered from dam methyltransferase following the passage of
the chromosomal replication fork. Cell 62:967-79.
Carr, K. M., and J. M. Kaguni. 2001. Stoichiometry of DnaA and DnaB protein
in initiation at the Escherichia coli chromosomal origin. J Biol Chem 276:4491925.
Castuma, C. E., E. Crooke, and A. Kornberg. 1993. Fluid membranes with
acidic domains activate DnaA, the initiator protein of replication in Escherichia
coli. J Biol Chem 268:24665-8.
Cheo, D. L., K. W. Bayles, and R. E. Yasbin. 1991. Cloning and
characterization of DNA damage-inducible promoter regions from Bacillus
subtilis. J Bacteriol 173:1696-703.
Corn, J. E., P. J. Pease, G. L. Hura, and J. M. Berger. 2005. Crosstalk between
primase subunits can act to regulate primer synthesis in trans. Mol Cell 20:391401.
Courcelle, J., A. Khodursky, B. Peter, P. O. Brown, and P. C. Hanawalt.
2001. Comparative gene expression profiles following UV exposure in wild-type
and SOS-deficient Escherichia coli. Genetics 158:41-64.
Cox, M. M. 2001. Recombinational DNA repair of damaged replication forks in
Escherichia coli: questions. Annu Rev Genet 35:53-82.
Cox, M. M., M. F. Goodman, K. N. Kreuzer, D. J. Sherratt, S. J. Sandler,
and K. J. Marians. 2000. The importance of repairing stalled replication forks.
Nature 404:37-41.
Crooke, E. 2001. Escherichia coli DnaA protein--phospholipid interactions: in
vitro and in vivo. Biochimie 83:19-23.
Crooke, E., C. E. Castuma, and A. Kornberg. 1992. The chromosome origin of
Escherichia coli stabilizes DnaA protein during rejuvenation by phospholipids. J
Biol Chem 267:16779-82.
Dalrymple, B. P., K. Kongsuwan, G. Wijffels, N. E. Dixon, and P. A.
Jennings. 2001. A universal protein-protein interaction motif in the eubacterial
DNA replication and repair systems. Proc Natl Acad Sci U S A 98:11627-32.
Davey, M. J., D. Jeruzalmi, J. Kuriyan, and M. O'Donnell. 2002. Motors and
switches: AAA+ machines within the replisome. Nat Rev Mol Cell Biol 3:826-35.
Dervyn, E., C. Suski, R. Daniel, C. Bruand, J. Chapuis, J. Errington, L.
Janniere, and S. D. Ehrlich. 2001. Two essential DNA polymerases at the
bacterial replication fork. Science 294:1716-9.
Donachie, W. D., and G. W. Blakely. 2003. Coupling the initiation of
chromosome replication to cell size in Escherichia coli. Curr Opin Microbiol
6:146-50.
31.
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
42.
43.
44.
45.
Duigou, S., S. D. Ehrlich, P. Noirot, and M. F. Noirot-Gros. 2004. Distinctive
genetic features exhibited by the Y-family DNA polymerases in Bacillus subtilis.
Mol Microbiol 54:439-51.
Fang, L., M. J. Davey, and M. O'Donnell. 1999. Replisome assembly at oriC,
the replication origin of E. coli, reveals an explanation for initiation sites outside
an origin. Mol Cell 4:541-53.
Felczak, M. M., and J. M. Kaguni. 2004. The box VII motif of Escherichia coli
DnaA protein is required for DnaA oligomerization at the E. coli replication
origin. J Biol Chem 279:51156-62.
Feng, W., L. Rodriguez-Menocal, G. Tolun, and G. D'Urso. 2003.
Schizosacchromyces pombe Dpb2 binds to origin DNA early in S phase and is
required for chromosomal DNA replication. Mol Biol Cell 14:3427-36.
Flores, M. J., S. D. Ehrlich, and B. Michel. 2002. Primosome assembly
requirement for replication restart in the Escherichia coli holDGl 0 replication
mutant. Mol Microbiol 44:783-92.
Friedberg, E. C., Graham C. Walker, and Wolfram Siede. 1995. DNA Repair
and Mutagenesis. ASM Press.
Fukuoka, T., S. Moriya, H. Yoshikawa, and N. Ogasawara. 1990. Purification
and characterization of an initiation protein for chromosomal replication, DnaA,
in Bacillus subtilis. J Biochem (Tokyo) 107:732-9.
Funnell, B. E., T. A. Baker, and A. Kornberg. 1986. Complete enzymatic
replication of plasmids containing the origin of the Escherichia coli chromosome.
J Biol Chem 261:5616-24.
Funnell, B. E., T. A. Baker, and A. Kornberg. 1987. In vitro assembly of a
prepriming complex at the origin of the Escherichia coli chromosome. J Biol
Chem 262:10327-34.
Garner, J., P. Durrer, J. Kitchen, J. Brunner, and E. Crooke. 1998.
Membrane-mediated release of nucleotide from an initiator of chromosomal
replication, Escherichia coli DnaA, occurs with insertion of a distinct region of
the protein into the lipid bilayer. J Biol Chem 273:5167-73.
Gille, H., and W. Messer. 1991. Localized DNA melting and structural
pertubations in the origin of replication, oriC, of Escherichia coli in vitro and in
vivo. Embo J 10:1579-84.
Glinkowska, M., J. Majka, W. Messer, and G. Wegrzyn. 2003. The
mechanism of regulation of bacteriophage lambda pR promoter activity by
Escherichia coli DnaA protein. J Biol Chem 278:22250-6.
Gon, S., J. E. Camara, H. K. Klungsoyr, E. Crooke, K. Skarstad, and J.
Beckwith. 2006. A novel regulatory mechanism couples deoxyribonucleotide
synthesis and DNA replication in Escherichia coli. Embo J 25:1137-47.
Hartwell, L. H., and M. W. Unger. 1977. Unequal division in Saccharomyces
cerevisiae and its implications for the control of cell division. J Cell Biol 75:42235.
Hayashi, M., Y. Ogura, E. J. Harry, N. Ogasawara, and S. Moriya. 2005.
Bacillus subtilis YabA is involved in determining the timing and synchrony of
replication initiation. FEMS Microbiol Lett 247:73-9.
46.
47.
48.
49.
50.
51.
52.
53.
54.
55.
56.
57.
58.
59.
60.
61.
62.
Hingorani, M. M., and M. O'Donnell. 2000. Sliding clamps: a (tail)ored fit.
Curr Biol 10:R25-9.
Ireton, K., and A. D. Grossman. 1992. Coupling between gene expression and
DNA synthesis early during development in Bacillus subtilis. Proc Natl Acad Sci
U S A 89:8808-12.
Ishigo-Oka, D., N. Ogasawara, and S. Moriya. 2001. DnaD protein of Bacillus
subtilis interacts with DnaA, the initiator protein of replication. J Bacteriol
183:2148-50.
Jensen, R. B., S. C. Wang, and L. Shapiro. 2001. A moving DNA replication
factory in Caulobacter crescentus. Embo J 20:4952-63.
Johnson, A., and M. O'Donnell. 2005. Cellular DNA replicases: components
and dynamics at the replication fork. Annu Rev Biochem 74:283-315.
Johnston, G. C., J. R. Pringle, and L. H. Hartwell. 1977. Coordination of
growth with cell division in the yeast Saccharomyces cerevisiae. Exp Cell Res
105:79-98.
Jorgensen, P., and M. Tyers. 2004. How cells coordinate growth and division.
Curr Biol 14:R1014-27.
Karamata, D., and J. D. Gross. 1970. Isolation and genetic analysis of
temperature-sensitive mutants of B. subtilis defective in DNA synthesis. Mol Gen
Genet 108:277-87.
Katayama, T., and A. Kornberg. 1994. Hyperactive initiation of chromosomal
replication in vivo and in vitro by a mutant initiator protein, DnaAcos, of
Escherichia coli. J Biol Chem 269:12698-703.
Katayama, T., T. Kubota, K. Kurokawa, E. Crooke, and K. Sekimizu. 1998.
The initiator function of DnaA protein is negatively regulated by the sliding
clamp of the E. coli chromosomal replicase. Cell 94:61-71.
Kato, J., and T. Katayama. 2001. Hda, a novel DnaA-related protein, regulates
the replication cycle in Escherichia coli. Embo J 20:4253-62.
Kawai, Y., S. Moriya, and N. Ogasawara. 2003. Identification of a protein,
YneA, responsible for cell division suppression during the SOS response in
Bacillus subtilis. Mol Microbiol 47:1113-22.
Kawakami, H., M. Su'etsugu, and T. Katayama. 2006. An isolated Hda-clamp
complex is functional in the regulatory inactivation of DnaA and DNA
replication. J Struct Biol.
Kelman, Z., and M. O'Donnell. 1995. Structural and functional similarities of
prokaryotic and eukaryotic DNA polymerase sliding clamps. Nucleic Acids Res
23:3613-20.
Kikuchi, S., I. Shibuya, and K. Matsumoto. 2000. Viability of an Escherichia
coli pgsA null mutant lacking detectable phosphatidylglycerol and cardiolipin. J
Bacteriol 182:371-6.
Kim, S., H. G. Dallmann, C. S. McHenry, and K. J. Marians. 1996. Coupling
of a replicative polymerase and helicase: a tau-DnaB interaction mediates rapid
replication fork movement. Cell 84:643-50.
Kitchen, J. L., Z. Li, and E. Crooke. 1999. Electrostatic interactions during
acidic phospholipid reactivation of DnaA protein, the Escherichia coli initiator of
chromosomal replication. Biochemistry 38:6213-21.
63.
64.
65.
66.
67.
68.
69.
70.
71.
Kobayashi, K., S. D. Ehrlich, A. Albertini, G. Amati, K. K. Andersen, M.
Arnaud, K. Asai, S. Ashikaga, S. Aymerich, P. Bessieres, F. Boland, S. C.
Brignell, S. Bron, K. Bunai, J. Chapuis, L. C. Christiansen, A. Danchin, M.
Debarbouille, E. Dervyn, E. Deuerling, K. Devine, S. K. Devine, O. Dreesen,
J. Errington, S. Fillinger, S. J. Foster, Y. Fujita, A. Galizzi, R. Gardan, C.
Eschevins, T. Fukushima, K. Haga, C. R. Harwood, M. Hecker, D. Hosoya,
M. F. Hullo, H. Kakeshita, D. Karamata, Y. Kasahara, F. Kawamura, K.
Koga, P. Koski, R. Kuwana, D. Imamura, M. Ishimaru, S. Ishikawa, I. Ishio,
D. Le Coq, A. Masson, C. Mauel, R. Meima, R. P. Mellado, A. Moir, S.
Moriya, E. Nagakawa, H. Nanamiya, S. Nakai, P. Nygaard, M. Ogura, T.
Ohanan, M. O'Reilly, M. O'Rourke, Z. Pragai, H. M. Pooley, G. Rapoport, J.
P. Rawlins, L. A. Rivas, C. Rivolta, A. Sadaie, Y. Sadaie, M. Sarvas, T. Sato,
H. H. Saxild, E. Scanlan, W. Schumann, J. F. Seegers, J. Sekiguchi, A.
Sekowska, S. J. Seror, M. Simon, P. Stragier, R. Studer, H. Takamatsu, T.
Tanaka, M. Takeuchi, H. B. Thomaides, V. Vagner, J. M. van Dijl, K.
Watabe, A. Wipat, H. Yamamoto, M. Yamamoto, Y. Yamamoto, K. Yamane,
K. Yata, K. Yoshida, H. Yoshikawa, U. Zuber, and N. Ogasawara. 2003.
Essential Bacillus subtilis genes. Proc Natl Acad Sci U S A 100:4678-83.
Kong, X. P., R. Onrust, M. O'Donnell, and J. Kuriyan. 1992. Threedimensional structure of the beta subunit of E. coli DNA polymerase III
holoenzyme: a sliding DNA clamp. Cell 69:425-37.
Konieczny, I., and D. R. Helinski. 1997. The replication initiation protein of the
broad-host-range plasmid RK2 is activated by the ClpX chaperone. Proc Natl
Acad Sci U S A 94:14378-82.
Kornberg, A., and T. A. Baker. 1992. DNA Replication, Second ed. W.H.
Freeman and Company, New York.
Krause, M., and W. Messer. 1999. DnaA proteins of Escherichia coli and
Bacillus subtilis: coordinate actions with single-stranded DNA-binding protein
and interspecies inhibition during open complex formation at the replication
origins. Gene 228:123-32.
Krause, M., B. Ruckert, R. Lurz, and W. Messer. 1997. Complexes at the
replication origin of Bacillus subtilis with homologous and heterologous DnaA
protein. J Mol Biol 274:365-80.
Kunst, F., N. Ogasawara, I. Moszer, A. M. Albertini, G. Alloni, V. Azevedo,
M. G. Bertero, P. Bessieres, A. Bolotin, S. Borchert, R. Borriss, L. Boursier,
A. Brans, M. Braun, S. C. Brignell, S. Bron, S. Brouillet, C. V. Bruschi, B.
Caldwell, V. Capuano, N. M. Carter, S. K. Choi, J. J. Codani, I. F.
Connerton, A. Danchin, and et al. 1997. The complete genome sequence of the
gram-positive bacterium Bacillus subtilis. Nature 390:249-56.
Kurokawa, K., S. Nishida, A. Emoto, K. Sekimizu, and T. Katayama. 1999.
Replication cycle-coordinated change of the adenine nucleotide-bound forms of
DnaA protein in Escherichia coli. Embo J 18:6642-52.
Laurent, S. J. 1973. Initiation of deoxyribonucleic acid replication in a
temperature-sensitive mutant of B. subtilis: evidence for a transcriptional step. J
Bacteriol 116:141-5.
72.
73.
74.
75.
76.
77.
78.
79.
80.
81.
82.
83.
84.
85.
86.
87.
Le Chatelier, E., O. J. Becherel, E. d'Alencon, D. Canceill, S. D. Ehrlich, R.
P. Fuchs, and L. Janniere. 2004. Involvement of DnaE, the second replicative
DNA polymerase from Bacillus subtilis, in DNA mutagenesis. J Biol Chem
279:1757-67.
Lee, J. B., R. K. Hite, S. M. Hamdan, X. S. Xie, C. C. Richardson, and A. M.
van Oijen. 2006. DNA primase acts as a molecular brake in DNA replication.
Nature 439:621-4.
Lee, Y. S., and D. S. Hwang. 1997. Occlusion of RNA polymerase by
oligomerization of DnaA protein over the dnaA promoter of Escherichia coli. J
Biol Chem 272:83-8.
Lemon, K. P., and A. D. Grossman. 1998. Localization of bacterial DNA
polymerase: evidence for a factory model of replication. Science 282:1516-9.
Lemon, K. P., and A. D. Grossman. 2000. Movement of replicating DNA
through a stationary replisome. Mol Cell 6:1321-30.
Lemon, K. P., S. Moriya, N. Ogasawara, and A. D. Grossman. 2002.
Chromosome Replication and Segregation, p. 73-86. In A. L. Sonenshein, J. A.
Hoch, and R. Losick (ed.), Bacillus subtilis and its closest relatives. ASM Press,
Washington, DC.
Leonhardt, H., H. P. Rahn, P. Weinzierl, A. Sporbert, T. Cremer, D. Zink,
and M. C. Cardoso. 2000. Dynamics of DNA replication factories in living cells.
J Cell Biol 149:271-80.
Leu, F. P., R. Georgescu, and M. O'Donnell. 2003. Mechanism of the E. coli
tau processivity switch during lagging-strand synthesis. Mol Cell 11:315-27.
Li, Z., J. L. Kitchen, K. Boeneman, P. Anand, and E. Crooke. 2005.
Restoration of growth to acidic phospholipid-deficient cells by DnaA(L366K) is
independent of its capacity for nucleotide binding and exchange and requires
DnaA. J Biol Chem 280:9796-801.
Little, J. W. 1984. Autodigestion of lexA and phage lambda repressors. Proc Natl
Acad Sci U S A 81:1375-9.
Liu, G., K. Begg, A. Geddes, and W. D. Donachie. 2001. Transcription of
essential cell division genes is linked to chromosome replication in Escherichia
coli. Mol Microbiol 40:909-16.
Lobner-Olesen, A., K. Skarstad, F. G. Hansen, K. von Meyenburg, and E.
Boye. 1989. The DnaA protein determines the initiation mass of Escherichia coli
K-12. Cell 57:881-9.
Lopez de Saro, F. J., R. E. Georgescu, M. F. Goodman, and M. O'Donnell.
2003. Competitive processivity-clamp usage by DNA polymerases during DNA
replication and repair. Embo J 22:6408-18.
Lopez de Saro, F. J., R. E. Georgescu, and M. O'Donnell. 2003. A peptide
switch regulates DNA polymerase processivity. Proc Natl Acad Sci U S A
100:14689-94.
Love, P. E., M. J. Lyle, and R. E. Yasbin. 1985. DNA-damage-inducible (din)
loci are transcriptionally activated in competent Bacillus subtilis. Proc Natl Acad
Sci U S A 82:6201-5.
Love, P. E., and R. E. Yasbin. 1984. Genetic characterization of the inducible
SOS-like system of Bacillus subtilis. J Bacteriol 160:910-20.
88.
89.
90.
91.
92.
93.
94.
95.
96.
97.
98.
99.
100.
101.
102.
103.
Lovett, C. M., Jr., T. M. O'Gara, and J. N. Woodruff. 1994. Analysis of the
SOS inducing signal in Bacillus subtilis using Escherichia coli LexA as a probe. J
Bacteriol 176:4914-23.
Marians, K. J. 2000. Replication and recombination intersect. Curr Opin Genet
Dev 10:151-6.
Marsin, S., S. McGovern, S. D. Ehrlich, C. Bruand, and P. Polard. 2001.
Early steps of Bacillus subtilis primosome assembly. J Biol Chem 276:45818-25.
Marszalek, J., and J. M. Kaguni. 1994. DnaA protein directs the binding of
DnaB protein in initiation of DNA replication in Escherichia coli. J Biol Chem
269:4883-90.
McGarry, K. C., V. T. Ryan, J. E. Grimwade, and A. C. Leonard. 2004. Two
discriminatory binding sites in the Escherichia coli replication origin are required
for DNA strand opening by initiator DnaA-ATP. Proc Natl Acad Sci U S A
101:2811-6.
Messer, W. 2002. The bacterial replication initiator DnaA. DnaA and oriC, the
bacterial mode to initiate DNA replication. FEMS Microbiol Rev 26:355-74.
Messer, W., F. Blaesing, D. Jakimowicz, M. Krause, J. Majka, J. Nardmann,
S. Schaper, H. Seitz, C. Speck, C. Weigel, G. Wegrzyn, M. Welzeck, and J.
Zakrzewska-Czerwinska. 2001. Bacterial replication initiator DnaA. Rules for
DnaA binding and roles of DnaA in origin unwinding and helicase loading.
Biochimie 83:5-12.
Messer, W., and C. Weigel. 1997. DnaA initiator--also a transcription factor.
Mol Microbiol 24:1-6.
Miller, M. C., J. B. Resnick, B. T. Smith, and C. M. Lovett, Jr. 1996. The
bacillus subtilis dinR gene codes for the analogue of Escherichia coli LexA.
Purification and characterization of the DinR protein. J Biol Chem 271:33502-8.
Morigen, E. Boye, K. Skarstad, and A. Lobner-Olesen. 2001. Regulation of
chromosomal replication by DnaA protein availability in Escherichia coli: effects
of the datA region. Biochim Biophys Acta 1521:73-80.
Moriya, S., T. Fukuoka, N. Ogasawara, and H. Yoshikawa. 1988. Regulation
of initiation of the chromosomal replication by DnaA-boxes in the origin region
of the Bacillus subtilis chromosome. Embo J 7:2911-7.
Moriya, S., Y. Imai, A. K. Hassan, and N. Ogasawara. 1999. Regulation of
initiation of Bacillus subtilis chromosome replication. Plasmid 41:17-29.
Morrison, A., H. Araki, A. B. Clark, R. K. Hamatake, and A. Sugino. 1990. A
third essential DNA polymerase in S. cerevisiae. Cell 62:1143-51.
Nishida, S., K. Fujimitsu, K. Sekimizu, T. Ohmura, T. Ueda, and T.
Katayama. 2002. A nucleotide switch in the Escherichia coli DnaA protein
initiates chromosomal replication: evidnece from a mutant DnaA protein
defective in regulatory ATP hydrolysis in vitro and in vivo. J Biol Chem
277:14986-95.
Noirot-Gros, M. F., E. Dervyn, L. J. Wu, P. Mervelet, J. Errington, S. D.
Ehrlich, and P. Noirot. 2002. An expanded view of bacterial DNA replication.
Proc Natl Acad Sci U S A 99:8342-7.
Noirot-Gros, M. F., M. Velten, M. Yoshimura, S. McGovern, T. Morimoto, S.
D. Ehrlich, N. Ogasawara, P. Polard, and P. Noirot. 2006. Functional
104.
105.
106.
107.
108.
109.
110.
111.
112.
113.
114.
115.
116.
117.
118.
dissection of YabA, a negative regulator of DNA replication initiation in Bacillus
subtilis. Proc Natl Acad Sci U S A 103:2368-73.
Norris, V., J. Fralick, and A. Danchin. 2000. A SeqA hyperstructure and its
interactions direct the replication and sequestration of DNA. Mol Microbiol
37:696-702.
O'Donnell, M., J. Kuriyan, X. P. Kong, P. T. Stukenberg, and R. Onrust.
1992. The sliding clamp of DNA polymerase III holoenzyme encircles DNA. Mol
Biol Cell 3:953-7.
Ogasawara, N., S. Moriya, K. von Meyenburg, F. G. Hansen, and H.
Yoshikawa. 1985. Conservation of genes and their organization in the
chromosomal replication origin region of Bacillus subtilis and Escherichia coli.
Embo J 4:3345-50.
Ogawa, T., and T. Okazaki. 1994. Cell cycle-dependent transcription from the
gid and mioC promoters of Escherichia coli. J Bacteriol 176:1609-15.
Ogawa, T., Y. Yamada, T. Kuroda, T. Kishi, and S. Moriya. 2002. The datA
locus predominantly contributes to the initiator titration mechanism in the control
of replication initiation in Escherichia coli. Mol Microbiol 44:1367-75.
Ogura, Y., Y. Imai, N. Ogasawara, and S. Moriya. 2001. Autoregulation of the
dnaA-dnaN operon and effects of DnaA protein levels on replication initiation in
Bacillus subtilis. J Bacteriol 183:3833-41.
Onrust, R., and M. O'Donnell. 1993. DNA polymerase III accessory proteins.
II. Characterization of delta and delta'. J Biol Chem 268:11766-72.
Ote, T., M. Hashimoto, Y. Ikeuchi, M. Su'etsugu, T. Suzuki, T. Katayama,
and J. Kato. 2006. Involvement of the Escherichia coli folate-binding protein
YgfZ in RNA modification and regulation of chromosomal replication initiation.
Mol Microbiol 59:265-75.
Pak, M., and S. Wickner. 1997. Mechanism of protein remodeling by ClpA
chaperone. Proc Natl Acad Sci U S A 94:4901-6.
Polard, P., S. Marsin, S. McGovern, M. Velten, D. B. Wigley, S. D. Ehrlich,
and C. Bruand. 2002. Restart of DNA replication in Gram-positive bacteria:
functional characterisation of the Bacillus subtilis PriA initiator. Nucleic Acids
Res 30:1593-605.
Quillardet, P., M. A. Rouffaud, and P. Bouige. 2003. DNA array analysis of
gene expression in response to UV irradiation in Escherichia coli. Res Microbiol
154:559-72.
Rand, L., J. Hinds, B. Springer, P. Sander, R. S. Buxton, and E. O. Davis.
2003. The majority of inducible DNA repair genes in Mycobacterium tuberculosis
are induced independently of RecA. Mol Microbiol 50:1031-42.
Rocha, E. 2002. Is there a role for replication fork asymmetry in the distribution
of genes in bacterial genomes? Trends Microbiol 10:393-5.
Rokop, M. E., J. M. Auchtung, and A. D. Grossman. 2004. Control of DNA
replication initiation by recruitment of an essential initiation protein to the
membrane of Bacillus subtilis. Mol Microbiol 52:1757-67.
Rowland, S. L., W. F. Burkholder, K. A. Cunningham, M. W. Maciejewski,
A. D. Grossman, and G. F. King. 2004. Structure and mechanism of action of
119.
120.
121.
122.
123.
124.
125.
126.
127.
128.
129.
130.
131.
132.
133.
134.
Sda, an inhibitor of the histidine kinases that regulate initiation of sporulation in
Bacillussubtilis. Mol. Cell in press.
Ruvolo, M. V., K. E. Mach, and W. F. Burkholder. 2006. Proteolysis of the
replication checkpoint protein Sda is necessary for the efficient initiation of
sporulation after transient replication stress in Bacillus subtilis. Mol Microbiol
60:1490-508.
Ryan, V. T., J. E. Grimwade, J. E. Camara, E. Crooke, and A. C. Leonard.
2004. Escherichia coli prereplication complex assembly is regulated by dynamic
interplay among Fis, IHF and DnaA. Mol Microbiol 51:1347-59.
Salazar, L., E. Guerrero, Y. Casart, L. Turcios, and F. Bartoli. 2003.
Transcription analysis of the dnaA gene and oriC region of the chromosome of
Mycobacterium smegmatis and Mycobacterium bovis BCG, and its regulation by
the DnaA protein. Microbiology 149:773-84.
Sandler, S. J., and K. J. Marians. 2000. Role of PriA in replication fork
reactivation in Escherichia coli. J Bacteriol 182:9-13.
Sanjanwala, B., and A. T. Ganesan. 1989. DNA polymerase III gene of Bacillus
subtilis. Proc Natl Acad Sci U S A 86:4421-4.
Sanjanwala, B., and A. T. Ganesan. 1991. Genetic structure and domains of
DNA polymerase III of Bacillus subtilis. Mol Gen Genet 226:467-72.
Sassanfar, M., and J. W. Roberts. 1990. Nature of the SOS-inducing signal in
Escherichia coli. The involvement of DNA replication. J Mol Biol 212:79-96.
Sekimizu, K., D. Bramhill, and A. Kornberg. 1987. ATP activates dnaA protein
in initiating replication of plasmids bearing the origin of the E. coli chromosome.
Cell 50:259-65.
Simmons, L. A., A. M. Breier, N. R. Cozzarelli, and J. M. Kaguni. 2004.
Hyperinitiation of DNA replication in Escherichia coli leads to replication fork
collapse and inviability. Mol Microbiol 51:349-58.
Skarstad, K., and E. Boye. 1994. The initiator protein DnaA: evolution,
properties and function. Biochim Biophys Acta 1217:111-30.
Soultanas, P. 2002. A functional interaction between the putative primosomal
protein DnaI and the main replicative DNA helicase DnaB in Bacillus. Nucleic
Acids Res 30:966-74.
Speck, C., C. Weigel, and W. Messer. 1999. ATP- and ADP-dnaA protein, a
molecular switch in gene regulation. Embo J 18:6169-76.
Su'etsugu, M., T. R. Shimuta, T. Ishida, H. Kawakami, and T. Katayama.
2005. Protein associations in DnaA-ATP hydrolysis mediated by the Hdareplicase clamp complex. J Biol Chem 280:6528-36.
Su'etsugu, M., M. Takata, T. Kubota, Y. Matsuda, and T. Katayama. 2004.
Molecular mechanism of DNA replication-coupled inactivation of the initiator
protein in Escherichia coli: interaction of DnaA with the sliding clamp-loaded
DNA and the sliding clamp-Hda complex. Genes Cells 9:509-22.
Sun, L., and J. A. Fuchs. 1992. Escherichia coli ribonucleotide reductase
expression is cell cycle regulated. Mol Biol Cell 3:1095-105.
Sun, L., B. A. Jacobson, B. S. Dien, F. Srienc, and J. A. Fuchs. 1994. Cell
cycle regulation of the Escherichia coli nrd operon: requirement for a cis-acting
upstream AT-rich sequence. J Bacteriol 176:2415-26.
135.
136.
137.
138.
139.
140.
141.
142.
143.
Sutton, M. D., K. M. Carr, M. Vicente, and J. M. Kaguni. 1998. Escherichia
coli DnaA protein. The N-terminal domain and loading of DnaB helicase at the E.
coli chromosomal origin. J Biol Chem 273:34255-62.
Theisen, P. W., J. E. Grimwade, A. C. Leonard, J. A. Bogan, and C. E.
Helmstetter. 1993. Correlation of gene transcription with the time of initiation of
chromosome replication in Escherichia coli. Mol Microbiol 10:575-84.
Velten, M., S. McGovern, S. Marsin, S. D. Ehrlich, P. Noirot, and P. Polard.
2003. A two-protein strategy for the functional loading of a cellular replicative
DNA helicase. Mol Cell 11:1009-20.
Wake, R., and G. King. 1997. A tale of two terminators: crystal structures
sharpen the debate on DNA replication fork arrest mechanisms. Structure 5:1-5.
Wake, R. G. 1997. Replication fork arrest and termination of chromosome
replication in Bacillus subtilis. FEMS Microbiol Lett 153:247-54.
Wake, R. G., and J. Errington. 1995. Chromosome partitioning in bacteria.
Annu Rev Genet 29:41-67.
Witkin, E. M. 1976. Ultraviolet mutagenesis and inducible DNA repair in
Escherichia coli. Bacteriol Rev 40:869-907.
Zheng, W., Z. Li, K. Skarstad, and E. Crooke. 2001. Mutations in DnaA
protein suppress the growth arrest of acidic phospholipid-deficient Escherichia
coli cells. Embo J 20:1164-72.
Zhou, B. B., and S. J. Elledge. 2000. The DNA damage response: putting
checkpoints in perspective. Nature 408:433-9.
Chapter 2
A transcriptional response to replication status mediated by the
conserved bacterial replication protein DnaA
Alexi I. Goranov, Luba Katz, Adam M. Breier, Christopher B. Burge, and Alan D. Grossman
This chapter was previously published in PNAS, 2005, 102 (36): 12932-7.
For this chapter, Adam Breier helped set up the chromatin immunoprecipitation assay to study
DnaA association with promoter regions, and L. Kats and C. Burge helped with comparative
genomic studies investigating conservation of the DnaA binding sites across bacterial genomes.
Abstract
Organisms respond to perturbations in DNA replication. We characterized the global
transcriptional response to inhibition of DNA replication in Bacillus subtilis. We focused on
changes that were independent of the known recA-dependent SOS response. We found that
overlapping sets of genes are affected by perturbations in replication elongation or initiation, and
that this transcriptional response serves to inhibit cell division and maintain cell viability.
Approximately 20 of the operons (>50 genes) affected have potential DnaA binding sites and are
probably regulated directly by DnaA, the highly conserved replication initiation protein and
transcription factor. Many of these genes have homologues and recognizable DnaA binding sites
in other bacteria, indicating that a DnaA-mediated response, elicited by changes in DNA
replication status, may be conserved.
Introduction
Complete duplication and segregation of genomic material is essential for the production of
viable progeny. Prokaryotes and eukaryotes have several mechanisms to ensure that
chromosomes are fully duplicated, including mechanisms to repair and restart damaged or
arrested replication forks (9, 72). Such replication repair and restart mechanisms play a central
role in the bacterial cell-cycle as bacteria frequently experience replication fork arrests, even
under normal growth conditions (9).
Bacteria such as Escherichiacoli and Bacillus subtilis, much like eukaryotic cells, coordinate
cell division with DNA replication (16, 30). The SOS response is one response used to repair
DNA damage and to coordinate cell division with replication (16). During the SOS response, the
highly conserved recombination protein RecA causes the induction of transcription of a set of
genes involved in DNA repair and the inhibition of cell division (16). These genes are normally
repressed by the sequence specific binding of the transcriptional repressor LexA. RecA elicits the
SOS response upon binding single-stranded DNA and stimulating the autocleavage of LexA
(16). RecA-inducible, LexA-repressible genes have been extensively characterized in several
bacterial species including B. subtilis (7, 31, 52, 71). In B. subtilis, cell-division and
development (sporulation) are regulated, in part, by a RecA-dependent mechanism (24, 30).
Many bacteria also have RecA-independent pathways for inhibiting cell division and
development in response to replication fork arrest, but these are not well characterized (24, 30,
39, 40). Inhibiting replication initiation in B. subtilis also inhibits spore formation by a RecAindependent pathway (5, 25, 36). This inhibition of spore formation is mediated by the
transcriptional activation of sda, which indirectly reduces the expression of many genes needed
for sporulation (5, 56).
We set out to identify genes whose transcription is affected independently of recA in
response to perturbations in replication in B. subtilis, and to identify the regulatory factors that
bring about this alteration in gene expression. Using whole genome DNA microarrays, we found
that many genes are affected by inhibition of both replication elongation and initiation,
independently of RecA and Sda. We found that this RecA-independent transcriptional response
helps coordinate DNA replication with cell division by regulating the expression offtsL, an
essential cell division gene.
Our findings demonstrate that DnaA is a critical mediator of the RecA-independent response.
DnaA, the DNA replication initiation protein, is virtually ubiquitous among bacteria and is
structurally and functionally similar to Origin Recognition Complex (ORC) proteins from
eukaryotes and archea (14, 32, 43). DnaA is also a sequence-specific transcription factor (5, 44,
51). Our results indicate that 56 genes in 20 operons may be directly regulated by DnaA, and
bioinformatic analysis indicates that DnaA may regulate many of the homologous genes in
diverse bacterial species. Thus the RecA-independent, DnaA-mediated transcriptional response
appears highly conserved.
Materials and Methods
Strains and alleles. Standard procedures were used for strain construction and growth
conditions (20). B. subtilis strains used, with genotypes, are listed in Table 1.
AyneAB::spc is a deletion-insertion that removes the 3' 151 nucleotides (of 315) of yneA, the
yneA-yneB intergenic region, and the 5' 162 nucleotides (of 651) of yneB and replaces them with
spc (spectinomycin-resistance) cassette.
The Pspac-ftsL fusion was made by amplifying the completeftsL ORF by PCR and inserting
it downstream from the LacI-repressible-IPTG-inducible promoter Pspac in the vector pAG58
(26) generating pAIG6. This plasmid was integrated by single crossover into the chromosome of
strain AIG 105 to generate strain AIG107, containing a Pspac fused toftsL and the downstream
gene pbpB,ftsL+::Pspac-ftsL-pbpB. The resulting strain contains two functional copies offtsL,
one under its native promoter and one under Pspac control, and a single copy of the essential
gene pbpB under Pspac control.
AIG4 (recA::neoAsda) was made by transforming an sda null mutant (5) with genomic DNA
from SL7360 (recA::neo) (59). In the double mutant, the presence of the sda deletion mutation
was verified by PCR, and the recA mutation was verified by testing for sensitivity to ultraviolet
light.
A(dnaA-oriC-dnaN)::spc is a deletion-insertion mutation that replaces these genes with a
spectinomycin-resistance (spc) cassette. This construct removes the 1325 3' nucleotides (of
1338) of dnaA, the intergenic region between dnaA and dnaN, including oriC, and the 1050 5'
nucleotides (of 1134) of dnaN. This construct was generated through PCR (69) and transformed
into AIG36. The resulting strain AIG200 replicates from an ectopic DnaA-independent origin,
oriN and has dnaN expressed from an ectopic locus, amyE. The construct was verified by PCR
and growth was dependent upon expression of the ectopic dnaN. The dnaN expression construct
(pMMB 19) places the entire dnaN ORF under control of the xylose-inducible, glucoserepressible promoter PxylA in the plasmid pDR154 and integrates it in the chromosomal amyE
locus by a double cross-over. The plasmid origin of replication, oriN (originally from pLS32
(21)) was taken from plasmid pDL 110 (38) and cloned into vector pMMB 117 to generate
Table 1. B. subtilis strains used.
Strain
Relevant Genotype; comments; reference
AG174
trpC2,pheAl; lab wild type (a.k.a., JH642)
AIG4
recA::neo, Asda
AIG38
dnaC30ts, metB5, ilvAl (obtained from the Bacillus Genetic Stock Center,
strain 1A20) (28)
AIG105
AyneAB::spc
AIG107
AyneAB::spc, ftsL+::pAIG6 (Pspac-ftsL+-pbpB, cat)
AIG200
AdnaA-oriC-dnaN::spc, amyE::pMMB26(PxyMA-dnaN, cat),
spoIIIJ::pMMB 13 8(oriN, kan), pheA1, trpC+
KPL73
dnaD23ts chr::Tn917QHU151(mls)(28, 37)
KI 1365
recA260, dnaB+-zhb-83::Tn917(mls) (25, 37)
KI 1366
recA260, dnaB19ts-zhb-83::Tn917(mls) (25, 28, 37)
iAll strains are derived from AG174 and contain the trpC and pheA mutations, except, AIG38.
plasmid pMMB 138 which integrates into the chromosome at spolllJat 3590. The replication of
oriN is independent of DnaA (21).
DNA microarrays. DNA microarrays were prepared using PCR products from >99% of the
annotated B. subtilis open reading frames spotted onto Coming GAPS II slides, essentially as
described previously (3). Prior to hybridization with biological samples, arrays were
prehybridized for at least 45 min at 420 C in 1% BSA, 5x SSC, 0.1% SDS, washed in water, and
dried (3).
Culture samples were immediately mixed with an equal volume of methanol (pre-chilled to 200 C). Samples were then spun to pellet the cells, the supernatant was discarded and cell pellets
were frozen at -800 C until further use. RNA was extracted using Qiagen RNeasy kits combined
with on-column DNase treatment according to the manufacturer's recommendations. RNA was
then precipitated with ethanol/LiCl and resuspended in RNase-free water to a desired
concentration, usually >1.0 mg/ml. The quality of RNA was checked on agarose gels by
visualizing the integrity of the 23S and 16S rRNA.
To generate cDNA, RNA from the different experimental conditions was reverse-transcribed
in the presence of amino-allyl-dUTP, followed by coupling to Cy5 for all experimental samples,
or Cy3 for all reference RNA. Reference RNA was made by pooling RNA samples from various
strains grown under condition similar to the experimental conditions.
For reverse transcriptase reactions, 10 jtg RNA template was mixed with 2.5 jtg random
hexamers (in 18 pl) and incubated at 700 C for 10min and on ice for 5 min. Reverse transcription
reactions were then started by the addition of a cocktail resulting in a final mix of RNA template,
random hexamer primers, and 300U Superscript Reverse Transcriptase II (Invitrogen), IX RT
buffer, 10mM DTT, and deoxyribonucleoside triphosphates (0.5 mM each dATP, dCTP, and
dGTP; 0.1 mM dTTP; 0.4 mM aminoallyl-dUTP) in a final volume of 30 jLl. The labeling
reactions were incubated at 250 C for 10 min, at 42 0 C for 70 min, and then shifted to 70 0 C for 15
min to stop the reactions. RNA in the reactions was degraded by adding NaOH (33 mM final
concentration) and incubating at 700 C for 10 min. HCI (33mM) was added to each reaction to
neutralize the pH.
Reactions were purified with Qiagen MinElute kits and eluted in 10 jtl volumes and 0.5 ýll
IM NaHCO 3 (pH 9.0) was added to adjust the pH for the coupling reactions. To couple the
fluorescent dyes to cDNA, I lafreshly dissolved Cy3 or Cy5 dye (Amersham) was added to
cDNA and incubated for 1 h in the dark, mixing every 15 min. Reactions were quenched by
incubation with 1.4M hydroxylamine for 15 min.
Each experimental (Cy5-labeled) sample was mixed with an aliquot of reference RNA (Cy3labeled) and mixed samples were purified with Qiagen MinElute kits. The labeled samples were
mixed with 10 ýtg salmon sperm DNA and 0.8 [Lg yeast tRNA and the volume was adjusted to 14
jll. The samples were heated to 100 0 C for 5 min, spun down, mixed with 2X hybridization buffer
(0.05% SDS, 5X SSC, 25% formamide final concentration) and hybridized to DNA on a
microarray for at least 16 h at 42 0 C . Following hybridization, arrays were washed with lx SSC,
0.2% SDS for 5 min at 42 0 C, followed by a 5 min wash with 0.1x SSC, 0.2% SDS at room
temperature, and a final 5 min wash in 0.Ix SSC at room temperature. Arrays were spun to
remove extra liquid and dried with nitrogen gas.
Arrays were scanned and analyzed with GenePix 3.0 software (Axon Instruments, Inc.). The
signal intensity for each spot (gene) {(Cy5/Cy3)g} was normalized to the total signal intensity
{(Cy5/Cy3)t} on the array (essentially the sum of all the spots). Normalized ratios of
experimental RNA abundance over reference RNA was obtained for each spot for which 80% of
the pixels had intensities at least one standard deviation above background. In an average
experiment 96% of all genes gave such a signal. For genes of interest, we verified that the
intensities of 80% of the pixels in the spot were at least two standard deviations above
background.
To compare two conditions, the normalized signals from condition A (e.g., cells treated with
HPUra) were divided by the corresponding signals from condition B (cells not treated with
HPUra); {(Cy5/Cy3)g/(Cy5/Cy3)t}A
+
{(Cy5/Cy3)g/(Cy5/Cy3)t}B.
Since all experiments were done at least in triplicate, we report the average ratio of ratios
from all repetitions. Statistical Analysis of Microarrays (SAM) (68) was used for all
experiments. The input data contained the replicate ratios of experimental over reference in log2
format. Thus, for each gene, at least 6 values were input for calculation of significance: three
from one condition and three from the other. For all experiments, an effect on a gene is
considered statistically significant if there is less than 1%probability that this change occurred
by chance (false positive discovery rate less than 1.0%).
Identification of homologous transcription units. Transcription units of homologous
genes are relatively easy to recognize in different organisms if the operon structure is conserved.
However, transcription units of homologous genes sometimes become rearranged during
evolution: even in closely related species the relative order of genes within operons can be
different, genes become lost or dispersed throughout a genome, and monocistronic genes can
become incorporated into multicistronic operons. This evolution of operons presents a challenge
in finding a homologous upstream regulatory region for a given transcriptional entity. We
developed a program called FindHomolFirstGene (written in the Perl programming language)
which can identify most cases of operon rearrangements among genomes and determine which
gene is the first gene in an orthologous operon.
As an example, consider an operon in B. subtilis that is composed of three genes A-B-C. A
step-by-step decision tree to identify the first gene in the orthologous operon A'-B'-C'-of B.
halodurans is described below:
1). Best matches to the B. subtilis proteins A, B, and C are found in B. haloduranswith
BLASTP (identity cut-off of 30%).
2). Using GenBank annotation, relative genomic position of B. haloduransorthologs is
examined.
a). In the simplest case, A, B, and C all have single well-conserved homologs in B.
halodurans--A', B' and C'--that are clustered together in the same order and orientation, and the
distance between A' and gene X upstream of it on the chromosome is greater than 50 bases (or X
is less than 50 bases away but is expressed from the opposite strand). A' then is the first gene in
the B. haloduransA'-B'-C' operon.
b). A produces no BLASTP hit in B. halodurans,but B' and C' are clustered orthologs of B
and C, and B' is more than 50 bases away from X (or X is less than 50 bases away but is
expressed from the opposite strand). B' then would be called the first gene in the B. halodurans
B'-C' operon.
c). A', B', and C' are orthologs of A, B, C, but C' is the 5'-most gene in B. halodurans. C'
then is the first gene in the C'-A'-B' operon.
d). A', B', and C' are all homologs of A, B, and C, but the distance between A' and X
(where X is in the same orientation and A') is less than 50 bases. Such cases are examined
manually, and if X appeared to have a biological function similar to A', B', and C', then X is
called the first gene in the operon. If the function of X is unknown than the program
FindHomolFirstGene cannot determine which is the first gene in the B. haloduransoperon. If
the function of X appears dissimilar, than A' would be considered the first gene of the operon.
e). A', B', and C' were dispersed throughout the genome of B. halodurans,etc. The program
will not be able to determine the order of genes in B. halodurans.
f). None of the genes A, B, and C produced any hits in B. halodurans. Again, the program
will not be able to determine the order of genes in this organism.
3). Finally, 500 bases of sequence upstream of the initiator codon for each of the 5'-most genes
are obtained and searched for consensus binding sites for DnaA. The probability of finding a site
matching the consensus in any 500 bp region follows a Poisson distribution and is 2.33% (for
44% GC content). The probability of finding 2 sites with a single mismatch in any 500 bp region
(Poisson distribution with a mean of 0.483) is 8.5%.
Media and growth conditions. Cells were grown in defined minimal medium supplemented
with glucose (1%), glutamate (0.1%) and required amino acids.
For experiments in which replication elongation was arrested with 6-hydroxy-phenylazouracil (HPUra), cells were grown at 370 C and HPUra (38 jig/ml final concentration) was added
during mid-exponential growth. For experiments using temperature-sensitive mutants, cells were
grown at 320 C and shifted to 46 0 C during mid-exponential growth. For microarray analysis 7.5
ml samples were taken from treated and untreated cultures grown in parallel before and 15 min,
30 min, and 60 min after addition of HPUra or 30 min, 60 min, and 90 min after shift to nonpermissive temperature.
Chromatin Immuno-Precipitations. DNA fragments bound to DnaA were immunoprecipitated as described (38) except that extracts were successively incubated with 1:10,000
diluted chicken anti-DnaA, 1:500 diluted donkey anti-chicken, and 3%protein A-sepharose
beads, all for one hour at room temperature. Serial dilutions of total DNA and immunoprecipitated DNA were analyzed by PCR. Equal volumes (10 l) were loaded onto a 2% agarose
gel stained with ethidium bromide. Similar results were obtained with DNA from at least two
independent immunopreciptiations. Primer sequences are available upon request.
Microscopy and cell length determinations. Cells were stained with the vital dye FM4-64
(200 ng/ml; Molecular Probes) to visualize membranes and 4',6'-diamidino-2-phenylindole
(DAPI) (40 to 80 ng/ml) to visualize DNA. Microscopy was performed essentially as described
(35). Briefly, stained cells were placed on 1%agarose pads, and images were captured with a
Nikon E800 microscope equipped with a Hamamatsu digital camera. Improvision OpenLabs 2.0
software was used to process images.
Results
Inhibiting elongation of DNA replication affects multiple genes independently of RecA
and Sda
Production of stalled replication forks by arresting replication elongation is known to induce
the recA-dependent SOS response (16). In B. subtilis, RecA affects expression of approximately
300 genes (Goranov et. al. manuscript in preparation). In addition, Sda, which is induced by
replication arrest (5), indirectly affects the expression of over 100 genes involved in sporulation
and other stationary phase phenomena, all controlled by SpoOA (5, 45). To identify regulatory
responses that are independent of recA and sda, some of our analyses were done in recA sda
double mutants.
To inhibit replication elongation, we treated cell cultures with HPUra, which binds to the
catalytic (alpha) subunit of DNA polymerase, encoded by polC (46). In the recA sda double
mutant, treatment with HPUra affected (99% confidence) the mRNA levels of 108 genes in 55
operons that were also affected in wild type cells (Fig. 1; Table 2). The 55 operons contain a
total of 131 genes but the changes in 23 of the genes did not pass our statistical criteria (see
Materials and Methods). Fifteen of the operons had increased and 40 had decreased mRNA
levels after inhibition of replication elongation. In 43 of the 55 operons, the magnitude of these
effects was two to seven fold less in recA mutant cells than in recA+ cells (see Discussion).
We also arrested replication elongation using a temperature sensitive mutation (dnaC30ts) in
the gene for the replicative DNA helicase. Upon shift of the helicase mutant (dnaC30ts,recA+,
sda+ ) to nonpermissive temperature, replication elongation rapidly stops (57). We measured
changes in mRNA levels in the helicase mutant at various times after shift to non-permissive
temperature and found that the response was remarkably similar to that caused by replication
arrest due to treatment with HPUra. Of the 131 genes in the 55 operons that were affected by
HPUra in the recA sda double mutant, 107 were also affected by temperature shift in the helicase
mutant (Fig. 1; Table 2), including peroxide-inducible genes, iron-regulated genes, and genes
involved in nucleotide biosynthesis, DNA replication, and cell-division (see below). The extent
of the effects was similar to that observed in wild type cells treated with HPUra (Fig. 1, Table 2).
Thus, the vast majority of the effects of HPUra on mRNA levels are likely caused by the effects
of HPUra on replication elongation and not by some other effect on cell physiology. These
results indicate that two different mechanisms of inhibiting replication elongation induce a
common transcriptional program that is independent of the well characterized recA-dependent
SOS response.
~.A-""''''''
~""""'''.4b,\..'''''''
p:t-.- ....
- ~-::,:;.~.c+
~cb+
..
D
Figure 1
62
Figure 1. Changes in mRNA levels in response to inhibition of replication. The relative
mRNA levels in cells blocked for replication was determined using cDNA microarrays. Results
are shown for genes in the 55 operons that were significantly affected by blocking replication
elongation with HPUra in both wild type and the recA sda double mutant.
(A-E) Brightest green represents greater than or equal to 3.5-fold decrease and brightest red
represents greater than or equal to 3.5-fold increase in relative mRNA levels in response to a
block in replication. Gray cells indicate no data for the particular gene in the particular
experiment. Genes are organized by operon and functional categories and are presented in the
same order as in Table 1. Arrows next to operon names or descriptions indicate the direction of
the effect.
(A-B) Inhibition of replication elongation with HPUra. A. Wild type (strain AG174); B.
recA sda double mutant (strain AIG4). Samples from parallel cell cultures, untreated or treated
with HPUra, were taken at various times for a direct comparison. Data presented are from 60
min after treatment with HPUra, except for results for the Fur and PerR regulons which were
maximally affected 15 min after replication fork arrest with HPUra.
(C-E) For experiments with temperature sensitive mutants, cells were grown at permissive
temperature (320 C) to mid-exponential phase, then were shifted to non-permissive temperature
(46 0 C) and samples taken at various times after shift. Data from 90 minutes after temperature
shift are presented. C. Temperature sensitive helicase mutant dnaCts (strain AIG38) compared
to wild type (strain AG 174). D. Temperature sensitive initiation mutant dnaBts recA (strain
KI 1366) compared to dnaB+recA (strain KI1365). E. Temperature sensitive initiation mutant
dnaDts (strain KPL73) compared to wild type (AG174). F. Presence of potential DnaA binding
sites. A black cell indicates the presence of at least 2 potential DnaA binding sites (allowing one
mismatch from consensus) within the 500 bp upstream of the first gene of the operon.
Many genes respond to inhibition of either elongation or initiation of DNA replication
The changes in gene expression caused by arresting replication elongation might be due to
the presence of a stalled replication fork, and-or to the absence of active replication. To
distinguish between these possibilities, we arrested replication in the absence of significant fork
stalling. This was done by inhibiting replication initiation by shifting a temperature sensitive
dnaB (dnaBts) mutant to the non-permissive temperature. B. subtilis dnaB encodes a protein
required for replication initiation, and inactivation of this protein prevents initiation of
replication but allows elongation to continue (28, 55). Many genes (approximately 84 in 26
operons) affected by inhibiting replication elongation in cells lacking recA and sda were also
affected by inhibiting replication initiation in dnaBts, recA mutant (Fig. 1, Table 2). Similar
results were observed by inhibiting replication initiation using a temperature sensitive mutation
in dnaD (Fig. 1, Table 2).
Since a set of genes responds to perturbations in both initiation and elongation of DNA
replication, it appears that the mechanism(s) regulating expression of these genes does not
require detection of stalled replication forks.
Affected genes are involved in many essential cellular processes
The genes affected by perturbations in replication initiation and elongation (independently of
recA and sda) are involved in many aspects of bacterial physiology (Fig. 1, Table 2). The genes
are involved in replication (dnaA, dnaN, dnaB), cell-division (yllB-ylxA-ftsL-pbpB), amino acid
and nucleotide biosynthesis (lysC, proBA, pur and pyr operons, nrdEF), and perhaps translation
and ribosome assembly (yqeH,ywlC), as well as other unknown functions.
Other operons are only affected by inhibiting replication elongation. One of the earliest and
strongest effects of blocking replication elongation was the induction of the PerR regulon
Table 2. Effects of inhibiting DNA replication on gene expression
Gene
wt
recA
dnaCts dnaB19 dnaD23
sda
recA
Fold change after replication inhibition
DNA replication and recombination
dnaA
-7.7
-4.4
-5.6
-4.1
-7.4
dnaN
-7.5
-3.8
-4.3
-3.1
-5.5
2
dnaB
-2.4
-2.2
-3.1
ypvA
1.8
1.6
[1.0]
4.0
-2.5
-2.1
-2.7
-2.4
-4.4
-3.2
-2.0
-1.8
[-1.6]
-1.5
.
DnaA
Binding Sites
0 mm I1mm
Gene Product/Function
3
initiation of chromosome replication
11
DNA polymerase III (beta subunit)
3
1
initiation of chromosome replication
1.7
0
3
similar to helicase
-1.9
[-1.5]
-3.3
-2.5
0
2
similar to unknown proteins (mraZ)
S-adenosyl-methyltransferase mraW
-2.2
[1.1]
-1.8
cell-division protein
-2.8
[1.5]
-1.6
penicillin-binding protein 2B
Cell-division
yllB
ylxA
ftsL
b B
Nucleotide metabolism
ymaA
[1.5]
[1.8]
[-1.1]
3.3
2.8
0
1
unknown essential gene
nrdE
nrdF
[1.7]
2.8
[1.7]
2.1
[1.1]
3.1
2.7
1
2
[1.2]
1.9
3.1
ribonucleoside-diphosphate reductase
ribonucleoside-diphosphate reductase
yimaB
uA
[1.6]
[1.8]
[1.3]
5.7
3.5
-8.4
-1.8
-21.1
-2.6
-2.0
0
0
purine biosynthesis
purE
-6.3
-1.9
-9.4
[-1.6]
[-1.2]
0
1
purK
-12.6
-18.2
-11.3
-16.7
-15.2
-19.4
-15.9
-1.9
-2.1
-1.9
-1.8
-1.9
-1.9
-1.9
-9.9
-8.6
-8.9
-6.9
-9.0
-7.5
-6.5
-1.7
-1.9
-1.8
-1.9
-1.7
[-1.4]
[-1.4]
[-1.6]
[-1.3]
[-1.6]
[-1.5]
[-1.5]
purine biosynthesis
purine biosynthesis
purine biosynthesis
purine biosynthesis
purine biosynthesis
purine biosynthesis
purine biosynthesis
purine biosynthesis
-13.1
-4.5
-18.9
-21.7
-1.6
-1.4
-1.6
-1.7
-9.4
-7.7
-12.4
-12.3
[-1.3]
[-1.2]
[-1.4]
[-1.3]
[-1.2]
[-1.4]
[-1.2]
[-1.4]
purine biosynthesis
purine biosynthesis
purine biosynthesis
purine biosynthesis
-3.0
-7.9
-6.2
-1.5
-3.6
-2.9
-2.9
-2.5
-2.7
prE
-3.1
-1.6
-2.5
-2.0
-2.9
-2.7
-2.9
-1.2
-3.2
-3.1
-3.6
-3.6
[-1.5]
[-1.1]
[-1.1]
[-1.2]
[-1.2]
[-1.1]
[1.0]
[1.0]
[1.0]
0
-7.5
-2.4
-7.1
-5.5
-6.9
-4.9
[-1.4]
-1.8
-2.2
-2.1
[-1.4]
-2.1
-1.9
-1.9
-1.9
xpt
-10.4
-1.7
-31.4
[-1.3]
[-1.3]
0
-2.4
[-1.2]
-4.8
[-1.2]
[-1.3]
prB
purC
purS
purQ
purL
purF
purM
purN
purH
purD
pyrP
pyrB
pyrC
pyrAA
pyrAB
pyrK
pyrD
pyrF
buX
-1.9
-1.9
unknown essential gene
2
pyrimidine biosynthesis
pyrimidine biosynthesis
pyrimidine biosynthesis
pyrimidine biosynthesis
pyrimidine biosynthesis
pyrimidine biosynthesis
pyrimidine biosynthesis
pyrimidine biosynthesis
pyrimidine biosynthesis
1
xanthine phosphoribosyltransferase
xantine permease
Antibiotic production
0
3
sublancin 168 lantibiotic biosynthesis
sunA
-3.2
[-1.8]
-2.4
[-1.3]
-2.0
sunT
-3.7
-1.6
-3.9
-2.2
-1.8
ppsA
ppsB
ppsD
-2.6
-1.9
[-1.2]
[1.0]
[-1.1]
-2.6
-1.7
-2.1
[1.0]
[-1.0]
plipastatin synthesis
-2.1
-2.8
-1.7
-1.9
-1.8
-2.5
[1.1]
[1.1]
[1.0]
[1.6]
plipastatin synthesis
plipastatin synthesis
Adaptation to at pical conditions
-8.2
-16.2
-2.6
-2.4
yveK 3
-4.6
ppsE
sublancin 168 lantibiotic transporter
4
0
1
0
plipastatin synthesis
unknown; similar to capsular
polysaccharide biosynthesis proteins
yveL
-1.7
-1.7
-8.7
-6.4
-3.2
unknown
yveM
yveN
yveO
-2.8
-3.1
-2.4
-2.0
-9.6
-14.2
-7.7
-10.7
-4.1
-5.8
unknown
unknown
-1.7
-1.9
[-1.4]
[-1.4]
-1.6
-2.1
-6.5
-22.9
-6.8
-9.8
-3.6
-5.8
unknown; similar to glycosyltransferase
unknown; similar to glycosyltransferase
-2.0
-1.9
-12.0
-11.6
-8.7
-8.9
-3.8
-3.4
similar to capsular polysaccharide biosynt.
unknown; similar to polysaccharide
yveP
yveQ
yveR
biosynt.
-3.5
-1.7
[1.1]
[-1.2]
[1.0]
1
2
similar to delta-endotoxin
yqxM
-2.8
-1.9
-6.8
-10.2
-6.1
0
0
unknown
sipW
-3.2
-1.6
-16.3
-16.0
-6.9
tasA
-4.7
[-1.5]
-59.5
-21.9
-13.2
-1.8
-1.8
0
2
repressor of the pectin utilization operon
-3.2
-1.9
0
1
unknown
Transcriptional regulation
kdgR
-2.3
-1.6
-3.0
signal peptidase I
1
antimicrobial spore component
Metabolism of carbohydrates
yvfB 3
[-1.3]
[-1.2]
-3.6
yvfC
yvfD
-2.1
-2.0
-13.3
-6.1
-4.6
unknown; similar to UDP-gal-P transferase
[-1.2]
[-1.5]
-2.9
-3.2
-2.3
unknown; similar to serine 0-
yvfE
-2.1
-1.9
-14.1
-7.0
-4.3
acetyltransferase
unknown; similar to polysaccharide
yvfF
-1.8
-1.8
-8.1
-3.6
-2.6
biosynt.
unknown
citZ
2.1
[1.4]
[1.2]
[1.3]
1.8
icd
mdh
[1.2]
[1.2]
1.5
[1.5]
[1.1]
[1.1]
[1.1]
[1.4]
[1.4]
[1.5]
citB
2.5
1.8
4.5
[1.3]
[1.4]
M
0
3
isocitrate dehydrogenase
malate dehydrogenase
0
3
aconitate hydratase
xylan degradation
unknowns; similar to NADH-dependent
xynA
-3.0
-1.9
-4.2
[-1.6]
[-1.4]
0
2
yugJ
1.9
1.5
[-2.0]
1.7
1
3.5
[1.2]
[1.6]
0
[-1.7]
[1.3]
0
1.6
2.1
yugK
Iron responsive genes, Fur regulon
dhbA4'5 -11.2 -3.1
-19.0
[1.4]
dhbB
-29.7 -3.0
-106.4
[1.2]
dhbC
-28.7 -3.5
-92.3
[1.3]
dhbE
dhbF
-16.9
-4.5
-3.4
-2.2
-164.8
-22.0
[1.4]
[1.2]
citrate synthase II (major)
butanol dehydrogenase
1
siderophore synthesis
[1.1]
isochorismatase
[1.2]
isochorismate synthase
[1.3]
[1.2]
2,3-dihydroxybenzoate-AMP ligase
2,3-dihydroxybenzoate biosynthesis
3.8
3.1
4.5
4.9
-1.6
-1.4
-1.5
-1.5
-19.4
-17.0
-17.1
-13.6
-1.8
[-1.7]
[-1.5]
-1.8
-1.8
-1.7
-1.7
-2.0
0
4
unknown
unknown
unknown
unknown
22.6
-2.3
-96.9
[1.0]
[-1.5]
0
4
unknown
unknown
unknown
5.2
13.6
-2.6
--2.5
-2.5
-11.9
[-1.2]
[-1.7]
[-1.1]
[-1.3]
0
0
2
1
15.7
-1.6
-64.3
[1.0]
[-1.5]
0
3
ferrichrome ABC transporter
4.0
14.6
-1.6
-1.8
-16.9
-33.3
[-1.4]
[1.4]
[-1.4]
1
unknown
[-1.4]
1
0
2
unknown
14.0
-1.8
-46.1
[1.2]
[-1.3]
0
2
unknown
16.4
--3.9
-22.6
3.0
[1.1]
0
3
unknown
18.5
5.0
-4.0
-3.7
-23.6
-15.2
[-1.2]
[-1.5]
[1.1]
[1.2]
12.2
1.7
2.7
-1.9
11-1.2]
[-1.5]
-41.8
-7.4
-16.3
[-1.7]
[2.0]
[-1.3]
-1.9
[-1.2]
[-1.2]
6.6
[-1.5]
-16.1
[-1.2]
[-1.2]
-1.7
[1.1]
[-1.1]
[-1.5]
0
3
multidrug-efflux transporter
[1.2]
[1.3]
[1.5]
1.6
[1.6]
1.6
[1.7]
[1.7]
1.8
0
3
probable iron-regulated ABC transporter
probable iron-egulated ABC transporter
probable cysteine desulphorase
[1.5]
[1.4]
[1.5]
[1.5]
1.9
2.2
4.6
2.2
2.7
2
8
checkpoint inhibitor of sporulation
0
0
chromosome partitioning; sporulation
Transport
-1.8
mdr4
yur Y4
2.3
yurX
csd
2.3
2.2
1.9
1.9
11.9
_yurV
yurI
2.0
2.1
[1.2]
[1.7]
unknown
unknown
0
0
iron-uptake system
iron-uptake system
iron-uptake system
unknown
similar to NifN homologue
probable iron regulated ABC transporter
Sporulation
sda
2.0
soj
[-1.5]
-1.7
1.5
-2.3
-1.8
spoOJ
[-1.4]
-1.5
1.6
-2.0
-1.9
-2.3
-1.7
[2.0]
1.8
[-1.1]
0
3
flagellar basal-body rod protein
-15.4
-4.2
chromosome positioning; sporulation
Mobility
fliB
Amino acid biosynthesis
sC
-5.6
-2.6
-4.3
1
2
diaminopimelate/lysine biosynthesis
2
5
-3.3
-3.6
Not represented on arrays
-1.9
-2.3
[-1.1]
-1.9
-1.7
-2.4
[-1.1]
-1.8
unknown; similar to unknown proteins
similar to unknown proteins
shikimate 5-dehydrogenase
yqeK
-2.2
-2.7
-2.2
-1.5
[-1.4]
-1.4
-2.7
-3.3
-2.1
[-1.0]
[-1.1]
[1.0]
[-1.4]
[-1.2]
[-1.3]
proB
-1.9
-2.8
-2.3
-2.7
-2.4
-2.3
-2.1
1
[-1.6]
-1.5
[-1.3]
0
proA
proline biosynthesis
proline biosynthesis
23
2.3
[-1.2]
[-1.0]
0
1
alkyl hydroperoxide reductase
yqeG
yqeH
aroD
ygel
-yqe
Detoxification
S
5
hypothetical RNA binding protein
similar to unknown proteins
unknown; similar to unknown proteins
alkyl hydroperoxide reductase
ahpF
8.6
3.7
2.3
[-1.1]
[-1.1]
katA 4,6
81.6
15.1
6.1
[1.1]
[1.1]
0
2
vegetative catalase 1
perR 4,6
3.1
2.0
5.1
[1.0]
[-1.1]
0
2
Repressor of the perooxide inducible
6.5
1.7
5.9
[1.4]
[1.1]
0
1
zosA 4,6
26.8
1.8
175.2
[1.2]
[1.2]
0
2
Zn transporter
nap
-1.7
-1.6
-2.2
-1.9
-2.6
0
2
carboxylesterase NA
1
2
regulon
mrgA
4'
,6
metalloregulation DNA-binding stress
_protein
Unknown
ywzC
ywfO
wgA
[1.1]
[1.1]
[-1.5]
[1.3]
[1.0]
2.8
3.1
2.0
2.1
3.9
5.0
2.7
2.9
2.3
2.6
yydA
yydB
yydC
yydD
[-1.5]
-2.4
-1.7
-2.4
[-1.1]
-1.5
[-1.2]
[-1.5]
-1.7
[-1.2]
[-1.3]
[-1.3]
[-1.4]
-2.0
-1.6
-2.0
-1.5
-1.8
2
7
[1.1]
-1.6
-1.8
[-1.2]
-1.7
[-1.1]
0
0
unknown
unknown
unknown
similar to unknown proteins
unknown
[-1.2]
0
1
unknown
fn E
-2.0
similar to unknown proteins
similar to unknown proteins
fA
-1.8
-1.6
[1.1]
[1.1]
wlC7
-3.2
-2.2
-3.9
-4.5
-3.2
3
6
Predicted translation factor
xkC
ycM
-2.1
-7.9
-2.1
-2.0
[-1.4]
-4.9
[1.1]
[1.0]
0
0
0
0
0
2
yoqM
-1.4
1.6
-2.0
-1.7
[-1.2]
unknown; similar to DNA repair proteins
-2.2
1.9
[1.0]
[1.1]
[1.0]
[1.1]
[1.2]
0
1
unknown
eeA
1.8
1.5
1.7
3.8
3.3
0
1
unknown; similar to unknown proteins
unknown
unknown
'Numbers indicate the average ratio of relative mRNA levels for the indicated gene in cells
blocked for replication compared to that in cells replicating DNA. Results are shown for genes
in the 55 operons (transcription units) that were significantly affected by blocking replication
elongation with HPUra in both wild type and the recA sda double mutant (except for soj and
spoOJ). Data are the same as in Figure 1 and are averages from at least three independent
experiments. Ratios less than 1 are reported as the negative reciprocal (e.g., 0.5 is reported as 2.0). Ratios are within the 99% confidence interval unless bracketed. Genes are grouped by
operons (operons separated by thick lines) within functional categories.
For wild type (wt) and the recA sda mutant, replication was inhibited with HPUra. The
maximal effect was usually observed 60 min after treatment and, unless otherwise noted, data are
reported for relative mRNA levels 60 min after replication arrest (addition of HPUra) compared
to parallel samples with no arrest.
For the temperature sensitive mutants (helicase-ts (dnaC30); dnaB19, and dnaD23), results
are from 90 min after shift to non-permissive temperature compared to wild type cells treated
similarly.
The number of potential DnaA binding sites with perfect match to consensus (0 MM) or one
mismatch (1 MM) from consensus within 500 bp of the start of the indicated gene is indicated.
The sequence 5'-TT(a/t)TnCACA-3' (42), was used as consensus.
2dnaB was
not reproducibly detected on the PCR-based microarrays due to the poor quality
of the PCR-generated spot. Data shown are from experiments using oligonucleotide
microarrays.
3These
operons were previously detected as part of the sigma-H regulon (3).
4The maximal effect for these transcription units was 15 min after replication arrest with
HPUra and these data are presented (for wt and recA sda).
5These
operons were previously identified as part of the Fur regulon (2).
6These
operons were previously identified as part of the PerR regulon (17).
7ywlC
is not significantly expressed during mid-exponential growth, but is expressed during
late exponential growth and perhaps early stationary phase. The strong effects on ywlC
transcription were observed during late exponential phase and the data presented are from
experiments examining expression under these conditions. In the experiments during midexponential growth ywlC mRNA levels only decreased -1.5-fold in both wild type and recA sda
cells 60 min after replication arrest with HPUra.
(peroxide-inducible) and repression of the Fur regulon (iron-responsive). These responses are
known to help cells survive oxidizing conditions (67).
The regulatory response to replication status is mediated in part by DnaA
One of the most interesting effects of replication status is on expression of genes known to be
controlled by DnaA. In B. subtilis, DnaA, the essential and highly conserved replication
initiation protein and transcription factor, is known to repress expression of two genes, dnaA and
dnaN (51), and to activate expression of sda (5). Consistent with previous findings (5, 49), we
observed that the expression of dnaA and dnaN was decreased and that of sda was increased by
inhibition of either replication initiation or elongation (Fig. 1, Table 2).
We searched for potential DnaA binding sites in the regulatory regions upstream (within 500
bp upstream of the start codon) of operons containing genes affected in response to inhibition of
replication elongation in the recA sda mutant. Using a strict consensus sequence for the DnaA
binding site TT(a/t)TNCACA (42) and allowing only one mismatch, we found that of the 55
operons affected by inhibition of replication elongation, 34 (71 genes) contained at least two
potential DnaA binding sites (Fig. 1; 2A; Table 2). Eighteen of these 34 operons were also
significantly affected by inhibition of replication initiation (Fig. 1; Table 2). An additional 15
operons (53 genes) contained one potential DnaA binding site (Fig. 1; Table 2).
Assuming a Poisson distribution of DnaA binding sites across the genome, it is expected that
less than I of every 10 random operons will have two or more DnaA binding sites (8.5% chance
of finding >1 DnaA binding site within a 500bp region based on 33% GC content of B. subtilis).
The total number of operons in the B. subtilis genome with consensus binding sites or >1 site
with one mismatch agrees with this estimate. In contrast, 62% of the 55 identified operons had
more than one potential DnaA binding site. There are operons with >1 potential binding site that
were not affect under the conditions analyzed. These potential sites might not be properly
positioned to affect transcription. There might also be additional regulatory factors affecting
expression of these operons, or we do not fully understand what constitutes a DnaA binding site.
Binding of DnaA to putative regulatory regions in vivo
To test if DnaA is associated with some of the regulatory regions of the affected operons, we
used chromatin immuno-precipitation (ChIP) with anti-DnaA antibodies (Fig. 2B). The 6
promoters with putative DnaA binding sites tested, dnaA, sda, yllB (ftsL), dnaB, ywlC, and yydA,
all were significantly enriched in the immuno-precipitates (Fig. 2B). This enrichment was
dependent on DnaA; there was no significant enrichment in immuno-precipitates from a dnaAnull mutant (Fig. 2B). Two control regions tested, yxbB and pksL, were not enriched in the
immuno-precipitates (Fig. 2B).
Our results indicate that DnaA binds specifically to the promoter regions of dnaA, sda,yllB
(ftsL), dnaB, ywlC, and yydA (Fig. 2B). Based on this, we propose that in B. subtilis DnaA
controls a global regulatory response to perturbations in replication. In addition, we suspect that
DnaA also binds to potential sites upstream of other identified operones. Based on this, we
suggest that at least 19 operons (52 genes) are regulated directly by DnaA.
Repression of an essential cell division gene couples replication status to cell-division in
a recA-independent manner
One of the most important and conserved aspects of the cellular response to DNA damage
and perturbations in replication is the inhibition of cell division. The characterized mechanisms
for inhibiting cell division in E. coli and B. subtilis involve the RecA-dependent induction of
division inhibitors (16, 30). There are also RecA-independent mechanisms for inhibiting cell
division (39, 40), although the genes responsible for this inhibition have not been identified.
A
'f"
"
",.
... II ., •
'/"'
"
.--•••••
t
..
.II I ••
•••
~
~.
,u, ••
dn_e
y_le
.,
I
••
.unA
__
-+
ypvA
...,L.
•
..
,t1.
~
_ t.
,%1••••
__
~
•••••
...........
• _~_ l __nrdE
I •
_
ade
.
yxbB
pksL
~
D
0
~
~
1I1I
YYdA ••
ykuN
kdgR
.
~
__
Y/lB~.
ngB
yeiN
•
••
t
sda
ywlC
ty.c
~
It
IiiIII -
yliS
p-
III. ...
••
.....
,t1- yurV
--_-
IP Total
dnaA
:~~c
-------t-.---.-~--.t----l-
IP Total
.........
dnaB
""-pyr-P
----_.....-----_--_t
_______
B
dnaA
••
•
~iI:i~!I
1£1.1<~15
dnaA
dnaA-
yyetA
Figure 2
73
Figure 2. DnaA appears to directly regulate at least 20 operons. (A) Shown are the
promoter regions of the operons with 2 or more potential DnaA binding sites that are affected by
inhibition both initiation and elongation of DNA replication. The operon containing nrdE is also
included due to the presence of DnaA binding sites in the ymaA ORF, shown as a dashed line.
The relative location and direction of potential DnaA binding sites (arrowheads) in the regions
500 bp upstream of the first gene in the operon are shown. Solid arrowheads represent
consensus DnaA binding sites, and open arrowheads represent single mismatch sites. The bar at
the top indicates the distance in base pairs from the start of the gene indicated at the right.
Published reports summarizing RNA polymerase binding sites were used to label the
transcription start sites, indicated by an arrow and +1 (5, 27).
(B) DnaA binds in vivo to the promoters of genes affected by perturbations in replication.
Cultures of dnaA+ (AG174) or dnaA- (AIG200) strains growing exponentially were treated with
formaldehyde and collected for chromatin immuno-preciptiation analysis. DnaA and cross-linked
DNA were immuno-precipitated with an anti-DnaA antibody. Total DNA was collected from the
same samples before the addition of antibody. The cross-links were reversed and the precipitated
DNA was analyzed by PCR with primers targeting the promoter regions of the genes listed,
except for the dnaA primers which amplify the oriC region between dnaA and dnaN. We
obtained similar results with primers for the DnaA promoter as well (not shown). pksL and yxbB
are genes with no putative DnaA bindings sites (negative controls).
In the dnaA strain, the oriC region is deleted and no PCR product is detected. This strain
initiates replication from an ectopic origin, oriN, which does not require DnaA (21).
Perturbations in replication caused a recA-independent decrease in expression of an operon
that contains genes required for cell division. mRNA levels for yllB, ylxA,ftsL, and pbpB
decreased in the absence of ongoing replication (Fig. 1, Table 2). BothftsL and pbpB are
essential for cell division (12). FtsL is an unstable protein (10, 53); a decrease inftsL mRNA
levels quickly causes a decrease in FtsL protein and inhibits cell division (11). Conversely,
PbpB is likely to be a stable protein. Depleting PbpB alone in B. subtilis results in very slow
inhibition of cell-division (data not shown), and the PbpB homologue in E.coli, FtsI, has been
reported to be a stable protein (15, 19).
If a decrease in the expression of this operon contributes significantly to the inhibition of cell
division in the absence of replication, then continued expression of these genes under conditions
of replication fork arrest should allow cell division to continue. To test this hypothesis, we
constructed a strain that has two copies offtsL, one under its normal regulation and one
controlled by the LacI-repressible-IPTG-inducible promoter Pspac, and a single copy of pbpB
under control of Pspac. This rendered the cells dependent on inducer (IPTG) for growth, since
expression ofpbpB was IPTG dependent. In this strain in the presence of IPTG, the mRNA
levels offtsL after replication arrest are approximately equal to mRNA levels in the untreated
wild type strain as shown by microarrays (data not shown). To focus on the effects offtsL and
pbpB in the recA-independent response, we also deleted yneA, the gene responsible for much of
the recA-dependent inhibition of cell division (30).
We measured cell length (an indicator of cell division frequency) following replication fork
arrest induced by the addition of HPUra. There was a significant difference in the size
distribution betweenftsL+ and Pspac-ftsL-pbpB cells (Fig. 3A). The average length offtsL+cells
150 min after replication fork arrest was 5.0 ± 2.8 [tm (average cell length ± standard deviation)
compared to 3.2 ± 1.6 [tm for the Pspac-ftsL-pbpB cells (P<0.05). In contrast, prior to replication
arrest, the average cell lengths were virtually indistinguishable, 1.8 ± 0.42 tm and 2.0 + 0.53 gtm
forftsL+ and Pspac-ftsL-pbpB cells, respectively (Fig. 3B).
In yneA + cells the effects were similar (data not shown) but the average cell-length was
longer due to the contribution of YneA to inhibiting cell-division. Similar results were obtained
by placing the whole operon (yllB-ylxA-ftsL-pbpB) under the control of an inducible promoter
(data not shown).
In addition to effects on cell division, constitutive expression offtsL-pbpB during replication
fork arrest caused an increase in cell lysis (Fig. 3C) and a decrease in cell viability (Fig. 3D).
Even though there is a general loss of viability after prolonged exposure to HPUra, the PspacftsL-pbpB strain experienced a 4-8-fold greater loss in viability 120 min after replication fork
arrest, as compared to the identically treatedftsL+ strain (Fig. 3D). These results indicate that
regulation of theftsL operon in response to replication status is important for the proper control
of cell division and maintenance of cell viability.
Even under conditions where both the recA-dependent and recA-independent effects on celldivision were bypassed by expression offtsL in ayneA null mutant, there was a partial inhibition
of cell division after replication fork arrest (compare 2.0 Jim before and 3.2 glm after replication
arrest in strain AIG107). We suspect that this inhibition of cell division is due to nucleoid
occlusion (70). In cells arrested for replication elongation, the nucleoid had an aberrant, noncompact morphology that filled most of the cytoplasm (not shown).
FtsL is found at the site of cell division and its presence depends on formation of the FtsZring (Z-ring). Since Z-ring formation is independent of FtsL (11, 61), the regulation of cell
division exerted by FtsL depletion should act after Z-ring formation. Consistent with this
B.
-'' ' Rh'
<:>
<:>
~
o
.~,o
5
;;,
'
o
SO
100
TlrT'le (min) ah1!r HPU,.
no
addition
o
60
Timf! .tt.!or HPUr.
120
addItion
110
(min)
Figure 3
77
Figure 3. Regulation offtsL contributes to the inhibition of cell division. Strains AIG105
(ftsL , AyneAB) and AIG 107 (Pspac-ftsL-pbpB,AyneAB) were grown to mid-exponential phase
with 1 mM IPTG to induce expression from Pspac. Cells were treated with HPUra to arrest
replication, or were left untreated.
(A) Cell-length distribution 150 min after replication fork arrest with HPUra. The length of
712 and 733 cells was measured forftsL÷ (AIG105; gray bars) and Pspac-ftsL-pbpB (AIG107;
black bars) strains, respectively. (B) Cell-length distribution without replication arrest. The
length of 388 and 407 cells was measured forftsL÷ (AIGI05) and Pspac-ftsL-pbpB (AIG107)
strains, respectively. (C) Cell growth measured by optical density at 600nm. Parallel cultures of
ftsL+ (AIG 05; gray symbols) and Pspac-ftsL-pbpB (AIG107; black symbols) strains were either
treated with HPUra at time Omin (round symbols) or left untreated (square symbols). These are
data from a representative experiment. (D) Viability offtsL + (AIG105; gray bars) and Pspac-ftsLpbpB (AIG 107; black bars) strains after treatment with HPUra. Aliquots of cells from the same
cultures as in (C) were withdrawn and plated to assess colony forming units (CFU) at indicated
times after addition of HPUra. CFU are normalized to the CFU present in each culture prior to
the addition of HPUra (time 0 min). Similar results were observed upon multiple repetitions.
hypothesis, HPUra treatment and DNA damage did not abolish Z-ring formation in yneA null
mutants (data not shown, (30)).
Conservation of genes and potential DnaA binding sites among bacterial species
Since DnaA is found in virtually all bacteria, and several of the genes affected in B. subtilis
have homologues in E. coli that are also regulated by DnaA, we thought that DnaA might be
controlling a similar set of genes in other organisms. We first searched several bacterial
genomes for homologues of the genes that are affected by replication status and have putative
DnaA binding sites in B. subtilis. We then searched for potential DnaA binding sites in the
regions upstream of these conserved genes. Strikingly, many of the homologous genes have
putative DnaA binding sites (Fig. 4; Table 3). In all bacterial genomes that we searched, the
number of identified homologous operons with more than one DnaA binding site is higher than
expected by chance (Fig. 4; Table 3). These findings indicate that these homologous operons
with potential DnaA binding sites are likely to be regulated directly by DnaA in several different
bacterial species.
Discussion
Perturbations in DNA replication are fairly common, even during normal growth (9) and
cells respond to these perturbations by altering gene expression and cell physiology to increase
their chances of survival. We analyzed the global transcriptional response elicited by inhibiting
different steps of DNA replication in the bacterium B. subtilis and found that expression of over
100 genes is altered independently of the well characterized recA-dependent SOS response. This
response alters the expression of genes involved in several physiological processes, including
Figure 4
80
Figure 4. Conservation of potential DnaA binding sites in other bacteria. Shown are the first
genes of the 31 operons in B. subtilis that are affected by inhibition of replication elongation,
have more than one potential DnaA binding site, and have homologues in the genomes of the
other bacteria. The yllB operon containsftsL, and the ymaA operon contains nrdE-F.Seven
different genomes were obtained from GenBank and were searched for homologues of the B.
subtilis genes whose transcription was affected by inhibiting DNA replication: B. halodurans
(Bh), B. anthracis(Ba), Listeria innocua (Li), Streptococcuspneumoniae (Sp), Clostridium
acetobutylicum(Ca),E. coli (Ec), Vibrio cholerae (Vc), and Yersiniapestis (Yp). A white space
in a column indicates no homologous gene (i.e., no gene with >30% identity) in the searched
genome, or that the operon structure in B. subtilis was not conserved within the given genome.
For each identified homologue, we searched 500 bp upstream of the start site of the first open
reading frame in the operon for the presence of consensus DnaA binding sites allowing for 0 or 1
mismatch.
Black boxes - more than one putative DnaA binding site with 1-mismatch or at least one
consensus DnaA binding site; gray boxes - single binding site with 1 mismatch; white boxes with
diagonal line- no biding sites matching the consensus or with 1-mismatch.
In all examined genomes, the number of homologous genes that have a consensus site, or two or
more DnaA binding sites with one mismatch was higher than expected by chance. The chance of
finding sites in similar locations in multiple genomes is considerably lower than that for an
individual genome. The numerical data for this figure are presented in Table 3.
DNA replication and cell division. In addition, our results show that there are mechanisms for
detecting the absence of replication which are independent of sensing stalled replication forks.
In recA null cells, the transcriptional effect on most genes was less than that in wild type cells
(Fig. 1, Table 2). The vast majority of these operons have no discernable LexA binding sites and
are not known to be repressed by LexA. Therefore, the differences caused by the presence or
absence of RecA are likely due to the ability of RecA to process stalled replication forks (8, 54,
64). This processing could generate or amplify the signal that elicits the transcriptional response.
The differences could also be an indirect result of induction of one or more LexA-repressed
genes and the activities of their products at the replication forks (64-66).
Conserved regulation by DnaA
A significant portion of the recA-independent transcriptional response appears to be mediated
directly by the conserved replication initiation protein and transcription factor DnaA. Many of
the genes affected have potential DnaA binding sites in their regulatory regions and DnaA is
associated with at least several of these regions in vivo. In addition, some of the genes affected
in B. subtilis have homologues in other species with potential DnaA binding sites upstream of the
open reading frames, indicating that the regulatory response my be highly conserved in bacteria.
Altogether, we propose that at least 20 operons (56 genes) are likely to be regulated directly
by DnaA (Fig. 2). We employed stringent criteria for identifying potential DnaA binding sites
and regulated genes (>1 binding site with 0 or 1 mismatch upstream of the operon affected).
There are examples in E. coli where one site with I mismatch and another more degenerate site
are sufficient for DnaA-dependent regulation. Additionally, some genes controlled by DnaA
have binding sites downstream of the promoter regions, sometimes within an open reading frame
(18, 58). These cases would have been missed by our analysis. Additional DnaA targets might
include operons like soj-spoOJ(that has a consensus site within the soj open reading frame),
yqxM-sipW-tasA (that has >15 sites with 2 mismatches), and yeeA (that has 1 site with 1
mismatch). These three operons responded to both inhibition of initiation and elongation of DNA
replication. Thus, it seems likely that the DnaA regulon includes more genes than those
identified here.
Inhibiting transcription of an essential cell division gene contributes to coupling cell
division to DNA replication
We observed that expression of the essential cell division geneftsL is inhibited by
perturbations in replication and that this inhibition is important for maintaining cell viability in
B. subtilis. Many bacteria have FtsL homologues which are likely unstable and DnaA regulated
(Fig. 4; Supplemental Table 3) (4). Therefore, the inhibition of cell division by DnaA-mediated
inhibition offtsL may be conserved. Decreased synthesis of an unstable protein is an effective
way to regulate gene expression, and in the case offtsL, inhibit cell division.
The transcriptional response to replication perturbations couples replication to
development, metabolism, and perhaps translation
The transcriptional response to perturbations in replication induces expression of sda in a
DnaA-dependent manner (5, 56). Sda couples sporulation to DNA replication status (5). If this
coordination is lost, cells that experience DNA damage continue to develop and lose viability
(5). Thus, at least two of the operons regulated by this global response (sda,ftsL) affect cell
viability.
Increased transcription of nrdEF,which encodes nucleotide reductase needed for
deoxyribonucleotide biosynthesis, is a common response to DNA damage and replication fork
arrest in several organisms (1, 23). The increase in ribonucleotide reductase in Saccharomyces
Table 3. Conservation of DnaA binding sites in other bacteria.
Bs
0, 3
0,3
3,11
3, 1
0,3
0, 3
0,2
0,:2
1,2
0,3
1, 2
0,2
0,2
2,8
0,2
1, 1
0,4
0,2
0,2
0,2
0,3
0,2
1, 2
0,3
2, 5
0,4
0,3
3, 6
1, 2
2, 7'
Bh
0, 2
0,4
6,8
0,3
0,0
0, 1
0,4
zosA
#operons
#operons
with DnaA
ites§ (%)
Gene
citB
citZ
dnaA
dnaB
jhuD
flgB
katA
kdgR
lysC
mdr
ymaA*
perR
pyrP
sda
xynA
ybbB
yclN
ydbN
yfiY
yhfQ
kuN
yllBt
yokG
ypvA
yqeG
yuil
urY
)yIC
ywzC
yyd4
0, 2
0, 1
0,2
0,2
1,5
0,6
0, 1
0,1
0,2
0, 1
Ba
0,2
0,3
8,9
0,2
0,2
Li
0, 1
0,1
9,11
2, 3
0,1
Sp
0, 1
0,2
0,3
0, 0
0,1
0, 5
0,0
0,3
3,4
0,
0,0
0,4
0,0
0, 1
0,1
0,I1
0,1
0,2
3,6
0,2
0,0
0,2
1, 5
0,0
3, 4
0,2
2, 3
, 1
5, 4
1,3
4, 6
0, 3
1,4
0, 2
551:
0,2
25
35
(64)
17
(68)
Ec
0, 1
0,0
1,2
Vc
0, 1
0,2
1,4
Yp
0, 0
1,1
1,2
0, 0
0, 0
0, 1
0, 1
0,1
0,0
0,2T
1, 3
0,2
0, 3
0, 3
0,2
0,1
0, 4
0, 2
0,1
0, 2
0,2
0,1
0,3
0,1
0,0
0,1
0,0
0,4
0,4
0,1
1,4
0,2
0,3
0,2
0,2
0, 1
0, 1
Ca
0,1
0,3
6,6
0,7
0,0
0,0
0,3
0, 1
0, 1
0,4
0,1
1,3
0,0
0, 0
0,0
1,0
0,0
0,1
0, 3
0,3
9, 6
0,0
0,3
0,1
0,0
0,2
0,2
1,0
4, 5
0, 2
4, 3
1, 1
1,3
26
0, 3
21
0, 1
14
0,0
16
0, 2
11
0, 3
16
0, 2
11
20
(77)
10
(48)
8
(64)
10
(62)
5
(45)
5
(31)
8
(62)
0, 1
This table lists the first genes of the 3 1 operons in B. subtilis that are affected by inhibition of
replication elongation, have more than one potential DnaA binding site, and have homologues in
the genomes of the other bacteria we searched. Seven different genomes were obtained from
GenBank and were searched for homologues of the B. subtilis genes whose transcription was
affected by' inhibiting DNA replication: B. halodurans(Bh), B. anthracis(Ba), Listeria innocua
(Li), Streptococcus pneumoniae (Sp), Clostridium acetobutylicum(Ca), E. coli (Ec), Vibrio
cholerae (Vc), and Yersiniapestis (Yp). A blank space in a column indicates that we did not
detect a gene with >30% identity in the searched genome, or that the operon structure in B.
subtilis was not conserved within the given genome. For each identified homologue, we searched
500 bp upstream of the start site of the first open reading frame in the operon for the presence of
consensus DnaA binding sites, allowing for 0 or 1 mismatch. The number of sites with no
mismatches is indicated first followed by the number with one mismatch. In all examined
genomes, the number of homologous genes that have a consensus site, or two or more DnaA
binding sites with one mismatch was higher than expected by chance. The chance of finding sites
in similar locations in multiple genomes is considerably lower than that for an individual
genome.
*The ymaA gene is the first one in the operon that contains the ribonucleotide
reductase genes
nrdE and nrdF.The operon has only one potential DnaA binding site within the promoter region,
but has 3 more potential DnaA binding sites within ymaA.
tftsL is the third gene of the yllB operon, and it is transcribed from the same promoter(s) as
yllB. The structure of this operon is highly conserved.
IFor the B. subtilis calculation, we used the number of total B. subtilis operons that we
initially identified by microarray analysis (55 operons) of which 35 have potential DnaA binding
sites. This includes the 34 operons with more than one potential DnaA binding site and the ymaA
operon.
4Refers only to the operons identified in our study by microarray analysis in B. subtilis or by
homology in other organisms. This category includes operons with one or more consensus DnaA
binding sites, or operons with two or more binding sites with one mismatch, since the probability
of finding a single consensus site is less than the probability of finding 2 sites with one
mismatch.
IThese genes have less than 30% identity with the B. subtilis homologues, but have a similar
proposed enzymatic function and thus are included.
cerevisiae contributes to survival after DNA damage; higher levels of dNTPs are thought to be
necessary for error-prone DNA replication (6). In E. coli the increase in dNTP levels is thought
to contribute to the RecA-dependent regression and repair of stalled replication forks (54).
Expression of the ymaA-nrdE-nrdF-ymaB operon is positively affected by inhibition of either
replication initiation or elongation in B. subtilis. There is one potential DnaA binding site with
one mismatch within the ymaA promoter region and two 1-mismatch and one consensus site
within ymaA (Fig. 2A). Based on these observations we suggest that the ymaA operon in B.
subtilis is regulated by DnaA.
Many of the genes affected by perturbations in replication have no known function, but have
been shown to be essential. Two genes, ywlC and yqeH, may be involved in translation, judging
by their homology to translation factors. Thus, the transcriptional response to perturbations in
replication may adapt cellular physiology by modulating the translational machinery.
Accumulation of the ATP-bound form of DnaA may affect gene expression
We suspect that variations in the amount of DnaA bound to ATP affect transcription in
response to perturbations in replication. The ATP bound form of DnaA is active in replication
initiation and in modulating transcription (33, 41, 60, 62). In E. coli, the amount of ATP-DnaA
is regulated during the cell-cycle by ongoing replication via an intermediary regulatory protein,
Hda (29, 34). B. subtilis lacks an obvious orthologue of Hda but may still regulate ATP
hydrolysis by DnaA (48). Arrest of replication elongation leads to an accumulation of ATPDnaA in E. coli (34). We suggest that arrest of replication elongation or initiation allows high
levels of ATP-DnaA to accumulate and persist which alters the transcription of many genes.
Since the levels of ATP-DnaA fluctuate from 20-80% of total DnaA during a normal E. coli
cell cycle, the expression of at least some of the DnaA regulated genes fluctuate as well (22, 34,
50, 63). The fluctuation of ATP-DnaA levels is one of the mechanisms that regulates the timing
of initiation of DNA replication (13, 34, 47). We suggest that cycling of ATP-DnaA levels
couples replication and other physiological processes including cell division and development.
This DnaA-mediated effect on gene expression has at least two roles: 1) to time and regulate
cell-cycle gene expression and 2) to induce a response to perturbations in replication that help
the cells survive.
Acknowledgments
This work was supported in part by NIH grants GM41934 to ADG, HG002439 to CBB and a
NIH postdoctoral fellowship to LK. We thank E. Kuester-Shoeck and C. Lee for technical help,
M. Barker Berkmen for materials used to construct AIG200, and T. Baker, F. Solomon, S. P.
Bell, and C. Lee for suggestions and comments on the manuscript.
References
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
Augustin, L. B., B. A. Jacobson, and J. A. Fuchs. 1994. Escherichia coli Fis and DnaA
proteins bind specifically to the nrd promoter region and affect expression of an nrd-lac
fusion. J Bacteriol 176:378-87.
Baichoo, N., T. Wang, R. Ye, and J. D. Helmann. 2002. Global analysis of the Bacillus
subtilis Fur regulon and the iron starvation stimulon. Mol Microbiol 45:1613-29.
Britton, R. A., P. Eichenberger, J. E. Gonzalez-Pastor, P. Fawcett, R. Monson, R.
Losick, and A. D. Grossman. 2002. Genome-wide analysis of the stationary-phase
sigma factor (sigma-H) regulon of Bacillus subtilis. J Bacteriol 184:4881-90.
Buddelmeijer, N., N. Judson, D. Boyd, J. J. Mekalanos, and J. Beckwith. 2002.
YgbQ, a cell division protein in Escherichia coli and Vibrio cholerae, localizes in
codependent fashion with FtsL to the division site. Proc Natl Acad Sci U S A 99:631621.
Burkholder, W. F., I. Kurtser, and A. D. Grossman. 2001. Replication initiation
proteins regulate a developmental checkpoint in Bacillus subtilis. Cell 104:269-79.
Chabes, A., B. Georgieva, V. Domkin, X. Zhao, R. Rothstein, and L. Thelander.
2003. Survival of DNA damage in yeast directly depends on increased dNTP levels
allowed by relaxed feedback inhibition of ribonucleotide reductase. Cell 112:391-401.
Courcelle, J., A. Khodursky, B. Peter, P. O. Brown, and P. C. Hanawalt. 2001.
Comparative gene expression profiles following UV exposure in wild-type and SOSdeficient Escherichia coli. Genetics 158:41-64.
Cox, M. M. 2001. Recombinational DNA repair of damaged replication forks in
Escherichia coli: questions. Annu Rev Genet 35:53-82.
Cox, M. M., M. F. Goodman, K. N. Kreuzer, D. J. Sherratt, S. J. Sandier, and K. J.
Marians. 2000. The importance of repairing stalled replication forks. Nature 404:37-41.
Daniel, R. A., and J. Errington. 2000. Intrinsic instability of the essential cell division
protein FtsL of Bacillus subtilis and a role for DivIB protein in FtsL turnover. Mol
Microbiol 36:278-89.
Daniel, R. A., E. J. Harry, V. L. Katis, R. G. Wake, and J. Errington. 1998.
Characterization of the essential cell division gene ftsL(yIID) of Bacillus subtilis and its
role in the assembly of the division apparatus. Mol Microbiol 29:593-604.
Daniel, R. A., A. M. Williams, and J. Errington. 1996. A complex four-gene operon
containing essential cell division gene pbpB in Bacillus subtilis. J Bacteriol 178:2343-50.
Donachie, W. D., and G. W. Blakely. 2003. Coupling the initiation of chromosome
replication to cell size in Escherichia coli. Curr Opin Microbiol 6:146-50.
Erzberger, J. P., M. M. Pirruccello, and J. M. Berger. 2002. The structure of bacterial
DnaA: implications for general mechanisms underlying DNA replication initiation. Embo
J 21:4763-73.
Fraipont, C., M. Adam, M. Nguyen-Disteche, W. Keck, J. Van Beeumen, J. A.
Ayala, B. Granier, H. Hara, and J. M. Ghuysen. 1994. Engineering and
overexpression of periplasmic forms of the penicillin-binding protein 3 of Escherichia
coli. Biochem J 298 ( Pt 1):189-95.
Friedberg, E. C., Graham C. Walker, and Wolfram Siede. 1995. DNA Repair and
Mutagenesis. ASM Press.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
Fuangthong, M., A. F. Herbig, N. Bsat, and J. D. Helmann. 2002. Regulation of the
Bacillus subtilis fur and perR genes by PerR: not all members of the PerR regulon are
peroxide inducible. J Bacteriol 184:3276-86.
Glinkowska, M., J. Majka, W. Messer, and G. Wegrzyn. 2003. The mechanism of
regulation of bacteriophage lambda pR promoter activity by Escherichia coli DnaA
protein. J Biol Chem 278:22250-6.
Goffin, C., C. Fraipont, J. Ayala, M. Terrak, M. Nguyen-Disteche, and J. M.
Ghuysen. 1996. The non-penicillin-binding module of the tripartite penicillin-binding
protein 3 of Escherichia coli is required for folding and/or stability of the penicillinbinding module and the membrane-anchoring module confers cell septation activity on
the folded structure. J Bacteriol 178:5402-9.
Harwood, C. R., and S. M. Cutting. 1990. Molecular Biological Methods for Bacillus.
John Wiley & Sons, Chichester, England.
Hassan, A. K., S. Moriya, M. Ogura, T. Tanaka, F. Kawamura, and N. Ogasawara.
1997. Suppression of initiation defects of chromosome replication in Bacillus subtilis
dnaA and oriC-deleted mutants by integration of a plasmid replicon into the
chromosomes. J Bacteriol 179:2494-502.
Herrick, J., M. Kohiyama, T. Atlung, and F. G. Hansen. 1996. The initiation mess?
Mol Microbiol 19:659-66.
Huang, M., Z. Zhou, and S. J. Elledge. 1998. The DNA replication and damage
checkpoint pathways induce transcription by inhibition of the Crtl repressor. Cell
94:595-605.
Ireton, K., and A. D. Grossman. 1992. Coupling between gene expression and DNA
synthesis early during development in Bacillus subtilis. Proc Natl Acad Sci U S A
89:8808-12.
Ireton, K., and A. D. Grossman. 1994. A developmental checkpoint couples the
initiation of sporulation to DNA replication in Bacillus subtilis. Embo J 13:1566-73.
Jaacks, K. J., J. Healy, R. Losick, and A. D. Grossman. 1989. Identification and
characterization of genes controlled by the sporulation-regulatory gene spo0H in Bacillus
subtilis. J Bacteriol 171:4121-9.
Jarmer, H., T. S. Larsen, A. Krogh, H. H. Saxild, S. Brunak, and S. Knudsen. 2001.
Sigma A recognition sites in the Bacillus subtilis genome. Microbiology 147:2417-24.
Karamata, D., and J. D. Gross. 1970. Isolation and genetic analysis of temperaturesensitive mutants of B. subtilis defective in DNA synthesis. Mol Gen Genet 108:277-87.
Kato, J., and T. Katayama. 2001. Hda, a novel DnaA-related protein, regulates the
replication cycle in Escherichia coli. Embo J 20:4253-62.
Kawai, Y., S. Moriya, and N. Ogasawara. 2003. Identification of a protein, YneA,
responsible for cell division suppression during the SOS response in Bacillus subtilis.
Mol Microbiol 47:1113-22.
Khil, P. P., and R. D. Camerini-Otero. 2002. Over 1000 genes are involved in the DNA
damage response of Escherichia coli. Mol Microbiol 44:89-105.
Kornberg, A., and T. A. Baker. 1992. DNA Replication, Second ed. W.H. Freeman and
Company, New York.
Krause, M., B. Ruckert, R. Lurz, and W. Messer. 1997. Complexes at the replication
origin of Bacillus subtilis with homologous and heterologous DnaA protein. J Mol Biol
274:365-80.
34.
35.
36.
37.
38.
39.
40.
41.
42.
43.
44.
45.
46.
47.
48.
49.
50.
Kurokawa, K., S. Nishida, A. Emoto, K. Sekimizu, and T. Katayama. 1999.
Replication cycle-coordinated change of the adenine nucleotide-bound forms of DnaA
protein in Escherichia coli. Embo J 18:6642-52.
Lee, P. S., D. C. Lin, S. Moriya, and A. D. Grossman. 2003. Effects of the
chromosome partitioning protein SpoOJ (ParB) on oriC positioning and replication
initiation in Bacillus subtilis. J Bacteriol 185:1326-37.
Lemon, K. P., and A. D. Grossman. 2001. The extrusion-capture model for
chromosome partitioning in bacteria. Genes Dev 15:2031-41.
Lemon, K. P., I. Kurtser, J. Wu, and A. D. Grossman. 2000. Control of initiation of
sporulation by replication initiation genes in Bacillus subtilis. J Bacteriol 182:2989-91.
Lin, D. C., and A. D. Grossman. 1998. Identification and characterization of a bacterial
chromosome partitioning site. Cell 92:675-85.
Liu, G., K. Begg, A. Geddes, and W. D. Donachie. 2001. Transcription of essential cell
division genes is linked to chromosome replication in Escherichia coli. Mol Microbiol
40:909-16.
Love, P. E., and R. E. Yasbin. 1984. Genetic characterization of the inducible SOS-like
system of Bacillus subtilis. J Bacteriol 160:910-20.
McGarry, K. C., V. T. Ryan, J. E. Grimwade, and A. C. Leonard. 2004. Two
discriminatory binding sites in the Escherichia coli replication origin are required for
DNA strand opening by initiator DnaA-ATP. Proc Natl Acad Sci U S A 101:2811-6.
Messer, W. 2002. The bacterial replication initiator DnaA. DnaA and oriC, the bacterial
mode to initiate DNA replication. FEMS Microbiol Rev 26:355-74.
Messer, W., F. Blaesing, D. Jakimowicz, M. Krause, J. Majka, J. Nardmann, S.
Schaper, H. Seitz, C. Speck, C. Weigel, G. Wegrzyn, M. Welzeck, and J.
Zakrzewska-Czerwinska. 2001. Bacterial replication initiator DnaA. Rules for DnaA
binding and roles of DnaA in origin unwinding and helicase loading. Biochimie 83:5-12.
Messer, W., and C. Weigel. 1997. DnaA initiator--also a transcription factor. Mol
Microbiol 24:1-6.
Molle, V., M. Fujita, S. T. Jensen, P. Eichenberger, J. E. Gonzalez-Pastor, J. S. Liu,
and R. Losick. 2003. The SpoOA regulon of Bacillus subtilis. Mol Microbiol 50:1683701.
Neville, M. M., and N. C. Brown. 1972. Inhibition of a discrete bacterial DNA
polymerase by 6-(p-hydroxyphenylazo)-uracil and 6-(p-hydroxyphenylazo-)-isocytosine.
Nat New Biol 240:80-2.
Nishida, S., K. Fujimitsu, K. Sekimizu, T. Ohmura, T. Ueda, and T. Katayama.
2002. A nucleotide switch in the Escherichia coli DnaA protein initiates chromosomal
replication: evidnece from a mutant DnaA protein defective in regulatory ATP hydrolysis
in vitro and in vivo. J Biol Chem 277:14986-95.
Noirot-Gros, M. F., E. Dervyn, L. J. Wu, P. Mervelet, J. Errington, S. D. Ehrlich,
and P. Noirot. 2002. An expanded view of bacterial DNA replication. Proc Natl Acad
Sci U S A 99:8342-7.
Ogasawara, N., S. Moriya, K. von Meyenburg, F. G. Hansen, and H. Yoshikawa.
1985. Conservation of genes and their organization in the chromosomal replication origin
region of Bacillus subtilis and Escherichia coli. Embo J 4:3345-50.
Ogawa, T., and T. Okazaki. 1994. Cell cycle-dependent transcription from the gid and
mioC promoters of Escherichia coli. J Bacteriol 176:1609-15.
51.
52.
53.
54.
55.
56.
57.
58.
59.
60.
61.
62.
63.
64.
65.
66.
67.
Ogura, Y., Y. Imai, N. Ogasawara, and S. Moriya. 2001. Autoregulation of the dnaAdnaN operon and effects of DnaA protein levels on replication initiation in Bacillus
subtilis. J Bacteriol 183:3833-41.
Rand, L., J. Hinds, B. Springer, P. Sander, R. S. Buxton, and E. O. Davis. 2003. The
majority of inducible DNA repair genes in Mycobacterium tuberculosis are induced
independently of RecA. Mol Microbiol 50:1031-42.
Robson, S. A., K. A. Michie, J. P. Mackay, E. Harry, and G. F. King. 2002. The
Bacillus subtilis cell division proteins FtsL and DivIC are intrinsically unstable and do
not interact with one another in the absence of other septasomal components. Mol
Microbiol 44:663-74.
Robu, M. E., R. B. Inman, and M. M. Cox. 2001. RecA protein promotes the
regression of stalled replication forks in vitro. Proc Natl Acad Sci U S A 98:8211-8.
Rokop, M. E., J. M. Auchtung, and A. D. Grossman. 2004. Control of DNA
replication initiation by recruitment of an essential initiation protein to the membrane of
Bacillus subtilis. Mol Microbiol 52:1757-67.
Rowland, S. L., W. F. Burkholder, K. A. Cunningham, M. W. Maciejewski, A. D.
Grossman, and G. F. King. 2004. Structure and mechanism of action of Sda, an
inhibitor of the histidine kinases that regulate initiation of sporulation in Bacillus subtilis.
Mol. Cell in press.
Sakamoto, Y., S. Nakai, S. Moriya, H. Yoshikawa, and N. Ogasawara. 1995. The
Bacillus subtilis dnaC gene encodes a protein homologous to the DnaB helicase of
Escherichia coli. Microbiology 141 ( Pt 3):641-4.
Schaefer, C., and W. Messer. 1991. DnaA protein/DNA interaction. Modulation of the
recognition sequence. Mol Gen Genet 226:34-40.
Sciochetti, S. A., P. J. Piggot, D. J. Sherratt, and G. Blakely. 1999. The ripX locus of
Bacillus subtilis encodes a site-specific recombinase involved in proper chromosome
partitioning. J Bacteriol 181:6053-62.
Sekimizu, K., D. Bramhill, and A. Kornberg. 1987. ATP activates dnaA protein in
initiating replication of plasmids bearing the origin of the E. coli chromosome. Cell
50:259-65.
Sievers, J., and J. Errington. 2000. The Bacillus subtilis cell division protein FtsL
localizes to sites of septation and interacts with DivIC. Mol Microbiol 36:846-55.
Speck, C., C. Weigel, and W. Messer. 1999. ATP- and ADP-dnaA protein, a molecular
switch in gene regulation. Embo J 18:6169-76.
Sun, L., and J. A. Fuchs. 1992. Escherichia coli ribonucleotide reductase expression is
cell cycle regulated. Mol Biol Cell 3:1095-105.
Sutton, M. D., J. M. Duzen, and R. W. Maul. 2005. Mutant forms of the Escherichia
colibeta sliding clamp that distinguish between its roles in replication and DNA
polymerase V-dependent translesion DNA synthesis. Mol Microbiol 55:1751-66.
Sutton, M. D., S. Murli, T. Opperman, C. Klein, and G. C. Walker. 2001. umuDCdnaQ Interaction and its implications for cell cycle regulation and SOS mutagenesis in
Escherichia coli. J Bacteriol 183:1085-9.
Sutton, M. D., and G. C. Walker. 2001. Managing DNA polymerases: coordinating
DNA replication, DNA repair, and DNA recombination. Proc Natl Acad Sci U S A
98:8342-9.
Touati, D. 2000. Iron and oxidative stress in bacteria. Arch Biochem Biophys 373:1-6.
68.
69.
70.
71.
72.
Tusher, V. G., R. Tibshirani, and G. Chu. 2001. Significance analysis of microarrays
applied to the ionizing radiation response. Proc Natl Acad Sci U S A 98:5116-21.
Wach, A. 1996. PCR-synthesis of marker cassettes with long flanking homology regions
for gene disruptions in S. cerevisiae. Yeast 12:259-65.
Wu, L. J., and J. Errington. 2004. Coordination of cell division and chromosome
segregation by a nucleoid occlusion protein in Bacillus subtilis. Cell 117:915-25.
Yasbin, R. E., D. Cheo, and K. W. Bayles. 1991. The SOB system of Bacillus subtilis:
a global regulon involved in DNA repair and differentiation. Res Microbiol 142:885-92.
Zhou, B. B., and S. J. Elledge. 2000. The DNA damage response: putting checkpoints in
perspective. Nature 408:433-9.
Chapter 3
Characterization of the global transcriptional responses to different
types of DNA damage and disruption of replication in Bacillus subtilis
Alexi I. Goranov, Elke Kuester-Schoeck, Jue D. Wang, and Alan D. Grossman
This chapter was previously published in the Journal of Bacteriology, J Bacteriol. 2006
Aug; 188(15):5595-5605.
For this chapter, Elke Kuester-Schoeck performed the microarray experiments with MMC and
UV treatments, and Jade Wang performed the genomic microarrays after MMC treatment.
Abstract
DNA damage and perturbations in DNA replication can induce global transcriptional
responses that can help organisms repair the damage and survive. RecA is known to mediate
transcriptional responses to DNA damage in several bacterial species by inactivating the
repressor LexA and phage repressors. To gain insight into how Bacillussubtilis responds to
various types of DNA damage, we measured the effects of DNA damage and perturbations in
replication on mRNA levels using DNA microarrays. We perturbed replication either directly
with p-hydroxyphenylazo-uracil (HPUra), an inhibitor of DNA polymerase, or indirectly with
the DNA damaging reagents mitomycin C (MMC) and ultraviolet (UV) irradiation. Our results
indicate that the transcriptional responses to HPUra, MMC, and UV are only partially
overlapping. recA is the major transcriptional regulator under all of the tested conditions and
LexA appears to directly repress expression of 63 genes in 26 operons, including the 18 operons
previously identified as LexA targets. MMC and HPUra treatments caused induction of an
integrative and conjugative element (ICEBsl) and resident prophages (PBSX and SPB), which
affected expression of many host genes. Consistent with previous results, induction of these
mobile elements required recA. Induction of the phage appeared to require inactivation of LexA.
Unrepaired UV damage and treatment with MMC also affected expression of some of the genes
that are controlled by DnaA. Furthermore, MMC treatment caused an increase in origin proximal
gene dosage. Our results indicate that different types of DNA damage have different effects on
replication and on the global transcriptional profile.
Introduction
Proper maintenance of genomic content is a major task for all organisms. A variety of
cellular processes are devoted to faithfully replicating and segregating completed genomes prior
to cell division. Cells have also evolved several mechanisms for increasing the chances of
survival by monitoring and responding to the status of genomic integrity. When replication does
not proceed normally these mechanisms are triggered (22, 23, 71). The role of these mechanisms
is to repair the DNA replication defect and to delay subsequent cell-cycle events, such as cell
division, thus increasing the chances of cell survival (7, 22, 23, 27, 43, 71). The effects of these
surveillance mechanisms and subsequent responses are mediated in large part by altering the
transcription of many genes (7, 20, 23, 27).
One of the most well characterized mechanisms that detects DNA damage and replication
arrest is the bacterial SOS response (22, 69). There are two regulatory components to this
response: RecA and LexA. RecA is a highly conserved protein, with homologs present in
eukaryotes (e.g., scRad51, scDmcl) (42, 43). In the cell, there are usually limited amounts of
single stranded DNA (ssDNA) at the replication forks, but when DNA damage occurs, the
amount of ssDNA is thought to increase in both bacteria and eukaryotes (8, 22). RecA binds to
ssDNA exposed during DNA damage and catalyzes recombination processes inside the cell (22,
32, 43). RecA has also been shown to mediate replication fork reversal (a mechanism used to
repair a stalled replication fork on the leading strand), and to alter the accessibility of the 3' ends
of DNA to DNA polymerase (43).
When RecA is bound to ssDNA, it also stimulates the auto-cleavage of LexA (38, 41, 45,
59), also called DinR in Bacillus subtilis (56, 67, 68). LexA is a transcriptional repressor that
binds within the promoter region of target genes as a dimer and prevents transcription from these
promoters (1, 24, 45). Auto-cleavage of LexA relieves repression, thereby allowing transcription
of LexA-repressed genes (1, 45). Subsequent degradation of the cleaved LexA is important, at
least in E. coli, for the proper induction of LexA repressed genes (48).
The composition of the LexA regulon has been the subject of many studies in different
bacteria. Analysis of the global transcriptional response to DNA damage in E.coli established
that treatment with mitomycin C (MMC) causes a large transcriptional effect (approximately
1000 genes) but that only approximately 50 genes are likely to be regulated directly by LexA
(28). The regulation of the remaining genes was speculated to involve other stress-response
transcriptional regulators. Treatment of E. coli with ultraviolet light (UV) showed a similar
number of LexA-regulated genes, but the overall response was smaller than that caused by MMC
(14, 52). These studies concluded that RecA and LexA regulate 50 genes (14). RecA has also
been demonstrated to regulate the activation of mobile genetic elements in E. coli, most notably
the lysogenic phage lambda (57). Similar experiments with MMC treatment in M tuberculosis
also revealed that approximately 50 genes are regulated to some extent in a RecA-dependent
manner following DNA damage (54).
The LexA regulon of Bacillus subtilis has also been investigated (1, 9, 39). Genes known to
be repressed by LexA in B. subtilis are involved in transcriptional regulation (lexA, recA), DNA
repair (uvrAB, uvrC), recombination (recA, ruvAB), and cell division (yneA) (1, 9, 27, 39).
A total of 18 operons (approximately 30 genes) were identified as likely direct targets of
LexA using a computational approach combined with in vitro DNA binding experiments and in
vivo analysis of recA-dependent genes induced by UV irradiation (1). In addition to the genes
repressed by LexA, genes in several lysogenic phages (PBSX, SPB, and phi-105) and the
integrative and conjugative element ICEBs] are also known to be induced by DNA damage in B.
subtilis. (2, 50, 61, 66)
We examined the global transcriptional response to different types of DNA damage and
replication arrest in B. subtilis using whole genome DNA microarrays. We also tested the roles
of recA and lexA in regulating the transcriptional response to various perturbations in DNA
replication. Replication was inhibited directly by inhibiting the PolC (catalytic) subunit of DNA
polymerase with p-hydroxyphenylazo-uracil (HPUra) (6, 23). We also perturbed replication
indirectly by treatment of cells with two different agents, MMC and UV irradiation, that cause
DNA damage. Mitomycin C causes the formation of intra- and inter-strand DNA cross-links and
MMC mono-adducts (17). UV irradiation causes the formation of pyrimidine dimers (22). As
expected, our results show that RecA plays a major part in regulating the transcriptional response
under all of the conditions tested. In addition to the approximately 30 genes inl 8 operons
previously identified as direct targets of LexA (1), we identified another 31 genes in 8 operons
that are also likely to be direct targets of LexA. This brings the total number of operons likely to
be regulated directly by LexA to 26 (approximately 63 genes).
In addition to bacterial genes, we found that many genes from lysogenic bacteriophage and
the mobile genetic element ICEBsl required recA but not lexA for induction, as expected because
RecA facilitates cleavage of many phage repressor proteins (57). However, induction of many of
these genes was blocked in a mutant defective in cleavage of LexA, indicating that LexA plays a
role in the regulation of these mobile elements. The induced phage genes and their indirect
effects comprised more than half of the transcriptional response to replication arrest and MMC
treatment.
In addition to the recA-dependent transcriptional response, replication arrest also induces a
recA-independent response, which is mediated in part by the replication protein DnaA (23). Our
results indicate that replication fork arrest and DNA damage caused by MMC, UV irradiation, or
HPUra all induce a recA-dependent SOS response, but that the recA-independent response to
these perturbations was qualitatively and quantitatively different. MMC also caused a relative
increase of the dosage of the genes near the chromosomal origin of replication. This increase in
gene dosage was most likely caused by a reduced rate of elongation of replication. Our results
demonstrate that cells respond differently to various types of perturbations to DNA replication.
Those differences probably allow cells to respond to each challenge more specifically and
efficiently.
Materials and Methods
General methods and strain construction. B. subtilis strains are listed in Table 1. Genetic
manipulations were performed using standard protocols (25).
Media and growth conditions. For all experiments, cells were grown with vigorous shaking
at 300 C or 370C in S7 defined minimal medium with MOPS (morpholinepropanesulfonic acid)
buffer at a concentration of 50 mM rather than 100 mM (26); the medium was supplemented
with 0. 1%glutamate, the required amino acids (at 40 [tg/ml), and I % glucose. HPUra (a
generous gift from Neal Brown) and MMC (Sigma) were used at a final concentration of 38
4ig/ml and I jig/ml, respectively. For treatment with UV, cultures were transferred to a shallow
dish and irradiated with 25 mJ/m 2 (-50% killing) as previously described (1).
Use of DNA microarrays for gene expression profiling. DNA microarrays were prepared
either using PCR products from >99% of the annotated B. subtilis open reading frames spotted
Table 1. B. subtilis strains used.
Strains
Relevant Genotype; (reference)
JH642
trpC2pheAl; (51)
IRN444
trpC2pheAl
AIG4
trpC2pheAl recA::neo Asda; (23)
AIG105
trpC2pheAl AyneAB::spc; (23)
JJS39
trpC2Aupp; (21)
AIG246
trpC2 Aupp lexAind-; (21)
AIG250
trpC2 Aupp lexA::upp+-K7(phleo); (21)
AIG266
trpC2pheAl lexA::upp+-K7(phleo)AyneAB::spc; (This paper)
YB886
metBlO trpC2 xin-1 SPB0 ICEBslo (phage-defective, pd); (2, 70)
YB3000
metBlO trpC2 xin-1 SPB0 recA260::mls cat ICEBslo (phage-defective, pd); (10)
recA260::mls::cat; (37)
100
onto Corning GAPS slides, or 65-mer oligonucleotide library representing all of the annotated
ORF of the B. subtilis genome (Sigma-Genosys) essentially as described previously (1, 2, 5, 23).
Oligonucleotide microarrays were used only for the lexA(ind) set of experiments (lexA(ind), and
respective isogenic lexA÷ control strain JJS39). The observations from ologonucleotide
microarrays with respect to JJS39 were verified on PCR arrays to demonstrate compatibility
between oligonucleotide and PCR microarray results.
Exponentially growing cultures were treated with HPUra, MMC or UV and samples were
collected immediately before and 15 min, 30 min, and 60 min after treatment and processed as
previously described to generate labeled cDNA (1, 23). A similarly processed reference sample
was hybridized with each experimental sample for normalization. The reference sample
contained pooled total RNA from cell cultures grown in defined minimal medium and cultures
treated with DNA damaging agents, thus ensuring that all genes expressed under those
conditions are represented in the sample. Our microarray analysis includes every spot that has
>80% of the pixels at least one standard deviation over background in one or both Cy3 or Cy5
channels. Since virtually all phage genes are represented in the reference sample (Cy3 channel)
we obtain data for most of these genes even when we compare test samples of stains missing
these genes (phage defective strains for example).
All microarray experiments were done with at least three independent replicates and
evaluated using Significance Analysis of Microarrays (SAM)(65). For all experiments, we
picked the most stringent criteria that resulted in a predicted number of false positives of one or
less. Because the number of statistically significant genes varies from experiment to experiment
but the predicted number of false positives was always 1 or less, the calculated rate of
identification of false positives varied. For most experiments there was less than 1% probability
101
of identifying random effects (false positive discovery rate less than 1.0%). However, due to the
limited number of affected genes in phage cured strain backgrounds, the false positive discovery
rate increased to 2.9% in the MMC or UV treatment experiments. For all experiments we also
introduced cut-off values of 1.5-fold, i.e., only effects that were 21.5-fold were considered
significant and are reported.
Microarray data are deposited at Gene Expression Omnibus
(http://www.ncbi.nlm.nih.gov/geo) database with accession number GSE4673.
Use of genomic microarrays to study DNA replication. The same type of PCR-product
based microarray slides used for expression profiling was used for determining relative
chromosomal content. Asynchronous cultures were treated with MMC or mock-treated, and
samples were collected at various times after the treatment by mixing with an equal volume of
ice cold methanol. Chromosomal DNA was extracted and purified using G-100 Qiagen genomic
DNA purification columns, fragmented by digesting with HaeIII, and purified with Qiagen
QiaQuick PCR purification columns. The DNA fragments were mixed with random hexamers
and heated at 950 C for 5 minutes, then rapidly cooled on ice. Primer extension was conducted at
30'C overnight with Klenow fragment (3' to 5' exo-) and a dNTP mixture including aminoallyldUTP. DNA labeled with aminoallyl-dUTP was purified with Qiagen MinElute columns. Test
and reference samples (DNA obtained from cells with a single unreplicated chromosome) were
coupled to Cy5 and Cy3 dyes respectively, mixed, and hybridized to a microarray using the same
procedures as for expression profiling, to obtain ratios of test sample to reference for each
chromosomal locus. Different test samples were then compared by obtaining the ratio of these
ratios. Presented results are from a single representative experiment.
102
Thymidine Incorporation. Experiments were performed with cells grown to midexponential phase at 370 C. For pulse labeling, [3H] Methyl-thymidine (10 Rl) (80 mCi/mol, 1
mCi/ml, from Perkin-Elmer) was added to 200 pl of culture to a final concentration of 0.05
mCi/ml. The cells were incubated with the radioactive label for Imin at 370 C and were
subsequently mixed with an equal volume of 20% ice cold trichloroacetic acid (TCA) and
allowed to precipitate for at least 30 minutes on ice. Samples were then loaded on glass fiber
filters (GF6, Schleicher & Schuell), filtered with vacuum, washed with 3x10ml of ice cold 5%
TCA, and dried with ethanol. The dried filters were added to scintillation vials, mixed with 10ml
scintillation fluid, shaken vigorously, and processed with a scintillation counter. Background
was determined by treating killed cells in the same manner as the experimental samples.
Background was subtracted from the experimental results.
103
Results and discussion
DNA damage and replication fork arrest cause a broad transcriptional response,
largely mediated by RecA. We compared mRNA levels of virtually all B. subtilis genes in cells
treated with different agents that block replication directly or that cause DNA damage. The goal
was to identify the genes affected under each condition and understand, at a global level, the
types of cellular processes affected by the separate agents. We also sought to identify those genes
induced in a recA-dependent manner under each condition. Our experimental approach was to
treat exponentially growing cultures with the selected agent and compare mRNA levels of each
gene in treated cultures to the levels in mock treated cultures. The majority of changes in mRNA
abundance were expected to be due to changes in the rate of gene transcription, as opposed to
effects on mRNA stability. For each experiment, recA+ and recA- strains were analyzed in order
to determine which changes were recA-dependent.
We arrested DNA replication in exponentially growing cells at a concentration of the drug
HPUra at which replication appears completely inhibited. Comparison of the mRNA levels from
cells treated with HPUra for 60 min to levels from mock-treated cells at the same time point
revealed that the abundance of 668 mRNA species (-17%of all B. subtilis genes) was
significantly affected. Of these 668 genes, the expression of 357 increased, and the expression of
311 decreased (Fig. IA, column 3; Table 2).
When we analyzed the effect of DNA damage 60 min after treatment with MMC, we found
that a total of 464 genes were affected significantly. The expression of 347 genes increased and
117 genes decreased significantly (Fig. lA, column 1; Table 2). Of the 464 genes affected, 334
were also affected by HPUra. Of these 334 genes in common, 266 had increased and 68 had
decreased expression.
104
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Figure 1
105
Figure 1. DNA damage and replication arrest affect the expression of large number of
genes in a recA-dependent manner. The relative mRNA levels in cells treated with MMC,
HPUra, or UV or untreated cells or in various mutants were determined using microarrays.
Samples of parallel treated or untreated cell cultures, or mutant and wild type, were taken at
various times for a direct comparison. Samples were taken during mid-exponential growth and
when treated with HPUra, MMC, or UV, samples were taken 60 min. after treatment. The
average from three biological replicates is presented.
Data are presented as colored boxes (more like lines in panel A), with each box representing
a gene. In A and C, the brightest green represents a 2 8-fold decrease and the brightest red
represents a 2 8.0-fold increase in relative levels of mRNA. In B, the brightest green represents
a >3.5 fold decrease and the brightest red indicates a >3.5 fold increase in relative mRNA levels.
Gray cells indicate no data for that particular gene in the particular experiment. Black indicates
no change.
The letters M, H, and UV above a column indicates treatment with MMC, HPUra, or UV,
respectively. Relevant strain properties are also indicated, with pd- indicating the phagedefective strains.
A. Results are shown for the 798 genes that were significantly affected in wild type by either
HPUra or MMC treatment. Genes are ordered according to their position in the chromosome.
B. Data from the subset of genes that are likely to be directly regulated by LexA are
presented here. These data are extracted from panel A and include the 28 operons whose
expression was significantly affected in the lexA null mutant and that also have putative LexA
binding sites in the regulatory regions. All the previously proposed direct targets of LexA (1) are
also included. Data on expression of the previously proposed direct targets of LexA, in phage-
106
defective strains, both recA+ and recA-, were presented previously (1) and are included here for
completeness. The numeric data for all of the proposed direct targets of LexA are presented in
Table 4.
C. Data from the subset of genes that are in the mobile genetic elements ICEBsl, PBSX, and
SP8 are presented. These data, except for column 24, are from panel A.
columns 1, 14, 22; wild type strain (JH642) treated with MMC compared to the same strain
untreated. columns 2, 19, 29; recA strain (IRN444) treated with MMC compared to the same
strain untreated. columns 3, 15, 23; wild type strain (JH642) treated with HPUra compared to
the same strain untreated. columns 4, 17, 30; recA sda double mutant (AIG4) treated with
HPUra compared to the same strain untreated. Data from these experiments were previously
published (23) and are presented here for comparison. columns 5, 31; Phage-defective (pd-)
strain (YB886) treated with MMC compared to the same strain untreated. column 6; Phagedefective (pd-) recA strain (YB3000) treated with MMC compared to the same strain untreated.
columns 7, 16; Phage-defective (pd-) strain (YB886) treated with UV compared to the same
strain untreated. columns 8, 20; Phage-defective (pd-) recA strain (YB3000) treated with UV
compared to the same strain untreated. columns 9, 25; Aupp strain (JJS39) treated with HPUra
compared to the same strain untreated. columns 10, 18, 26; lexA(incr) Aupp strain (AIG246;
isogenic with JJS39) treated with HPUra compared to the same strain (AIG246) untreated.
columns 11, 21, 27; lexA yneAB strain (AIG266) treated with HPUra compared to the same
strain untreated. columns 12, 13, 28; lexA yneAB strain (AIG266) compared to lexA+ yneAB
strain (AIG105) during exponential growth. column 24; yneAB strain (AIG105) treated with
HPUra compared to the same strain untreated.
107
Table 2. DNA damage and perturbations in replication cause changes in expression of
many genes
a
WT
recA sda
WT
recA
pd-
pd- recA
pd-
pd- recA
HPUra
HPUra
MMC
MMC
MMC
MMC
UV
UV
Total # of genes
668
108
464
80
138
30
35
36
up
357
21
347
69
119
30
35
5
down
311
87
117
11
19
0
0
31
57(20)
57(20)
12(8)
12(3)
4(2)
4(1)
1(1)
8(6)
# of LexA genes
50
0
55
0
31
0
24
0
recA-dependent
-450
# DnaA
regulated
b
overlap
3
13
35
108
32
125
429
oriC prox.
genes up
mobile element d
PBSX (32)
SPB (187)
e
e
ICEBsl (24)e
aThe
0
0
20
35
45
12
0
0
193
0
193
5
2
0
1
0
30
0
30
0
1
0
1
0
149
0
143
5
1
0
0
0
14
0
20
0
0
0
0
0
number of genes whose mRNA levels are significantly affected by a given treatment in the
indicated strain are presented. Strains and data are the same as those in Fig 1. pd- refers to
phage defective strains.
108
bNumber
of genes previously proposed to be directly regulated by DnaA that are significantly
affected under the indicated condition (# of operons in parentheses).
CNumber of genes within -1.5% of the chromosome (-50 kb) on each side of oriC whose
expression was increased after the indicated treatment.
dTotal number of genes that belong to mobile genetic elements (ICEBsl, PBSX, and SPB) that
were affected significantly under the indicated conditions.
e total number of genes that belong to the element.
109
Since RecA is a known regulator of the transcriptional response to DNA damage, we tested
the effect of MMC treatment on cells lacking recA. Of the 464 genes affected by MMC in recA÷
cells, only 35 changed significantly in a recA null mutant after treatment with MMC (Fig lA,
column 2; Table 2). These results demonstrate, as expected, that recA largely mediates the
transcriptional response to MMC.
We previously analyzed the recA-independent, sda-independentresponse to replication arrest
with HPUra and found that only 108 of the 668 genes affected in wild-type (WT) cells were also
affected in a recA sda double mutant (23). Replication arrest negatively affects the expression of
many developmental genes through the induction of sda [(7, 23) Goranov and Grossman
unpublished results]. Sda is an inhibitor of sporulation. It inhibits activation of the transcription
factor SpoOA, thereby affecting expression of at least 100 genes (7, 46, 58). These results
indicate that of the 668 total genes affected by HPUra, recA affects the expression of
approximately 450 genes, sda affects -100, and 108 are independent of both sda and recA. These
results provide further evidence that RecA is a main, but not the only, contributor to the
transcriptional response to DNA damage and perturbations in replication. We also note that some
of the differences in gene expression between wt and recA strains may be due to more extensive
DNA degradation which happens in the recombination deficient recA cells (13).
LexA represses 26 operons containing 63 genes. Since RecA is a major regulator of the
transcriptional response to DNA damage and replication arrest, we were interested in
determining how RecA affects the expression of so many genes. RecA is known to affect gene
expression, in part, by catalyzing the auto-cleavage of the repressor LexA (45) and the cleavage
of some phage repressors (57). To address how many genes are directly repressed by LexA, we
identified genes that have increased expression in a lexA null mutant. This analysis was
110
performed in a yneAB-null mutant. yneA encodes a division inhibitor that is repressed by LexA
(27). In the absence of lexA, cells filament and grow poorly in minimal medium due to
expression of yneA. In our strain background, deletion of yneA suppresses the lexA growth defect
in minimal medium (data not shown).
Expression of 139 genes increased significantly in the lexA yneAB mutant as compared to the
lexA+yneAB strain (Fig. 1, columns 12, 13, 28; Table 4). We searched the 300 bp region
upstream of the start of these genes for the relaxed LexA binding site consensus sequence
GAACnnnnGTTC (where n=A, G, T, or C) (1, 9), allowing for one mismatch. Twenty-six of the
genes (all were the first gene in their putative operon) had at least one recognizable LexA
binding site with no more than one mismatch to the consensus sequence. These 26 operons
accounted for approximately 70 of the 139 genes whose expression was increased in a lexA null
mutant. Two other operons have also been reported to be regulated by LexA: aprX and ybaKcwlD (1). Au et al demonstrated that LexA bound these promoters in vitro. Though the
expression of these two operons was not affected significantly in our lexA null vs lexA+
comparisons, they did exhibit recA-dependent regulation (1).
Since LexA is inactivated in a RecA-dependent fashion, then the operons that we suspect to
be LexA-repressed should be induced by DNA damage and replication arrest in a recAdependent manner. We examined whether the 28 operons we suspect to be LexA-repressed are
induced by DNA damage and replication arrest in a recA-dependent manner in the experiments
above (Fig. 1). Twenty-six of the proposed LexA-repressed operons were significantly induced
under at least one of these conditions in wild type cells and were not induced in a recA null
mutant (Fig. 1, Table 4). The other two operons, yokEF and dltABCDE, were not affected by
HPUra, MMC, or UV.
111
We also tested the effects of the lexA null mutation on global changes in gene expression in
response to replication arrest. Over 600 genes had altered expression 60 min after replication
arrest (Fig 1A, column 11). These results indicate that, as expected, there are several regulatory
factors other than LexA that control gene expression in response to replication arrest.
In the lexA null mutant, LexA-repressed genes are constitutively expressed and replication
arrest should have little or no effect on their expression, unless there are other mechanisms
controlling expression of these genes. We found that of the 26 operons proposed to be regulated
directly by LexA, only the genes that belong to SPB were induced 60 min after replication arrest
with HPUra in the lexA null mutant (Fig. 1, columns 11, 21, 27; see below).
There are mutations in lexA, lexA(ind), that prevent (or greatly reduce) autocleavage of
LexA, but still allow LexA to function as a repressor (21). Genes that are repressed by LexA
should not be induced in the lexA(ind) mutant in response to DNA damage or arrest of DNA
replication. We arrested replication elongation with HPUra in lexA(ind) strains and performed
microarray analysis to assess the effects on global gene expression. Of the 26 operons regulated
by LexA, only tagC (dinC) was increased significantly in the lexA(ind) mutant 60 min after
replication fork arrest (Fig. 1B, column 18). The change in tagC expression was less than 20%
that in lexA + cells. The tagC promoter is one of the most highly induced promoters after DNA
damage (Table 4) and is likely extremely sensitive to LexA levels. We speculate that there might
be some limited proteolysis of the LexA(ind) protein and that the most sensitive promoters
become slightly induced. Furthermore, additional results indicate that the partial induction of
tagC is not due to lexA-independent regulation of tagC by DNA damage (see above).
Based on our results, we propose that a total of 26 operons containing at least 63 genes are
directly repressed by LexA, as these 26 operons displayed lexA- and recA-dependent induction
112
Table 3. MMC and UV treatments affect a few of the genes proposed to be regulated by
DnaA.
Putative DnaA-regulated genes that are also affected by the indicated treatment (fold-effect) wt
recA
pd-
pd- recA
pd-
pd- recA
Treatment
MMC
MMC
MMC
MMC
UV
UV
# operons
8
3
2
1
1
6
citZ (2.1)
ymaA (3.4)
yqeH (-2.4)
ymaA(3.7)
ywfO(2.3)
aroD (-1.7)
Genotype
b
flgB (-1.7)
nrdE (3.4)
kdgR (-1.9)
nrdF(3.5)
nap (-2.0)
ymaB (3.1)
nrdE (1.7)
pyrAA (-18)
ywfO (3.2)
ywgA (3.0)
nrdE (3.5)
dnaA (-3.5)
dnaN (-3.6)
nrdF(3.9)
ymaB (3.5)
sda (2.2)
yllB (-2.0)
nrdF(2.4)
pyrB (-21)
ylxA (1.6)
ymaB (1.9)
pyrC (-15)
ywfO (2.3)
yclN (-2.1)
pyrF (-7.6)
yclQ (-2.0)
pyrK (8.5)
yybB (-1.5)
ywgA (2.5)
pyrD (-5.8)
yclP (-1.8)
ywfO (1.6)
a Of the
aroD(-2.7)
ywfO (2.2)
ywgA (2.3)
57 genes proposed to be regulated directly by DnaA (23), only those that are also
affected by the indicated treatment are indicated. Numbers in parentheses indicate the foldeffects, with negative numbers indicating decreased expression.
bRelevant genotype is indicated. Strains used were as indicated in Figl.
c Genes that are internal in an operon are indented.
113
after replication arrest or DNA damage. These 26 operons include the 18 operons previously
proposed by Au et al. to be regulated directly by LexA. Although two operons, yokEF and
dltABCDE, had increased expression in lexA null mutants and contained putative LexA binding
sites, they were not induced by DNA damage. We suspect that they are probably not directly
regulated by LexA, and instead are indirectly affected by the constitutive expression of LexA
regulated genes.
Of the 8 additional operons we propose to be LexA regulated, one had previously been
proposed to be LexA repressed: dnaE-an essential, error-prone DNA polymerase (16, 36).
Another operon, yqjH (polY1), encodes a Y family DNA polymerase that has been demonstrated
to have a role in stationary phase mutagenesis and can interfere with replication (18, 19, 63). In
addition, the genes in the yozKL operon are also similar to the Y-family polymerases, so their
regulation by DNA damage is not surprising, though their role in repair of DNA damage is yet to
be determined. At least 5 of the proposed LexA-repressed operons (23 genes) are a part of the
prophage SPI3 in the B. subtilis genome: yolD-uvrX,yokHIJKL, yopTUVXYZ-yoqABC, yorHI,
yorBCDEFG(35). The presence of potential LexA binding sites in the promoters of these 5 SPB
operons indicates that LexA may be directly involved in their regulation. Indeed, LexA also
directly regulates at least one gene, xkdA, in the genome of another B. subtilis phage, PBSX (Fig.
1, (1)). Also one of the proposed LexA regulated SPB genes, uvrX, is homologous to the Y
family DNA polymerases [e.g. E.coli umuC, B.subtilisyqjW[(polY2), yqjH(polY1)] which are
regulated by LexA in both E. coli and B. subtilis (Fig. 1, (47, 60, 63)).
LexA is also known to directly regulate gene expression in other bacteriophages. In
coliphage 186, lytic gene expression is repressed by the phage repressor, cI (34). The activity of
cl is antagonized by an antirepressor, Tum, whose transcription is repressed by LexA (33). In the
114
Vibrio cholerae CTX prophage, LexA binds directly to a promoter required for phage
development (53). Removal of LexA repression results in increased expression from this
promoter and subsequent prophage induction (53).
In B. subtilis, additional genes also appear to be regulated by LexA indirectly. Of the 139
genes that increased in expression in lexA null mutants, 75 had no discernable LexA binding sites
and are probably controlled indirectly by LexA. Approximately half of these genes are in SP3.
Most of the 75 genes were also induced in a recA-dependent manner after MMC or HPUra
treatment (Fig. 1; and not shown). None of the genes repressed by LexA appears to be a
transcriptional regulator, but many are involved in DNA metabolism and can interfere with
replication (64). It is plausible that the interference with replication by LexA regulated genes
causes many of the detected changes in gene expression. Interference with replication also might
be how the tsi-23 mutation (gene not known) causes temperature induced SOS related
phenomena, most of which are recA-dependent (39). tsi-23 [22% linked to air (dal) locus (39)]
does not map near any of the LexA regulated genes described here.
Some of the genes encoding DNA recombination and repair proteins are induced during the
development of genetic competence (4, 15, 39, 40, 49, 55). However, most of the LexAregulated genes described here are not induced during competence development, the exceptions
being recA, lexA, yneAB, tagC, and dinB.
Induction of phage gene expression by DNA damage and replication arrest requires
RecA and the inactivation of LexA. Our results indicate that a large set of genes induced by
DNA damage in B. subtilis belong to the genomes of the prophage PBSX (32 total genes) and
SPB (187 total genes), and to the integrative and conjugative element ICEBsl (24 total genes).
115
Sixty min after treatment with HPUra or MMC, 193 of the 243 genes associated with PBSX,
SPB, and ICEBs] were induced (Fig 1C, columns 22, 23; Table 2).
The induction of genes in these elements was dependent on recA as phage and ICEBs1 genes
were not induced in recA null strains (Fig. 1C, columns 29, 30 (compared to columns 22, 23);
Table 2). However, unlike most LexA-repressed genes, some phage and ICEBs] genes were still
induced by HPUra in a lexA null strain (Fig. IC, column 27), indicating that genes in these
elements are regulated, at least in part, independently of LexA. The induction of SPB genes by
HPUra in a lexA strain was less than that in WT, at least in part, due to the increased background
expression level of SPB genes in untreated lexA" cultures (Fig. IC, columns 23, 27). It is likely
that the SPB induction in untreated lexA- cells is due to the constitutive expression of LexArepressed genes.
The induction of at least one LexA-repressed gene must play a role in the induction of PBSX
and SP8 gene expression, as their induction was prevented in a non-cleavable lexA(ind) mutant
(Fig. IC, column 26). Since the lexA(ind) mutation was in a different genetic background, we
confirmed that HPUra induces the expression ofprophage genes in the isogenic lexA+ strain (Fig.
1C, column 25). Curiously, in that strain background we saw no induction of ICEBs1 by HPUra
in either lexA + or lexA(ind) mutant, though ICEBsl was present in the genome (data not shown).
The explanation of this observation will require further study.
Our results are consistent with previously published reports that showed that the induction of
SP8 and ICEBsl is recA-dependent (2, 44). These mobile genetic elements are regulated
similarly to several other phages and the integrative and conjugative element SXT, which are
activated by the recA-dependent DNA damage response (3, 57). It is thought that this regulation
116
Table 4. LexA appears to repress approximately 63 genes in 26 operonsa
19
20
17
18
15
16
13
14
Name
21
Description
aprX
ybaK
1.2
1.1
1.8
1.7
1.6
1.3
1.9
2.6
1.0
1.0
-2.7
1.1
1.1
1.0
1.1
-1.2
1.2
1.1
cwlD
dinB
1.2
32.2
1.5
27.1
1.3
29.3
2.2
40.3
1.1
1.2
1.0
1.7
1.1
-1.2
1.1
-1.7
1.4
1.1
intracellular alkaline serine protease
unknown; similar to unknown proteins
N-acetylmuramoyl-L-alanine amidase
(germination)
nuclease inhibitor
dltAb
dltBb
1.7
1.9
-1.2
-1.1
-1.5
-1.4
1.1
1.1
1.1
1.1
-1.9
1.3
1.1
1.2
1.2
1.2
-1.3
-1.4
dltCO
1.8
1.0
-1.3
1.0
1.2
1.5
1.3
1.3
-1.1
dltDb
dltE"
dnaE
lexA
pcrA
ligA
yerH
1.7
1.6
1.5
4.5
1.7
1.8
1.6
1.0
1.1
1.3
1.4
1.3
1.4
1.0
1.1
1.0
2.1
1.9
1.8
1.6
-1.5
-1.3
1.3
2.8
2.3
2.8
2.2
-1.1
1.2
4.4
1.6
1.9
1.5
1.3
1.0
-1.3
1.2
1.2
1.2
1.4
1.3
-1.1
1.5
1.2
1.3
1.3
1.1
-1.5
1.4
1.4
1.3
1.4
1.1
-1.3
1.1
1.3
1.3
1.0
-1.3
-1.5
1.1
1.1
1.1
recA
parE
parC
ruvA
ruvB
queA
3.7
1.7
2.1
2.5
2.2
2.4
7.5
1.6
2.0
1.7
1.6
1.9
6.9
1.4
2.0
2.0
2.3
1.8
6.7
1.9
2.1
2.1
2.2
2.1
1.1
-1.1
1.0
-1.2
1.1
1.0
1.3
1.1
1.1
1.5
1.0
1.3
1.0
1.2
1.2
1.1
1.1
1.2
1.0
1.0
-1.2
1.1
1.2
1.2
-1.2
1.0
1.0
-1.3
-1.2
-1.3
tgt
2.2
1.9
2.2
2.1
1.0
1.4
1.1
1.3
1.0
tagC
uvrA
uvrB
uvrC
17.1
8.4
7.7
1.5
37.8
7.9
9.2
1.5
31.3
9.6
7.5
1.3
53.8
8.3
6.8
1.6
1.0
1.2
1.1
1.2
3.0
1.3
1.2
1.1
-1.3
1.2
1.3
1.3
1.1
1.2
1.0
1.0
1.8
1.0
1.1
1.0
involved in lipoteichoic acid biosynthesis
DNA polymerase III (alpha subunit)
transcriptional repressor of the SOS regulon
ATP-dependent DNA helicase
DNA ligase (NAD-dependent)
unknown; similar to unknown proteins
multifunctional protein involved in
homologous recombination and DNA repair
(LexA-autocleavage)
subunit of DNA topoisomerase IV
subunit of DNA topoisomerase IV
Holliday junction DNA helicase
Holliday junction DNA helicase
S-adenosylmethionine tRNA
ribosyltransferase
tRNA-guanine transglycosylase
possibly involved in polyglycerol
phosphate teichoic acid biosynthesis
excinuclease ABC (subunit A)
excinuclease ABC (subunit B)
excinuclease ABC (subunit C)
yolD
10.2
5.7
4.6
1.1
-1.0
1.5
1.0
1.1
-1.6
unknown
uvrX
xkdA
8.5
6.9
5.9
4.5
4.9
2.8
1.1
2.6
1.0
1.1
-1.7
1.1
1.0
1.0
1.0
-1.2
-1.6
1.4
ydiO
1.7
2.7
2.1
2.5
1.1
1.1
1.7
1.2
1.2
ydiP
yhaO
yhaN
yhaM
yhaZ
yhjD
1.9
3.1
1.1
1.7
19.9
10.5
2.8
2.7
1.1
1.5
8.0
7.8
2.1
2.4
1.1
1.5
6.4
7.0
2.8
3.2
1.3
1.9
10.8
7.7
1.1
1.0
1.0
1.1
1.1
1.1
1.3
1.2
1.1
1.1
1.3
1.5
1.8
1.1
-1.2
1.0
-1.3
1.1
1.3
-1.1
-1.2
1.0
-1.5
-1.4
1.1
1.1
-1.2
-1.1
-1.6
1.0
UV-damage repair protein
PBSX prophage
unknown
similar to DNA-methyltransferase
unknown
similar to DNA-methyltransferase
unknown; similar to unknown proteins
unknown; similar to unknown proteins
unknown; similar to CMP-binding factor
unknown; similar to DNA repair enzyme
unknown; similar to unknown proteins
yhjC
yhjB
yneA
1.5
1.5
94.8
1.2
1.3
20.4
1.6
1.9
19.7
1.6
1.8
24.9
-1.0
1.3
1.1
1.1
1.0
1.6
1.3
1.5
1.0
1.3
1.3
-1.1
1.0
1.1
-1.6
unknown
unknown
unknown
yneB
ynzC
31.9
8.2
16.0
4.0
9.0
4.6
11.9
4.3
1.0
1.1
1.9
1.2
-1.3
1.0
1.4
-1.2
-1.5
-1.2
unknown; similar to resolvase
unknown; similar to unknown proteins
yokEb
yoken
yopT
yopU
1.7
1.7
3.0
2.3
1.1
1.1
2.8
2.3
-1.5
-1.6
1.9
2.1
1.2
1.4
-1.0
-1.3
1.1
1.1
-1.2
1.1
1.0
1.2
-1.1
-1.2
1.3
1.3
1.2
1.2
1.0
1.0
1.0
-1.1
-1.2
-1.2
1.1
1.1
unknown
unknown
unknown
unknown
117
D-alanyl-D-alanine carrier protein ligase
D-alanine transfer from Dcp to
undecaprenol-phosphate
D-alanine carrier protein
to the poly(glycerophosphate) chain
yop V
yopW
yopX
yop Y
yopZ
yoqA
yoqB
yoqC
yoqH
yoql
2.9
3.4
2.5
1.4
2.3
2.1
2.5
2.3
1.7
1.8
15.0
14.4
16.3
8.4
15.3
19.5
13.9
18.8
5.9
5.2
16.4
13.7
7.1
5.0
9.2
12.6
21.5
17.9
12.6
9.5
-1.2
-1.5
1.2
1.1
1.1
-1.3
1.3
1.1
-1.3
1.0
1.3
1.0
1.0
1.1
1.0
1.0
1.0
1.2
1.0
1.0
1.4
1.0
1.3
1.6
1.4
1.9
1.7
1.1
1.2
1.7
1.5
1.2
1.7
1.0
1.6
1.6
1.3
1.6
1.4
1.1
1.1
1.1
1.2
1.2
1.1
-1.1
1.1
1.0
-1.1
-1.1
1.3
1.9
1.7
1.1
1.6
1.7
1.8
2.0
1.9
1.8
unknown
unknown
unknown
unknown
unknown
unknown
unknown
unknown
unknown
unknown
yoqJ
yoqK
yoqL
yorB
yorC
yorD
yorE
yorF
yorH
1.8
1.6
2.0
62.9
44.1
4.5
3.2
2.1
2.3
17.2
3.9
14.7
88.0
33.6
9.8
19.4
67.7
54.9
14.1
1.0
18.1
34.6
28.9
3.7
30.9
70.5
38.0
-1.3
1.0
1.1
1.6
-1.4
1.9
-1.4
1.0
1.4
1.1
1.0
1.1
1.0
1.0
-1.7
1.1
1.0
1.2
1.5
1.5
1.0
1.1
1.0
1.0
1.2
2.2
1.3
1.4
1.3
1.2
1.2
1.2
-1.5
1.6
2.3
2.4
1.1
1.4
-1.1
1.0
1.0
1.0
1.1
1.0
1.1
1.8
1.5
2.3
1.5
1.4
1.0
2.5
3.9
3.2
unknown
unknown
unknown
unknown
unknown
unknown
unknown
unknown
unknown
yorl
1.1
8.0
15.0
1.0
-1.2
1.3
-1.2
1.1
2.0
unknown; similar to DNA helicase
yozL
10.1
4.3
5.9
1.0
1.0
1.4
1.3
1.2
-1.6
unknown
yozK
yqjH
4.0
1.6
2.2
1.1
-1.0
1.2
1.6
2.0
1.0
1.0
1.4
1.2
1.1
1.0
1.1
1.0
-1.6
-1.3
unknown; similar to DNA repair protein
unknown; similar to DNA repair protein
yqj W
yqjX
1.7
3.1
2.1
3.4
2.0
4.7
2.8
4.3
1.1
1.0
-1.1
1.0
1.1
1.2
1.1
1.0
1.2
1.0
unknown
unknown
yqjY
yqjZ
1.3
1.4
1.0
1.0
-1.1
1.0
1.5
1.5
1.1
1.1
1.1
1.1
-1.4
-1.4
-1.1
1.1
1.1
1.0
unknown; similar to unknown proteins
unknown; similar to unknown proteins
aColumns same as in Figure 1. Numbers are normalized average
fold effects in mutant or treated culture. Negative numbers represent negative regulation
(e.g. 0.5 ratio is represented as -2.0)
bBold genes are not considered as LexA-regulated
118
allows the mobile genetic element a better chance of survival through leaving if the host cell is
being damaged.
We also found that expression of genes in PBSX and SPB is partially controlled by the
repressor, LexA, indicating that the full expression of at least one LexA repressed gene is
necessary for their induction of the phage genes. It is likely that the lack of increased expression
of recA, or the insufficient activation of RecA due to low abundance of ssDNA in the lexA(ind)
strain prevent full induction of phages (22). In addition, LexA also inhibits expression of at least
one operon in PBSX and SPB, and phage induction might be partly inhibited if these operons are
not induced. We were unable to determine whether LexA regulates the expression of genes in
ICEBsl, as this element was not induced in a lexA + strain isogenic to the lexA (ind-) strain. Our
data also indicates that LexA inactivation alone is not sufficient to trigger the full expression of
most of the genes of the mobile elements. This is a key difference between the majority of the
genes of the mobile elements and the direct targets of LexA.
We also observed that expression of 15 to 20 genes from the skin element had apparently
increased in expression after HPUra and MMC treatment (Fig lA, columns 1, 3). The skin
element is a defective prophage that disrupts a gene important for sporulation, sigK, and precise
excision of skin during sporulation restores the sigK open reading frame (31, 62). The increase in
expression of skin element genes was detected only for genes with high similarity to genes in
PBSX and was not detected when the same RNA was hybridized to spotted oligonucleotide
arrays (data not shown). Our interpretation of these results is that mRNA from some of the
PBSX genes, which are induced in response to DNA damage, cross-hybridize with the probes to
skin element genes, thus giving the appearance of up-regulation of the skin element. Consistent
with this, previous reports showed that the skin element is not induced by MMC treatment (30).
119
Phage gene expression and its indirect effects constitute a large portion of the genes
induced by DNA damage and replication arrest. As outlined above, many of the genes
induced in WT cells after DNA damage belong to prophage. The induction of phage genes is the
first step towards assembling phage particles and lysing the host cell to release these particles.
PBSX and SPB are both capable of assembling such particles and releasing them (50, 61, 66).
The induction of phage genes and subsequent particle assembly almost certainly have many
effects on cellular metabolism that could cause indirect effects on the host gene expression. We
were interested in assessing the transcriptional profiles of cells unable to induce the prophage
genes after DNA damage, thus eliminating any secondary effects we might observe due to phage
induction. We used a strain (YB886) that lacks SPB and cannot induce PBSX due to a mutation
xin-1(70), and also is missing ICEBs] (Auchtung and Grossman unpublished data).
In the phage-defective strain (YB886), 60 min after addition of MMC, in conditions identical
to the experiments outlined above, 138 genes were significantly affected as compared to the 464
genes affected by MMC in a phage + background. Of the 138 genes, expression of 119 increased
and expression of 19 decreased following treatment with MMC. This response was primarily
mediated by RecA, as the effect of MMC on most genes (125 of the total 138 affected genes)
was absent in the recA null mutant (Table 2).
The reduction of the overall number of genes affected by MMC in the phage-defective strain
indicates that there is a large effect of phage induction on host gene expression. Our analysis
indicates that besides the 198 phage and ICEBs1 genes, there were 184 non-phage genes that
were affected in the wild type but not in the phage-defective background after MMC treatment
(e.g., Fig. 1A, compare columns, I and 5; Table 5). Changes in expression of most of the host
genes were dependent on both recA and the presence of the phage (Table 5). These non-phage
120
Table 5. Phage induction affects the expression of many non-phage genes'
1
2
5
6
3
4
10
9
MMC
MMC
MMC
MMC
HPUra
HPUra
HPUra
Name
veg
WT
recA
pd-
pd- recA
WT
recA sda
HPUra
lexAindupp
Description
3.1
1.8
3.1
1.2
-1.2
-1.3
upp
-1.2
lexA
1.8
1.1
sspF
sigW
ybdO
ybeF
-1.7
-1.2
1.1
1.1
1.0
1.0
1.0
-1.2
-1.5
1.0
-1.2
1.1
-2.3
-1.3
-1.3
-1.6
-1.1
1.1
1.1
-2.3
1.1
-2.0
ybfG
1.7
21.5
28.5
119.0
1.0
1.4
1.5
1.4
1.5
1.5
1.6
1.2
-1.4
1.1
1.5
1.1
7.6
14.7
77.4
1.1
1.0
1.0
1.2
-1.0
1.2
1.0
-1.3
-1.9
1.0
6.4
1.0
1.0
1.0
1.2
-1.4
1.4
1.2
1.0
1.0
function unknown
small acid-soluble spore protein
(minor alpha/beta-type SASP)
gamP
yceK
tlpC
yclE
-2.3
-1.5
-1.8
2.5
-2.0
-1.1
1.0
1.2
1.0
1.0
-1.6
1.2
-2.5
-1.2
-1.3
1.1
1.1
1.1
-1.7
2.6
-1.3
1.1
1.1
1.1
1.1
1.0
1.0
1.2
-1.6
-1.2
-1.8
1.6
1.0
1.0
-4.3
1.0
yclF
ycnl
2.2
-1.4
-1.2
-1.2
1.3
1.1
-1.3
-1.4
1.6
-1.2
1.0
-1.3
1.1
-1.3
-1.2
-1.8
1.2
1.3
ycnK
ycsI
ydcO
ydfF
groEL
pspA
-1.5
-1.4
1.1
-1.5
-1.4
-1.6
-1.2
ydjH
-1.8
81.1
2.9
1.5
-2.2
-2.2
-1.3
1.3
1.0
1.3
-1.1
-1.3
1.1
1.3
1.1
1.2
-1.7
-1.9
-1.3
1.6
-1.2
1.2
-1.3
-1.5
1.4
7.9
4.2
1.8
-2.6
-2.8
1.1
1.1
-1.2
1.4
1.1
-1.2
1.0
1.1
-1.0
1.1
-1.1
-1.0
ydjI
yefC
yeeA
-2.2
1.8
2.2
-1.3
2.5
2.5
-2.0
1.5
1.3
-1.5
3.0
3.1
-2.7
1.5
1.8
-1.2
1.3
1.5
-1.0
1.5
1.5
-2.1
-1.1
24.3
-1.4
1.0
-1.7
-1.8
-1.6
2.4
2.3
1.4
-1.1
1.3
1.0
-1.3
1.2
1.2
1.2
1.2
1.2
yfmT
yfmS
-2.1
-2.2
-1.2
-1.3
-1.5
-1.6
-1.3
-1.5
-1.8
-1.8
1.1
1.1
1.3
1.2
1.1
1.0
1.1
1.0
yfmP
1.8
1.0
1.4
1.1
1.2
1.0
1.1
1.7
1.3
nagP
-2.9
-2.2
-1.3
-3.1
-1.7
-1.2
-1.5
-1.8
1.1
yfiJ
-1.7
-1.3
-1.3
-1.3
-1.4
1.0
1.1
-1.2
1.1
yfiL
yhbA
glpF
yhdE
hemZ
hemZ
yhflU
comK
wprA
yisP
yjaZ
cotW
yjcM
-1.8
1.4
1.6
-1.6
1.8
1.0
-2.0
-2.1
-1.6
-1.7
2.6
3.7
-4.2
-1.4
1.3
1.1
-1.3
1.1
-1.2
1.2
-1.9
-1.4
-1.3
1.0
1.3
-1.8
1.1
1.3
1.5
-1.2
1.4
1.1
-1.5
-2.1
-1.8
-1.2
1.2
1.2
-3.6
-1.5
1.0
1.1
-1.2
1.2
1.1
1.1
-1.7
-1.6
-1.4
1.0
-1.2
-1.9
-1.4
1.1
1.0
-1.8
1.6
1.0
1.1
-2.2
-2.1
1.1
1.1
2.9
-7.9
1.0
-1.2
-1.2
-1.5
1.0
1.0
1.0
-1.4
1.1
1.0
1.0
1.1
1.1
1.0
-1.2
1.1
1.1
1.1
-1.0
-1.8
1.2
1.1
-1.2
-1.0
1.0
1.2
-1.2
-1.3
1.3
1.3
1.0
-2.1
-1.4
-1.1
-1.2
1.0
-1.8
1.2
1.1
1.3
1.1
1.3
1.3
-1.6
-2.6
1.4
1.0
-1.2
-3.3
1.2
ybfE
ybJF
-2.0
121
RNA polymerase ECF-type sigma factor
unknown
unknown; similar to unknown proteins
unknown
unknown
unknown; similar to unknown proteins
IICBA component
unknown
methyl-accepting chemotaxis protein
unknown
unknown; similar to di-tripeptide ABC
transporter
unknown; similar to unknown proteins
unknown; similar to transcriptional regulator
(DeoR family)
unknown; similar to unknown proteins
unknown
unknown; similar to unknown proteins
class I heat-shock protein (chaperonin)
phage shock protein A homolog
unknown; similar to unknown proteins
unknown; similar to unknown proteins
unknown; similar to resolvase
unknown; similar to unknown proteins
unknown; similar to benzaldehyde
dehydrogenase
unknown
(MerR family)
putative PTS N-acetylglucosaminespecific enzyme IICB component
unknown; similar to two-component
sensor histidine kinase
unknown; similar to ABC transporter
(ATP-binding protein)
unknown; similar to unknown proteins
glycerol uptake facilitator
unknown; similar to unknown proteins
coproporphyrinogen III oxidase
coproporphyrinogen III oxidase
unknown; similar to biotin biosynthesis
competence transcription factor (CTF)
cell wall-associated protein precursor
unknown; similar to phytoene synthase
unknown; similar to unknown proteins
spore coat protein (insoluble fraction)
unknown
---
yjcN
yjcO
yjdF
-3.7
2.1
-2.1
-1.8
1.1
-1.2
-1.9
1.0
-1.8
-1.9
1.0
-1.3
-3.3
1.0
-1.8
-1.5
1.0
-1.2
-2.5
1.1
-1.2
-1.9
1.6
-1.2
1.2
1.2
1.0
xlyB
yjqA
20.0
-1.2
1.1
-1.2
1.0
1.1
1.0
1.1
1.4
1.1
1.1
1.1
10.1
5.6
10.7
3.9
unknown
N-acetylmuramoyl-L-alanine amidase
(PBSX prophage-mediated lysis)
3.9
1.1
unknown; similar to unknown proteins
yjqB
-1.2
-1.3
-1.2
1.2
1.1
-1.3
2.6
3.8
-1.1
1.1
1.1
1.0
1.9
2.8
1.0
unknown; similar to phage-related
replication protein
yjqC
5.0
8.9
1.5
unknown
dppC
proA
ispA
ispA
yko W
sigI
ykrK
spoOE
2.0
-3.0
-1.5
1.4
-2.5
-1.6
-1.7
-2.6
1.2
-1.9
1.1
1.0
-1.3
1.1
-1.1
-1.7
1.1
-1.8
1.4
1.2
-2.1
-1.2
-1.2
-1.7
1.2
-2.2
-1.1
-1.3
1.2
1.0
-2.1
1.8
-1.6
1.0
1.6
-2.3
-1.5
-1.4
-1.8
1.4
-1.3
1.0
1.2
1.1
-1.4
1.1
-1.6
1.0
1.1
1.0
1.0
1.1
1.0
1.2
-1.5
1.8
-1.8
1.0
1.0
-1.3
-1.4
1.0
-2.2
1.1
1.0
-1.2
-1.2
1.0
-1.2
-1.2
-1.2
kinD
-1.9
-1.1
-1.3
1.1
-1.7
-1.5
-1.2
-1.7
1.1
motB
motA
-2.4
-2.2
-2.6
1.8
-1.2
1.0
-1.5
1.0
-1.8
-1 .9
-1.9
1.2
-1.3
- 1.2
-2.8
-2.7
1.1
1.1
-1.1
1.1
-1.1
-1.2
1.0
1.1
motility protein (flagellar motor rotation)
motility protein (flagellar motor rotation)
-1.7
-3.1
1.1
-1.3
-1.6
1.1
unknown
1.2
1.5
1.0
1.0
-1.1
1.2
unknown
6.5
3.9
1.2
1.6
1.0
1.3
1.5
-1.2
1.0
1.1
unknown
1.2
-1.1
1.1
1.1
1.0
-1.7
unknown; similar to unknown proteins
3.3
1.9
2.6
2.2
1.4
1.1
1.9
1.6
1.1
2.6
3.0
1.0
-1.1
unknown; similar to unknown proteins
s
tran criptional regulator mediating
ykuR
ykuT
ykuU
yku V
yknT
2.0
2.7
2.7
2.7
6.7
1.0
1.6
1.1
1.0
1.1
1.0
1.2
1.0
1.0
1.2
unknown
unknown; similar to unknown proteins
unknown
unknown; similar to unknown proteins
unknown
mobA
moeB
moeA
2.6
2.0
2.0
1.1
1.2
1.2
mobB
moaE
moaD
yknU
ykn V
ctaB
ylbF
2.1
2.0
2.0
2.5
6.6
-2.0
-1.6
sigE
ykuA
ykuF
ykuH
ykul
ykuL
ccpC
-1.3
unknown
unknown
dipeptide ABC transporter (pennease)
(sporulation)
gamma-glutamyl phosphate reductase
major intracellular serine protease
major intracellular serine protease
unknown; similar to unknown proteins
RNA polymerase sigma factor
unknown
negative sporulation regulatory
phosphatase
ens
two-component s or histidine
kinase involved in the initiation
of sporulation
1.8
1.1
-1.2
1.0
1.0
1.1
1.0
-1.1
1.2
-1.1
1.0
1.0
1.1
1.5
-1.2
1.1
2.2
1.1
1.2
1.1
1.1
1.0
1.3
1.4
-1.6
-1.1
1.0
1.1
2.0
1.1
1.0
1.2
1.0
1.1
1.0
-1.2
-2.7
1.0
1.0
1.0
1.4
1.3
1.3
1.6
1.4
1.4
1.0
1.1
1.0
1.2
1.2
1.3
1.4
1.2
1.4
1.0
1.1
1.0
1.2
1.2
1.1
1.1
-1.1
-1.5
1.1
1.0
1.0
1.2
1.1
-1.3
-1.4
-1.1
1.3
1.3
1.3
-1.2
-1.1
-1.5
1.0
1.4
1.3
1.5
1.5
1.5
-1.9
-1.3
1.0
1.0
1.4
-1.1
1.0
-1.2
-1.2
1.3
1.3
1.3
1.1
1.2
1.1
1.0
1.4
1.3
1.4
-1.2
1.0
1.1
1.0
1.1
1.1
1.1
-1.3
1.1
1.1
1.1
2.7
1.5
1.2
1.0
1.1
1.8
1.1
-1.6
-1.8
sigG
cysH
3.0
-2.1
1.4
-1 .4
1.3
1.0
1.3
1.1
1.6
1.0
1.9
1.1
1.0
1.1
- 1.2
1.1
-2.4
1.1
ylqB
-2.7
-2.0
-1.8
-1.9
-4.8
1.0
-2.3
-1.6
-1.2
unknown
ylqD
topA
flgC
fliF
1.5
-2.2
-1.8
-1.8
1.0
-1.1
-1.2
1.1
1.2
-1.6
-1.7
-1.9
1.2
-1.1
-1.2
-1.2
1.0
-1.8
-2.1
-2.2
1.0
-1.1
-1.3
1.1
1.1
1.0
1.0
1.0
1.0
-1.5
-1.4
1.0
1.0
-1.7
1.0
1.0
unknown; similar to unknown proteins
DNA topoisomerase I
flagellar basal-body rod protein
flagellar basal-body M-ring protein
carbon catabolite repression
molybdopterin-guanine dinucleotide
122
biosynthesis
molybdopterin biosynthesis protein
molybdopterin biosynthesis protein
molybdopterin-guanine dinucleotide
biosynthesis
molybdopterin converting factor
molybdopterin converting factor
unknown; similar to ABC transporter
unknown; similar to ABC transporter
cytochrome caa3 oxidase
unknown; similar to unknown proteins
RNA polymerase sporulation mother
cell-specific (early) sigma factor
RNA polymerase sporulation
forespore-specific (late) sigma factor
phosphoadenosine phosphosulfate
fliG
fliH
ylxF
-1.8
-1.8
-1.7
1.1
1.0
1.0
-2.1
-1.2
-1.9
-2.0
-1.2
-1.2
-2.4
-2.3
-2.1
1.1
1.1
1.0
1.0
1.1
1.2
-1.4
-1.3
-1.2
-1.2
1.0
1.1
1.2
1.0
-1.2
1.1
-1.6
-1.5
-2.2
-1.3
1.2
1.1
-1.2
1.2
1.3
1.4
1.3
1.3
1.1
1.7
1.6
1.1
1.1
1.0
1.1
-1.2
1.5
1.0
1.1
-1.6
1.6
1.0
1.2
-1.2
1.1
1.1
-2.6
1.1
-1.3
-1.6
1.1
1.0
-1.7
-1.1
yncF
6.5
1.2
1.3
4.6
3.2
1.9
1.6
-1.4
-1.9
-1.8
23.6
1.3
1.2
1.0
1.5
1.0
1.1
1.2
-1.1
-1.1
2.2
-1.8
-1.8
-1.3
1.5
2.5
-1.1
-1.4
-2.0
-1.3
22.4
-1.1
1.8
-1.1
-1.2
1.1
citB
-1.1
1.9
1.2
1.0
-1.5
-1.3
1.2
1.1
1.0
1.0
1.0
-1.7
1.2
-1.5
1.1
1.1
yojH
-1.7
1.7
1.0
1.6
cgeE
-2.3
1.0
-1.2
-1.7
cgeD
yosN
-2.2
4.2
10.3
1.0
3.2
2.3
yosM
3.1
yorE
1.1
1.1
-1.2
1.0
1.0
1.0
ymdA
pksC
-1.7
-1.8
-2.4
-1.1
1.2
-1.4
1.1
pksJ
pksJ
2.0
1.1
1.5
1.8
spo VK
1.6
ynaE
yncD
ymfC
ymfK
dacC
dacC
yoeB
-1.7
1.1
1.1
-1.2
-1.2
-1.7
-1.2
-1.2
1.4
1.0
1.0
1.1
flagellar motor switch protein
1.1
1.1
unknow
1.1
-1.5
-1.5
-1.5
-1.4
unknown; similar to unknown proteins
involved inpolyketide synthesis
1.0
unknown; similar to unknown proteins
-2.7
-1.2
-1.3
-1.3
-1.3
unknown; similar to alanine racemase
flagellar assembly protein
unknown; similar to unknown proteins
unknown
polyketide synthase
polyketide synthase
disruption leads to the
production of immature spores
unknown
aconitate hydratase
penicillin-binding protein
penicillin-binding protein
-1.3
1.2
-1.2
1.2
unknown
-1.5
1.3
1.0
1.0
16.9
16.9
1.3
1.3
glutamate synthase (small subunit)
glutamate synthase (small subunit)
1.0
1.0
1.0
1.1
1.2
1.0
-1.2
1.1
-1.1
6.3
-1.4
-1.2
9.2
1.3
unknown
1.4
1.1
-2.7
3.8
-2.4
1.1
-1.2
unknown
-1.1
1.0
-1.6
-1.1
-1.0
1.4
1.1
1.7
1.2
1.0
1.1
1.2
1.1
1.1
1.1
1.0
1.4
-1.2
1.0
1.1
-2.2
1.1
1.0
-1.2
1.5
1.6
1.0
4.0
2.5
-1.4
4.4
6.4
1.0
2.0
1.3
1.0
-1.1
1.3
1.5
1.1
1.6
1.7
1.2
ligB
19.4
6.9
1.2
-1.1
yonJ
51.1
1.6
kdgK
kdul
yprB
ypfD
ypfB
-1.9
-1.9
sigX
gltB
gltB
yoaH
yobH
yobJ
yobN
yozB
yozC
yosP
-2.5
8.8
-2.0
-1.5
-1.5
1.1
1.2
-1.3
1.1
1.0
-1.2
unknown
unknown
unknown; similar to unknown
proteins
1.0
1.0
unknown; similar to unknown proteins
-1.2
1.1
1.2
1.1
3.7
5.7
1.1
1.1
-1.2
maturation of the spore
maturation of the spore
1.0
2.1
1.0
unknown; similar to unknown proteins
3.2
unknown
1.1
DNA ligase (ATP-dependent)
unknown
unknown
unknown
1.1
1.2
1.2
-1.1
30.9
9.6
1.2
49.4
5.4
1.0
-1.6
63.2
1.2
1.4
39.0
-1.6
-1.5
-1.5
- 1.5
-1.6
-1.4
-1.3
-1.7
-2.6
1.1
1.1
-1.3
-1.2
1.0
-1.5
1.0
1.0
2-keto-3-deoxygluconate kinase
5-keto-4-deoxyuronate isomerase
1.7
-2.2
1.5
1.2
-1.2
1.1
1.8
-1.5
1.8
-1.5
2.0
1.0
-1.1
1.4
-1.2
1.1
2.8
-1.3
3.1
1.0
1.0
unknown
-1.9
2.5
-1.1
1.1
-1.3
-1.4
1.3
1.5
-1.2
1.1
-1.2
1.0
-1.7
ECF-type sigma factor
penicillin-binding protein
2.0
1.4
1.0
1.2
1.1
1.3
1.1
1.1
2.6
1.9
1.0
1.5
-1.4
yqzF
yqzF
folD
-1.1
-1.7
-2.1
1.1
-1.3
-1.6
1.1
1.1
-1.5
1.0
-1.4
-1.7
-1.3
-1.7
-2.3
nusB
yqhQ
-1.7
-1.5
-1.6
1.2
-1.6
1.0
-1.4
1.0
tasA
yqxM
-3.3
-2.2
-2.4
-1.8
-2.3
1.2
-2.1
-1.2
dacF
spollM
yqkK
-1.2
-1.2
1.0
1.0
1.0
1.1
1.0
-1.1
-1.1
-1.2
1.1
1.0
-1.3
1.0
-1.2
1.0
-1.2
-1.2
1.1
-1.5
-1.5
-1.2
-1.9
-1.7
-1.1
1.0
1.1
-1.1
-4.7
-2.8
-1.5
-1.9
-1.1
1.1
123
1.0
1.1
1.1
unknown
unknown
unknown
required for dissolution of the septal cell wall
unknown
1.1
1.1
unknown; similar to unknown proteins
unknown; similar tounknown proteins
methylenetetetrahydrofolate dehydrogenase /
-1.1
-1.2
1.1
1.0
probable transcription termination
unknown; similar to unknown proteins
-2.5
-3.0
1.0
-1.1
methenyltetrahydrofolate cyclohydrolase
translocation-dependent antimicrobial
spore component
unknown
yqgY
yqgA
-2.7
-2.6
-1.9
-2.6
-2.2
-1.6
-1.4
-3.6
-3.5
-1.6
-1.1
-2.1
-1.6
-2.4
-1.6
-1.1
unknown; similar to unknown proteins
-1.4
1.0
unknown
gpr
-2.0
-1.6
-1.4
-1.3
1.1
1.1
1.1
-1.5
1.0
3.4
-1.1
3.2
1.7
-2.3
1.0
1.0
spore protease (degradation of SASPs)
N-acetylmuramoyl-L-alanine amidase
-1.1
1.1
(sporulation mother cell wall)
1.9
unknown; similar to ribonuclease inhibitor
1.0
1.3
-1.2
1.2
unknown
cwlH
9.4
1.3
1.2
-1.1
5.7
1.1
1.1
yrkK
1.0
1.0
1.1
-1.2
1.0
1.0
1.3
1.2
1.0
5.1
1.0
1.0
yrdD
yrpD
yral
yrrL
-1.5
3.8
11.2
-2.2
1.5
-1.7
1.3
-1.5
1.2
-1.4
1.2
-1.4
1.4
-1.2
1.1
-1.4
1.1
1.1
8.7
-3.3
-1.1
1.1
-1.3
1.1
aspS
hisS
ysoA
uvrC
mutSB
-2.4
-2.4
-1.8
1.5
- 1.4
-1.8
-1.8
-1.1
1.3
1.0
-1.6
-1.6
-1.1
1.6
-1 .2
-1.6
-1.4
-1.2
1.1
1.0
-1.3
-1.7
-1.6
-1.4
1.3
1.1
-1.2
-1.2
-1.2
1.1
1.2
1.2
1.3
-2.4
1.1
1.3
-1.2
-1.2
1.4
1.1
1.2
-1.5
-1.6
-1.1
yshC
-1.5
1.0
1.1
1.1
1.1
1.0
1.1
1.1
yrdF
1.7
1.1
1.2
1.3
2.8
-1.3
-1.5
-1.2
-1.9
-1.1
1.1
ytmN
2.1
1.2
1.4
1.0
3.4
1.0
1.2
yteJ
-1.5
1.1
-1.2
1.2
-1.8
1.2
1.2
sspA
3.2
1.1
1.3
-1.2
2.0
1.1
-1.1
1.0
1.1
1.1
ytoQ
-1.8
-1.5
-1.5
-1.6
-1.2
ytlR
ytlQ
-1.6
-1.6
1.1
1.0
-1.4
-1.3
1.0
-1.5
-1.5
bioA
-1.7
-1.7
-1.5
mcpA
mcpA
-2.3
-2.1
- 1.3
-1.5
-1.4
-1.1
yusT
yvrH
yvrN
yvrN
yvbF
2.3
-1.7
-1.6
-2.0
-1.1
-1.2
-1.3
-1.4
yvbX
-1.8
-1.9
-1.3
-1.2
1.3
-1.2
-1.4
-1.6
1.1
yvfO
1.6
yveS
yveA
1.5
1.2
1.0
1.0
1.0
1.5
1.1
1.1
1.1
unknown
unknown
unknown
unknown; similar to folate metabolism
aspartyl-tRNA synthetase
histidyl-tRNA synthetase
unknown; similar to unknown
proteins
excinuclease ABC (subunit C)
probable DNA mismatch repair protein
unknown; similar to DNA polymerase beta
unknown; similar to amino-acid
ABC transporter
unknown; similar to unknown proteins
small acid-soluble spore protein
(major alpha-type SASP)
unknown; similar to unknown proteins
unknown; similar to unknown proteins
-1.2
1.0
-1.1
1.1
-1.4
1.0
1.0
unknown; similar to unknown proteins
-1.3
-2.2
-2.2
1.0
1.1
-1.2
-1.2
-1.5
-1.5
methyl-accepting chemotaxis protein
methyl-accepting chemotaxis protein
-1.2
-1.3
-1.3
1.3
-1.3
-2.4
-2.8
1.1
unknown
-1.6
-1.2
-1.6
1.1
-1.1
-1.7
-1.7
1.0
-1.2
-1.5
-1.1
-1.3
1.0
-1.1
1.0
1.2
1.1
1.1
1.1
1.1
1.2
1.2
1.0
1.0
1.0
1.2
1.1
1.0
1.2
1.0
1.3
-1.8
-1.8
-1.3
-1.2
-1.7
-1.2
-2.8
-1.4
1.0
-1.3
-1.2
1.1
-1.4
-1.4
1.0
-1.2
1.1
1.0
1.0
-1.2
1.0
ywoF
-1.7
2.3
-1.7
-1.7
-1.1
ywjF
yweA
1.6
-3.6
1.1
-2.4
1.2
-2.5
spsE
1.8
1.0
yxjA
-1.8
yxjA
yxal
yydB
1.0
-1.7
-1.5
-1.2
-1.3
1.4
-2.4
1.0
1.1
-2.8
1.2
-4.8
1.0
1.2
1.1
1.1
1.4
-1.2
-1.5
-1.4
-2.1
-2.1
-1.5
-1.4
1.0
1.1
-1.6
-1.5
1.0
purA
-4.4
-2.7
yosO
27.2
1.6
yvdS
-1.6
-1.6
-1.2
-1.4
unknown
-1.3
unknown; similar to ABC transporter
- 1.3
1.1
unknown; similar to ABC transporter
1.0
unknown; similar to unknown proteins
1.1
unknown
- 1.2
-1.2
1.2
1.1
1.1
-2.5
1.0
1.1
-3.4
1.1
1.2
1.1
-1.1
1.0
-1.6
1.1
1.0
1.0
1.0
-1.4
-1.2
-1.1
-2.1
-1.4
-2.4
-1.2
1.0
-1.5
1.0
1.1
-1.7
1.0
1.1
-2.1
1.0
-1.8
1.0
-2.4
-1.9
-8.4
-1.8
-2.9
-2.5
1.1
1.0
-1.1
29.9
-1.1
-1.0
17.8
1.1
a Genes
aminotransferase
unknown
unknown; similar to unknown
unknown
unknown
unknown
reductase
unknown
spore coat polysaccharide synthesis
unknown; similar to pyrimidine
nucleoside transport
unknown; similar to pyrimidine
nucleoside transport
unknown
unknown
adenylosuccinate synthetase
affected significantly by DNA damage only in the presence of phages are listed.
Genes are ordered by chromosomal location.
Results are extracted from the data presented in Figure 1, and the column numbers represent the respective
124
proteins
unknown; similar to permease
columns in figure 1.
Numbers are the average of three replicates of microarray analysis. Negative numbers represent fold decrease
in expression.
125
genes fall in various categories indicating that the induction of phages impinges on many cellular
processes: 49 genes in cell envelope functions, 35 genes in intermediate metabolism, 24 genes in
information pathways, 10 genes in antibiotic production, stress and detoxification, and 66 genes
of unknown function. The observation that a large number of genes involved in cell-envelope
functions are affected by phage induction is not entirely surprising as phages need to modify the
cell envelope so that the phage particles will be released into the environment. Understanding
what events in the cell envelope bring about the observed changes in gene expression and
whether there is a different mechanism for the phages to affect gene expression will require
further studies.
Replication arrest and unrepaired UV damage affect the expression of genes regulated
by DnaA. DnaA, the replication initiation protein in bacteria, mediates a transcriptional response
to perturbations in replication that is independent of RecA (23). We had previously established
that replication arrest caused by HPUra affects the expression of DnaA-regulated genes including
dnaA, dnaN, sda, ywlC, andftsL (23). We were interested in investigating whether this DnaAmediated response was induced under any of the other conditions studied. MMC and UV
treatments affected a small subset of the genes regulated by DnaA (Table 2 and Table 3). In the
wild type strain, MMC affected 8 operons (12 genes) of the 20 operons proposed to be regulated
directly by DnaA (23). HPUra affects the expression of all 20 operons in a recA-independent
manner (23), yet MMC affected only 3 operons (total of 12 genes) in a recA-null background.
.MMC affected 2 putative DnaA operons (4 genes) in the phage-defective strain and only 1
operon (4 genes) in the phage-defective recA strain (Table 3). These results indicate that DnaAregulated genes are affected differently by MMC compared to replication arrest. Because MMC
treatment did not change the expression of dnaAN and sda, the best characterized DnaA targets,
126
it is possible that the genes affected by MMC are controlled by factors other than DnaA under
these conditions. In addition, it is as likely that after MMC treatment, DnaA may affect the
expression of only a subset of genes, due to different sensitivity of the promoters to DnaA
activity.
UV treatment affected only 1 operon in the phage-defective strain (Table 3). UV treatment in
the phage-defective recA strain affected the well characterized DnaA targets dnaAN and sda as
well as 4 more proposed DnaA targets (total of 8 genes in 6 operons). Only one of the operons
regulated by DnaA, ynzC-ywfO-ywgA, was affected by treatment with UV, MMC, and HPUra.
The direction of the effects (either increased or decreased) on all operons except yclNOP was the
same under all conditions where the operons were significantly affected.
The effects on more DnaA-regulated genes after UV damage in the recA null vs. recA+ strain
are likely due to the inability of recA mutants to repair the DNA damage, thus causing prolonged
replication arrest (data not shown) (11, 12, 29). The magnitude of the effect on DnaA regulated
genes after UV damage was not as large as in the HPUra experiments. One possibility is that UV
partially blocks replication in all cells. For example, the replisome may stall for some period of
time and then replication may restart until the replisome stalls again. Another possibility is that
replication is completely blocked in a subpopulation of cells and only these cells exhibit effects
on DnaA regulated genes, thus in the context of the whole population the effects on DnaA
regulated genes appears less than when cells are treated with HPUra. These possibilities could be
distinguished by analysis of gene expression in single cells.
In HPUra treated cells, the changes in expression of sda and yllB are less than that of other
affected genes. However, UV irradiation affected these genes and not others that are more
strongly affected by HPUra. One possible explanation for seeing only a subset of the proposed
127
DnaA-regulated genes after UV treatment is that expression of the proposed DnaA-regulated
genes may require factors in addition to DnaA, and that these additional factors are differentially
regulated by UV and HPUra treatments.
Our results indicate that the recA-independent, DnaA-dependent responses to the various
DNA damaging treatments and replication arrest are qualitatively and quantitatively different.
These differences are likely due to mechanistic differences of how MMC, UV and HPUra affect
replication. We speculate that since different events occur at the replication fork when replication
is arrested with various treatments, replication is affected differently, which leads to different
effects on DnaA and the DnaA-dependent transcriptional response. Furthermore, in addition to
causing DNA damage, MMC has the potential to damage proteins and, thus, could disrupt
protein function and disrupt other pathways that occur inside the cell (17). These effects on
protein function may influence the transcriptional response after DNA damage.
MMC causes a relative increase in origin proximal chromosomal loci. The expression of
a large number of genes within the origin proximal region appeared to be significantly increased
after treatment with MMC in recA+ and recA null strains when grown in rich as well as in
minimal medium (Fig 2A, Table 2, and data not shown). An analysis of the mRNA abundance,
irrespective of statistical significance, revealed that in cells treated with MMC >80% of the
genes in the origin proximal region have higher levels of gene transcripts in most experiments
(Fig. 2A). In only one experiment did we observe an effect on less than 80% of the origin
proximal genes, with only 56% of the origin proximal genes having higher mRNA abundance.
These effects on gene expression were usually less than 2-fold.
This apparent increase in origin proximal gene expression might be caused by an increase in
the rate of transcription of these genes. Alternatively, the effect on mRNA abundance of the
128
2-
MMC
MMC
A.
A.
10-
:1..
Ulm, 1i+
....
.....
-1-
2
....
.
-2·
-3
.... ...
....
....
...
..
...
....
...
....
.......
........ ...... ..
. ..
.
------------
....
.......
.
....
...
....
....
-- -.-........
.....
....
.....
....
..
.............
..
.....1.................
..........
.............
23
2
C.
...
.
....
HPUra
.. .
......... . ............
.......
.
. .......
. .
.....
...
.. .... .
....
. .
.
.
....
.
.
...
0
0-i
.P. -T w-
-1
|
-2
. . ...
.........
_....................
.........
........
I.......
..............................
.....
...
..
.....
...--..............
...
.......
..
...
...
.....
.....
...
...
.
·.
,.:.
r:"• "
. .~
.-. .. .....
• It •.:
I. - .•
................
---.....
...
..
...........
...
.........
..................
mRNA
tF. .JV.........
"1
2-
E.
UV
.
.. ...
..
S
-2
mRNA
|·~
......
... .. ... .......
•
.......
. ..........
d- recA"
+
recA
pd-
Dd- recA'
...
......
.......
..
...
...
. ..
...
.....
..
....
.....
........
......
.....
...
......
.........
-3
... _.....l~..........
.....................
....
...........
............
..................
. ....
.. ........
,
Figure 2
129
Figure 2. MMC treatment causes a relative increase in origin proximal gene expression and
gene dosage. The relative amount of mRNA (panels A, C, D, E, F) or chromosomal DNA (panel
B) of each gene in cells treated with MMC (A, B), HPUra (C, D), or UV (E, F) compared to
untreated was determined and the average change 60 min after treatment from three replicates is
plotted on the y-axis in log2 format. The position of each gene on the chromosome, with the
origin region in the middle, is plotted on the x-axis. The relevant genotype of each strain is
indicated in each panel. Pd- refers to phage-defective strains.
130
oriC-proximal genes might be caused by increased copies of the origin region. Such an effect on
gene dosage could be achieved if MMC were causing over-initiation of DNA replication, with
the newly assembled replication forks proceeding for a relatively short distance (-50 kb).
Alternatively, MMC could be slowing DNA replication elongation without affecting the rate of
initiation, thus effectively increasing replication in the origin region.
To test the hypothesis that there is a higher copy number of the genes near oriC in MMC
treated cells, we performed genomic microarray experiments. This type of experiment allows a
comparison to be made between the genomic compositions of different bacterial cultures. The
genomic microarray experiments revealed that cultures treated with MMC have an increased
(-2-fold) relative amount of origin proximal DNA compared to untreated cells (Fig. 2B). The
fold increase of a particular chromosomal locus was inversely proportional to the distance from
the origin, with virtually no increase in DNA content after 60 kb away from the origin.
Thymidine incorporation experiments demonstrated that the rate of replication decreases after
MMC treatment, but not to the level of that in HPUra treated cultures (Fig. 3), indicating that
MMC inhibits, although does not abolish, DNA replication. This intermediate effect of MMC is
consistent with the notion that MMC slows down elongation, causing a relative increase in origin
proximal DNA and a relative increase in transcripts from origin proximal genes. Under these
conditions MMC has a more toxic effect than HPUra 60 min after addition of the drug (data not
shown), indicating that the more severe effects on DNA replication by HPUra are not due to
higher toxicity.
Treatment of recA+ cells with UV irradiation also caused an increase in mRNA levels of the
origin-proximal genes (Fig. 2F) indicating increased gene dosage in this region. However, in a
recA null mutant, this increase was not observed (Fig. 2E). In contrast to these effects and those
131
18000
16000
5
14000
12000
10000
o
8000
6000
4000
2000
0
10
20
30
40
50
60
70
Time after treatment (min)
Figure 3
132
Figure 3. Effects of MMC and HPUra on DNA replication. A culture of wild type cells
(JH642) was grown to mid-exponential phase and spilt in three: one part was left untreated, one
was treated with MMC, and one was treated with HPUra. Aliquots were taken at indicated times
and the rate of DNA synthesis was determined by measuring the rate of incorporation of 3Hthymidine into DNA, with background subtracted (Materials and Methods). Data presented are
the average of two experiments with error bars indicating one standard deviation. mock
treatment, triangles; MMC, circles; HPUra, squares.
133
of MMC, treatment of either recA+ or recA null mutant cells with HPUra (to block replication)
did not cause an increase in mRNA levels of these genes (Fig. 2C, D). These results are
consistent with the notion that different mechanisms for blocking DNA replication have different
regulatory effects. We suggest that HPUra in recA+ and recA mutant cells and UV damage in
recA mutant cells causes a strong arrest of replication forks throughout the chromosome such
that there is little or no relative change in chromosomal content as compared to untreated cells.
We suggest that UV light has different effects in recA+ and recA cells because the recA mutant
cells are unable to repair the damage, whereas wild type cells repair the damage and resume
replication within 40 min. We propose that MMC causes a relative increase in chromosomal
content of origin proximal regions by slowing down replication fork progression without causing
a decrease in the rate of initiation of replication.
Our results demonstrate that the inhibition of replication elongation by different DNA
damaging agents or inactivation of replication components can have both unique and overlapping
effects on gene expression. The results also indicate that B. subtilis, and likely other bacteria, can
induce several different response pathways after DNA damage, but the extent of induction of
each pathway may vary such that the organisms have the best chance of survival.
Acknowledgements
This work was supported in part by NIH grants GM41934 to ADG. We thank Neal Brown
for the generous gift of HPUra, and Philippe Noirot and S.D. Ehrlich for providing us with
JJS39, lexA, and lexA(ind-) strains. We thank C. Lee and J. Auchtung for technical help, and T.
Baker, C. Lee, M. Berkmen, J. Auchtung, and L. Simmons for suggestions and comments on the
manuscript.
134
References
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
Au, N., E. Kuester-Schoeck, V. Mandava, L. E. Bothwell, S. P. Canny, K. Chachu, S.
A. Colavito, S. N. Fuller, E. S. Groban, L. A. Hensley, T. C. O'Brien, A. Shah, J. T.
Tierney, L. L. Tomm, T. M. O'Gara, A. I. Goranov, A. D. Grossman, and C. M.
Lovett. 2005. Genetic composition of the Bacillus subtilis SOS system. J Bacteriol
187:7655-66.
Auchtung, J. M., C. A. Lee, R. E. Monson, A. P. Lehman, and A. D. Grossman.
2005. Regulation of a Bacillus subtilis mobile genetic element by intercellular signaling
and the global DNA damage response. Proc Natl Acad Sci U S A 102:12554-9.
Beaber, J. W., B. Hochhut, and M. K. Waldor. 2004. SOS response promotes
horizontal dissemination of antibiotic resistance genes. Nature 427:72-4.
Berka, R. M., J. Hahn, M. Albano, I. Draskovic, M. Persuh, X. Cui, A. Sloma, W.
Widner, and D. Dubnau. 2002. Microarray analysis of the Bacillus subtilis K-state:
genome-wide expression changes dependent on ComK. Mol Microbiol 43:1331-45.
Britton, R. A., P. Eichenberger, J. E. Gonzalez-Pastor, P. Fawcett, R. Monson, R.
Losick, and A. D. Grossman. 2002. Genome-wide analysis of the stationary-phase
sigma factor (sigma-H) regulon of Bacillus subtilis. J Bacteriol 184:4881-90.
Brown, N. C. 1970. 6-(p-hydroxyphenylazo)-uracil: a selective inhibitor of host DNA
replication in phage-infected Bacillus subtilis. Proc Natl Acad Sci U S A 67:1454-61.
Burkholder, W. F., I. Kurtser, and A. D. Grossman. 2001. Replication initiation
proteins regulate a developmental checkpoint in Bacillus subtilis. Cell 104:269-79.
Byun, T. S., M. Pacek, M. C. Yee, J. C. Walter, and K. A. Cimprich. 2005. Functional
uncoupling of MCM helicase and DNA polymerase activities activates the ATRdependent checkpoint. Genes Dev 19:1040-52.
Cheo, D. L., K. W. Bayles, and R. E. Yasbin. 1991. Cloning and characterization of
DNA damage-inducible promoter regions from Bacillus subtilis. J Bacteriol 173:1696703.
Cheo, D. L., K. W. Bayles, and R. E. Yasbin. 1992. Molecular characterization of
regulatory elements controlling expression of the Bacillus subtilis recA+ gene. Biochimie
74:755-62.
Courcelle, J., C. Carswell-Crumpton, and P. C. Hanawalt. 1997. recF and recR are
required for the resumption of replication at DNA replication forks in Escherichia coli.
Proc Natl Acad Sci U S A 94:3714-9.
Courcelle, J., D. J. Crowley, and P. C. Hanawalt. 1999. Recovery of DNA replication
in UV-irradiated Escherichia coli requires both excision repair and recF protein function.
J Bacteriol 181:916-22.
Courcelle, J., and P. C. Hanawalt. 2003. RecA-dependent recovery of arrested DNA
replication forks. Annu Rev Genet 37:611-46.
Courcelle, J., A. Khodursky, B. Peter, P. O. Brown, and P. C. Hanawalt. 2001.
Comparative gene expression profiles following UV exposure in wild-type and SOSdeficient Escherichia coli. Genetics 158:41-64.
de Vos, W. M., S. C. de Vries, and G. Venema. 1983. Cloning and expression of the
Escherichia coli recA gene in Bacillus subtilis. Gene 25:301-8.
135
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
Dervyn, E., C. Suski, R. Daniel, C. Bruand, J. Chapuis, J. Errington, L. Janniere,
and S. D. Ehrlich. 2001. Two essential DNA polymerases at the bacterial replication
fork. Science 294:1716-9.
Dronkert, M. L., and R. Kanaar. 2001. Repair of DNA interstrand cross-links. Mutat
Res 486:217-47.
Duigou, S., S. D. Ehrlich, P. Noirot, and M. F. Noirot-Gros. 2004. Distinctive genetic
features exhibited by the Y-family DNA polymerases in Bacillus subtilis. Mol Microbiol
54:439-51.
Duigou, S., S. D. Ehrlich, P. Noirot, and M. F. Noirot-Gros. 2005. DNA polymerase I
acts in translesion synthesis mediated by the Y-polymerases in Bacillus subtilis. Mol
Microbiol 57:678-90.
Elledge, S. J., Z. Zhou, J. B. Allen, and T. A. Navas. 1993. DNA damage and cell cycle
regulation of ribonucleotide reductase. Bioessays 15:333-9.
Fabret, C., S. D. Ehrlich, and P. Noirot. 2002. A new mutation delivery system for
genome-scale approaches in Bacillus subtilis. Mol Microbiol 46:25-36.
Friedberg, E. C., Graham C. Walker, and Wolfram Siede. 1995. DNA Repair and
Mutagenesis. ASM Press.
Goranov, A. I., L. Katz, A. M. Breier, C. B. Burge, and A. D. Grossman. 2005. A
transcriptional response to replication status mediated by the conserved bacterial
replication protein DnaA. Proc Natl Acad Sci U S A 102:12932-7.
Groban, E. S., M. B. Johnson, P. Banky, P. G. Burnett, G. L. Calderon, E. C.
Dwyer, S. N. Fuller, B. Gebre, L. M. King, I. N. Sheren, L. D. Von Mutius, T. M.
O'Gara, and C. M. Lovett. 2005. Binding of the Bacillus subtilis LexA protein to the
SOS operator. Nucleic Acids Res 33:6287-95.
Harwood, C. R., and S. M. Cutting. 1990. Molecular Biological Methods for Bacillus.
John Wiley & Sons, Chichester, England.
Jaacks, K. J., J. Healy, R. Losick, and A. D. Grossman. 1989. Identification and
characterization of genes controlled by the sporulation-regulatory gene spo0H in Bacillus
subtilis. J Bacteriol 171:4121-9.
Kawai, Y., S. Moriya, and N. Ogasawara. 2003. Identification of a protein, YneA,
responsible for cell division suppression during the SOS response in Bacillus subtilis.
Mol Microbiol 47:1113-22.
Khil, P. P., and R. D. Camerini-Otero. 2002. Over 1000 genes are involved in the DNA
damage response of Escherichia coli. Mol Microbiol 44:89-105.
Koehler, D. R., J. Courcelle, and P. C. Hanawalt. 1996. Kinetics of pyrimidine(64)pyrimidone photoproduct repair in Escherichia coli. J Bacteriol 178:1347-50.
Krogh, S., M. O'Reilly, N. Nolan, and K. M. Devine. 1996. The phage-like element
PBSX and part of the skin element, which are resident at different locations on the
Bacillus subtilis chromosome, are highly homologous. Microbiology 142 (Pt 8):203140.
Kunkel, B., R. Losick, and P. Stragier. 1990. The Bacillus subtilis gene for the
development transcription factor sigma K is generated by excision of a dispensable DNA
element containing a sporulation recombinase gene. Genes Dev 4:525-35.
Kuzminov, A. 1999. Recombinational repair of DNA damage in Escherichia coli and
bacteriophage lambda. Microbiol Mol Biol Rev 63:751-813, table of contents.
136
33.
34.
35.
36.
37.
38.
39.
40.
41.
42.
43.
44.
45.
46.
47.
48.
49.
Lamont, I., A. M. Brumby, and J. B. Egan. 1989. UV induction of coliphage 186:
prophage induction as an SOS function. Proc Natl Acad Sci U S A 86:5492-6.
Lamont, I., H. Richardson, D. R. Carter, and J. B. Egan. 1993. Genes for the
establishment and maintenance of lysogeny by the temperate coliphage 186. J Bacteriol
175:5286-8.
Lazarevic, V., A. Dusterhoft, B. Soldo, H. Hilbert, C. Mauel, and D. Karamata.
1999. Nucleotide sequence of the Bacillus subtilis temperate bacteriophage SPbetac2.
Microbiology 145 ( Pt 5):1055-67.
Le Chatelier, E., O. J. Becherel, E. d'Alencon, D. Canceill, S. D. Ehrlich, R. P.
Fuchs, and L. Janniere. 2004. Involvement of DnaE, the second replicative DNA
polymerase from Bacillus subtilis, in DNA mutagenesis. J Biol Chem 279:1757-67.
Lemon, K. P., I. Kurtser, and A. D. Grossman. 2001. Effects of replication termination
mutants on chromosome partitioning in Bacillus subtilis. Proc Natl Acad Sci U S A
98:212-7.
Little, J. W. 1984. Autodigestion of lexA and phage lambda repressors. Proc Natl Acad
Sci U S A 81:1375-9.
Love, P. E., M. J. Lyle, and R. E. Yasbin. 1985. DNA-damage-inducible (din) loci are
transcriptionally activated in competent Bacillus subtilis. Proc Natl Acad Sci U S A
82:6201-5.
Lovett, C. M., Jr., P. E. Love, and R. E. Yasbin. 1989. Competence-specific induction
of the Bacillus subtilis RecA protein analog: evidence for dual regulation of a
recombination protein. J Bacteriol 171:2318-22.
Lovett, C. M., Jr., T. M. O'Gara, and J. N. Woodruff. 1994. Analysis of the SOS
inducing signal in Bacillus subtilis using Escherichia coli LexA as a probe. J Bacteriol
176:4914-23.
Lovett, C. M., Jr., and J. W. Roberts. 1985. Purification of a RecA protein analogue
from Bacillus subtilis. J Biol Chem 260:3305-13.
Lusetti, S. L., and M. M. Cox. 2002. The bacterial RecA protein and the
recombinational DNA repair of stalled replication forks. Annu Rev Biochem 71:71-100.
McVeigh, R. R., and R. E. Yasbin. 1996. Phenotypic differentiation of "smart" versus
"naive" bacteriophages of Bacillus subtilis. J Bacteriol 178:3399-401.
Miller, M. C., J. B. Resnick, B. T. Smith, and C. M. Lovett, Jr. 1996. The bacillus
subtilis dinR gene codes for the analogue of Escherichia coli LexA. Purification and
characterization of the DinR protein. J Biol Chem 271:33502-8.
Molle, V., M. Fujita, S. T. Jensen, P. Eichenberger, J. E. Gonzalez-Pastor, J. S. Liu,
and R. Losick. 2003. The SpoOA regulon of Bacillus subtilis. Mol Microbiol 50:1683701.
Napolitano, R., R. Janel-Bintz, J. Wagner, and R. P. Fuchs. 2000. All three SOSinducible DNA polymerases (Pol II, Pol IV and Pol V) are involved in induced
mutagenesis. Embo J 19:6259-65.
Neher, S. B., J. M. Flynn, R. T. Sauer, and T. A. Baker. 2003. Latent ClpXrecognition signals ensure LexA destruction after DNA damage. Genes Dev 17:1084-9.
Ogura, M., H. Yamaguchi, K. Kobayashi, N. Ogasawara, Y. Fujita, and T. Tanaka.
2002. Whole-genome analysis of genes regulated by the Bacillus subtilis competence
transcription factor ComK. J Bacteriol 184:2344-51.
137
50.
51.
52.
53.
54.
55.
56.
57.
58.
59.
60.
61.
62.
63.
64.
65.
66.
Okamoto, K., J. A. Mudd, and J. Marmur. 1968. Conversion of Bacillus subtilis DNA
to phage DNA following mitomycin C induction. J Mol Biol 34:429-37.
Perego, M., G. B. Spiegelman, and J. A. Hoch. 1988. Structure of the gene for the
transition state regulator, abrB: regulator synthesis is controlled by the spo0A sporulation
gene in Bacillus subtilis. Mol Microbiol 2:689-99.
Quillardet, P., M. A. Rouffaud, and P. Bouige. 2003. DNA array analysis of gene
expression in response to UV irradiation in Escherichia coli. Res Microbiol 154:559-72.
Quinones, M., H. H. Kimsey, and M. K. Waldor. 2005. LexA cleavage is required for
CTX prophage induction. Mol Cell 17:291-300.
Rand, L., J. Hinds, B. Springer, P. Sander, R. S. Buxton, and E. O. Davis. 2003. The
majority of inducible DNA repair genes in Mycobacterium tuberculosis are induced
independently of RecA. Mol Microbiol 50:1031-42.
Raymond-Denise, A., and N. Guillen. 1992. Expression of the Bacillus subtilis dinR
and recA genes after DNA damage and during competence. J Bacteriol 174:3171-6.
Raymond-Denise, A., and N. Guillen. 1991. Identification of dinR, a DNA damageinducible regulator gene of Bacillus subtilis. J Bacteriol 173:7084-91.
Roberts, J. W., and R. Devoret. 1983. Lysogenic Induction, p. 123-144. In R. W.
Hendrix, J. W. Roberts, F. W. Stahl, and R. A. Weisberg (ed.), Lambda II. Cold Spring
Harbor Laboratory, Cold Spring Harbor, New York.
Rowland, S. L., W. F. Burkholder, K. A. Cunningham, M. W. Maciejewski, A. D.
Grossman, and G. F. King. 2004. Structure and mechanism of action of Sda, an
inhibitor of the histidine kinases that regulate initiation of sporulation in Bacillus subtilis.
Mol Cell 13:689-701.
Sassanfar, M., and J. W. Roberts. 1990. Nature of the SOS-inducing signal in
Escherichia coli. The involvement of DNA replication. J Mol Biol 212:79-96.
Smith, B. T., and G. C. Walker. 1998. Mutagenesis and more: umuDC and the
Escherichia coli SOS response. Genetics 148:1599-610.
Steensma, H. Y., L. A. Robertson, and J. D. van Elsas. 1978. The occurrence and
taxonomic value of PBS X-like defective phages in the genus Bacillus. Antonie Van
Leeuwenhoek 44:353-66.
Stragier, P., B. Kunkel, L. Kroos, and R. Losick. 1989. Chromosomal rearrangement
generating a composite gene for a developmental transcription factor. Science 243:50712.
Sung, H. M., G. Yeamans, C. A. Ross, and R. E. Yasbin. 2003. Roles of YqjH and
YqjW, homologs of the Escherichia coli UmuC/DinB or Y superfamily of DNA
polymerases, in stationary-phase mutagenesis and UV-induced mutagenesis of Bacillus
subtilis. J Bacteriol 185:2153-60.
Sutton, M. D., I. Narumi, and G. C. Walker. 2002. Posttranslational modification of
the umuD-encoded subunit of Escherichia coli DNA polymerase V regulates its
interactions with the beta processivity clamp. Proc Natl Acad Sci U S A 99:5307-12.
Tusher, V. G., R. Tibshirani, and G. Chu. 2001. Significance analysis of microarrays
applied to the ionizing radiation response. Proc Natl Acad Sci U S A 98:5116-21.
Warner, F. D., G. A. Kitos, M. P. Romano, and H. E. Hemphill. 1977.
Characterization of SP3: a temperate bacteriophage from Bacillus subtilis 168M.
Canadian Jpurnal of Microbiology 23:45-51.
138
67.
68.
69.
70.
71.
Winterling, K. W., D. Chafin, J. J. Hayes, J. Sun, A. S. Levine, R. E. Yasbin, and R.
Woodgate. 1998. The Bacillus subtilis DinR binding site: redefinition of the consensus
sequence. J Bacteriol 180:2201-11.
Winterling, K. W., A. S. Levine, R. E. Yasbin, and R. Woodgate. 1997.
Characterization of DinR, the Bacillus subtilis SOS repressor. J Bacteriol 179:1698-703.
Witkin, E. M. 1976. Ultraviolet mutagenesis and inducible DNA repair in Escherichia
coli. Bacteriol Rev 40:869-907.
Yasbin, R. E., P. I. Fields, and B. J. Andersen. 1980. Properties of Bacillus subtilis 168
derivatives freed of their natural prophages. Gene 12:155-9.
Zhou, B. B., and S. J. Elledge. 2000. The DNA damage response: putting checkpoints in
perspective. Nature 408:433-9.
139
Chapter 4
Early association of replisome components with the origin of replication
provides a mechanism for regulating the initiation of DNA replication
140
Abstract
Multiple layers of regulation control the initiation of DNA replication and ensure that the
growth of an organism is coupled with the duplication of its genome and subsequent
proliferation. Using chromatin immunoprecipitation, we assayed the order in which
components of the replication machinery associate with the origin of replication in the
bacterium Bacillus subtilis. Contrary to current models, we observed that, in vivo,
primase and DNA polymerase holoenzyme associate with the origin before helicase. We
explored the possible regulatory role of this order of association and found that the clamp subunit of DNA polymerase holoenzyme stimulates initiation of DNA replication
by regulating a step of initiation before association of helicase with the origin. This
regulation likely coordinates helicase loading with the presence of DNA polymerase
holoenzyme and serves to increase genomic stability.
141
Initiation of DNA replication is tightly controlled so that the frequency of
chromosome replication stays proportional to the rate of cell growth. It is of great interest
to understand how organisms control the initiation of replication, as the regulation of
replication is central cell-cycle and genomic stability. The initiation of DNA replication
requires multiple events: the origins of replication are recognized and melted, and
components of the DNA replication machinery, including the DNA polymerase, primase,
P-clamp, clamp loader, and helicase associate with the origins. In eukaryotic cells, several
of these events are regulated (1). Replication initiation factors, such as ORC (origin
recognition complex) and Cdc6, and replication elongation factors, such as DNA Pol F
participate in different steps of replication initiation (1). It appears that DNA Pol S
associates with the origins of replication prior to origin unwinding and primase
association and stimulates late events during initiation (2-4).
In contrast, bacterial replication elongation components are only thought to
associate with the origin as a last step, after helicase activation (5-9). This model is based
on in vitro experiments with purified Escherichiacoli replication components, which
show that replication elongation factors, including DNA polymerase, primase, P-clamp
and clamp loader, require helicase to associate with the origin of replication (5, 7, 10).
To better understand the regulation of replication initiation in the bacterium Bacillus
subtilis in vivo, we investigated which replication components were associated with the
origin at different steps during the initiation of replication using (ChIP). We assayed the
association of single stranded DNA binding protein SSB, initiation factors [origin melting
protein and ORC analogue (DnaA), helicase loading protein (DnaB)], and various
142
components of the DNA replication machinery (helicase (DnaC), primase (DnaG), DNA
polymerase (PolC), P-clamp (DnaN), clamp loader/t-subunit (DnaX)) with the origin.
During replication initiation, DnaA, a highly conserved AAA+ protein, binds to the
chromosomal origin of replication (oriC) in a sequence specific manner and melts the
origin to expose ssDNA (8, 9). To assay if DnaA, regulates the association of replication
proteins with the origin, we synchronized replication in cells using a temperature
sensitive allele dnaA Its (11). We found that most replication components tested were not
associated with the origin at the non-permissive temperature, as judged by the relative
enrichment of origin DNA by ChIP (Fig.1A). Only SSB and helicase loader DnaB had
some association with the origin at the non-permissive temperature. This association at
non-permissive temperature could be due to some residual DnaA activity at nonpermissive temperature or to DnaA-independent recruitment of SSB and DnaB to the
origin.
Once cultures were shifted to the permissive temperature, thereby allowing
replication to proceed, most of the replication proteins tested had a 6-14 fold increase in
association with the origin (Fig. IA). These results indicate that DnaA is required for the
association of replisome components (P-clamp, clamp loader, primase, replicative
polymerase) and a helicase loader component (DnaB) and SSB with the origin, agreeing
with the current hypothesis that DnaA is one of the furthest upstream regulators of
initiation. For two proteins, PolC and DnaG, the increase in origin association after
temperature downshift was -2 fold. This lower enrichment is likely due to the low
abundance of PolC per cell (12) and/or some specific effect of dnaA its on initiation.
143
A
Synchronized with dnaA lIs
o arrested
• released
14212 ---
Q, 10
o
Dna A DnaB DnaC DnaO DnaN DnaX
B Synchronized
respective
ssb
with
pole
no ab
no
no
myc
OFP
tag
tag
C Synchronized with
ts allele
dnaD23ls
12
39
210
.no ab
..c
U
'-'.
o dna4lts
Cd)
EIdnaD23Is
]
fJdnaBl341s
u
't:
Dm/a121S
c:
U.l
'0
o
tJ..
Arrested
o min
DllaB [P-ed
2 min
5 min
DnaB IP-ed
Figure 1
144
Figure 1. Use of Chromatin Immunoprecipitation (ChIP) to assay the association of
replication proteins with the origin of replication. Cells were synchronized for
replication using various temperature sensitive alleles. Samples were cross-linked at
different times (0-15 min) after downshift to permissive temperature, and proteins of
interest were immunoprecipitated. Amounts of co-immunoprecipitated DNA were
analyzed by PCR of serial dilutions of immuno-precipitated (IP) and input DNA with
oriC primers and distal region (dnaD) primers. PCR reactions were analyzed on agarose
gels stained with ethidium bromide. Fold enrichment of origin DNA was calculated as
oriC (IP/input) + distal region (IP/input). Similar results were observed in two
independent experiments. A) Association of initiation components and replisome
components with the origin is stimulated by DnaA function. Cells were synchronized for
replication with dnaA ts by incubation at 500 C for 60 min. Association of various
proteins, indicated on the x-axis, with the origin was assayed by ChIP. White bars
represent fold enrichment of origin DNA before release of replication and black bars
represent fold enrichment 10min after release into synchronous replication at 350 C. No
antibody (no ab) control samples were processed identically as the rest except no primary
antibody was used. Control samples were incubated with anti-myc or anti-GFP antibody,
respectively, but no myc- or GFP-tagged proteins, respectively, were present in the
sample. B. subtilis strains carrying DnaC, DnaN, DnaX, and PolC GFP- or myc-tagged
proteins, where the tagged forms are the only versions of the protein present, are viable at
500 C, therefore the lack of enrichment of immunoprecipitated origin DNA is likely not
due to destabilization and degradation of the tagged proteins. B) DnaB requires DnaD
and DnaA but not DnaI for association with the origin. Cells were synchronized with the
145
respective ts allele for 60 min at 500C for dnaAlts or 450 C for all other alleles. Binding of
DnaB to the origin was assayed by ChIP before and 10 min (dnaAlts) or 2 min (all other
alleles) after replication was allowed to proceed. C) Association of DnaB with the origin
is transient. Cells were synchronized with dnaD23tsand the association of DnaB with the
origin of replication was assayed at indicated times after release into a synchronous round
of replication.
146
In addition, we investigated the association of helicase loader DnaB with the origin
in synchronous cultures inactivated for the helicase loaders DnaD, DnaB, and DnaI (1315). Our results indicate that, in addition to DnaA, the association of DnaB with the
origin depends on DnaD, but does not depend on DnaI (Fig. lA, IB, IC). In all
experiments, association of DnaB with the origin of replication increased after the shift to
permissive temperature and then decreased (Fig. 1B, IC, and not shown), indicating that
DnaB associates with the origin, stimulates initiation, and as replication proceeds,
dissociates from the origin. These in vivo results support previous in vitro helicase loader
assembly experiments (14, 16).
We also analyzed the association of helicase with the origin using temperature
sensitive alleles of DnaB, DnaD, and DnaI. In all three cases, helicase was not detected at
the origin in cells incubated at the non-permissive temperature, but robust association
was detected within 2 min after shift to the permissive temperature (Fig. 2A, 2B,
Appendix A Fig. 1, and data not shown). As previously observed (13), the association
was transient, suggesting that helicase had largely moved away from the origin 5-10 min
after temperature downshift (Fig. 2B, Appendix A Fig. 1, data not shown). Western
blotting showed that the failure to detect helicase association with the origin at nonpermissive temperature was not due to failure to precipitate helicase from the samples
(data not shown). These data indicate that DnaB, DnaD, and DnaI are required for
helicase association with the origin in vivo.
We investigated the association of replisome components with the origin under
conditions that prevent helicase loading (dnaB134ts cells at non-permissive temperature).
We assayed the association of SSB and several replisome components with the origin of
147
A
Initiation blocked by respective ts allele
9~
.c
8
~
7
5
6
~
5
.~
4
u
• dnaA Its
~ dl1aB 134ts
rJ dnaD23ts
UJ
3
"'0
"0
D dna/2ls
2-
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o
no ab
B
helicase
DnaC
~ - clamp
DnaN
Synchronization
PollII
PolC
T -
subunit
DnaX
with dnaB134ts
16
14
e; 12
Time after replication
resumes
.c
~
c:o
10
-£'c
8
~
IOmin
m2min
c:
UJ
fA5 min
"'0
"0
lI..
noab
C
helicase
DnaC
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DnaN
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PolC
Arrested
0
30
-subunit
DnaX
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JO
25
25
0:-
0:-
2,20
810
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E
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-B
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g
15
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U
'c
r::
'C
r::
U.l
U.l
::2 10
"'0
0
'0
u...
Bprimase
OnaG
10
u...
Figure 2
dnaAlts
dnaB/34ts
dna4!ts
148
dnaB/34ts
Figure 2. A) Association of replisome components with the origin does not require
helicase. Cells were synchronized for replication with the respective ts alleles.
Association of the replication components with the origin was assayed by ChIP as
described in Fig. 1. A) Association of replication elongation components with the origin
in cells arrested for replication at non-permissive temperature with indicated ts alleles. B)
The association of indicated replication components with the origin was tested after
replication was allowed to proceed in cells synchronized with dnaB134ts. C-D) SSB and
primase DnaG associate with the origin in a DnaA dependent, helicase-independent
manner. Cells were synchronized with either dnaB134ts or dnaAlts and the association of
SSB and primase was assayed by ChIP before (C), and 2 min (dnaB134ts) or 10min
(dnaAlts) after replication was allowed to proceed (D).
149
replication. In contrast to helicase, these other replisome subunits and SSB associated
with the origin at the non-permissive temperature in dnaB134ts cells (Fig. 2A-D). The
level of association of replisome subunits or SSB did not increase after allowing helicase
loading by shifting the cells to permissive temperature (Fig.2B-D), indicating that
association of these components with the origin is likely not stimulated by the presence of
helicase. We observed similar results with other temperature sensitive proteins that block
helicase loading, dnaD23ts, dnal2ts (Fig. 2A, Appendix A Fig. 1, data not shown) and a
different allele of dnaB, dnaBl9ts (data not shown), indicating that the observations are
not allele specific. Together our results indicate that, contrary to the E. coli in vitro
model (5, 8, 17), the association of the DNA polymerase holoenzyme components with
the origin of replication does not require loading of helicase, and that DnaA function,
likely melting of the origin, may be sufficient to recruit DNA polymerase holoenzyme
and primase.
The association of replisome components with the origin of replication before the
association of helicase raises the possibility that components of the replisome may
regulate helicase loading. To test this hypothesis, we investigated the effects on DNA
replication caused by dnaN (P-clamp) or dnaX (clamp loader/T-subunit of DNA
polymerase). To monitor replication we determined DNA to protein ratios of
exponentially growing bacterial cultures (18).
Since P-clamp and clamp loader/T-subunit are essential proteins we overexpressed
these genes to look for effects on DNA replication. Overexpressing dnaN causes a nearly
two-fold increase in DNA content as compared to wild type cells (DNA to protein ratio
normalized to WT ± standard deviation from three biological replicates: WT 1.0 + 0.06;
150
dnaN overexpression 1.9 + 0.02). In contrast, overexpressing dnaX for 3-4 generations
had no effect on replication (WT 1.0 ± 0.09; dnaXoverexpression. 0.9 ± 0.05). These
data indicate that P-clamp has a positive effect on DNA replication.
To distinguish whether P-clamp stimulates initiation or elongation of DNA
replication, we tested whether the endogenous origin of replication, oriC,was required
for the effects of P-clamp overexpression. Effects on elongation should be independent of
the origin of replication. We constructed oriC inactivated strains by integrating a
different origin of replication, oriN (19), close to the location of the endogenous origin
and deleting part of oriC to generate the oriC-Sallele. Overexpression of dnaN in oriN+
oriC-S strains has no effect on replication (oriN+ oriC-S 1.4 ± 0.03; oriN+ oriC-S dnaN
overexpression 1.3 + 0.02, values normalized to WT), indicating the regulatory effects of
P-clamp on replication depend on initiation from oriC, and that overexpression of 3clamp is likely not stimulating elongation. Since replication from oriN requires the same
replication components as oriC except DnaA (19), our data indicate that P-clamp
specifically stimulates DnaA-dependent initiation of replication.
To further investigate whether P-clamp stimulates the initiation of DNA replication,
we monitored the relative genomic content of asynchronous populations of exponentially
growing cells using genomic microarrays (20). We measured the relative genomic
content for approximately 4000 genes throughout the genome and compared the genomic
content prior to and after induction of dnaN overexpression. When we overexpressed
dnaN for 25 or 50 min in an oriC+strain we saw a relative increase of origin region DNA
(Fig. 3A and data not shown). This result is typically seen in cells overinitiating DNA
replication (20). In contrast, overexpression of dnaN for 25 or 50 min in a strain
151
A
B
oriC+
1
oriN+ oriC-S
1
origin
1
~I
00
<0
........J
•
•
500kb
-1
CJlr<"ltllc..)SOI11C
•
pc..')silion
<':"-:'hrOlTlosonlC
Wild type dnaN
~ dnaN overexpressing
D
C
500kb
•
-1
position
strain
E
22
~ 20
.:
18
S
e
E
.:
II
16
14
'i: 12
,~
~
10
Ii:
8
Omin
2 min
Helicase (DnaC)
5 min
Omin
2min
Helicase loader (DnaB)
Synchronized
with dnaB134ts
Figure 3
152
Omin
2 min
DnaA
Figure 3. Overproduction of P-clamp (DnaN) stimulates DNA replication in an oriCdependent manner and affects the association of helicase and helicase loader (DnaB)
with the origin, but not the association of DnaA. A-B) The effect of P-clamp (DnaN)
overproduction on the initiation of replication was assessed by genomic microarrays.
Data were plotted as chromosomal position vs. relative genomic abundance of
chromosomal loci 50 min (- 1 generation) after addition of inducer as compared to
uninduced samples. The position of the active origin of replication is indicated by an
arrow. dnaN mRNA levels in tested cultures was similar, as assayed by microarrays. A)
Cells replicating from the endogenous, DnaA-dependent oriC.Presented results are from
a single representative experiment. B) Cells replicating independently of DnaA from
oriN. C-E) Cells were synchronized with dnaB134ts allele. dnaN was overexpressed only
at non-permissive temperature. Binding of helicase (C), helicase loader (D), and DnaA
(E) to the origin region was assayed by ChIP. Representative data from at least two
independent experiments are shown.
153
replicating from oriN had no effect on DNA replication (Fig. 3B). These observations
support the idea that P-clamp stimulates the initiation of DNA replication.
By placing the only copy of dnaN under the regulation of a xylose-inducible,
glucose-repressible promoter PxylA and growing without inducer for 3 generations, we
reduced P-clamp levels to 60% of normal. That reduction caused a 20% decrease in net
replication as assayed by DNA/protein ratio measurements. Although inhibiting P-clamp
activity, which is required for elongation of DNA replication, induces the DNA damage
SOS response ((21), data not shown), we observed no induction of the DNA damage SOS
response in our cells, indicating that the decrease in replication is likely due to decreased
rate of replication initiation, rather than elongation (data not shown).
To understand how P-clamp affects initiation, we monitored the association of
DnaA, DnaB and helicase with the origin in cells overexpressing dnaN. In control cells
that are blocked for initiation with the dnaB134ts allele, helicase and helicase loader
(dnaB) are not associated with the origin (Fig. 2A, 2C, 3C, 3D). However, 2 min after
release from this block, there is a distinct increase in DnaB and helicase binding to the
origin as assayed by ChIP (Fig. 2C, 3C). In similarly synchronized cells, overexpression
of dnaN caused -2-fold increase in association of DnaB and helicase at the origin after 2
min (Fig.3C-3D). These results indicate that P-clamp increases replication initiation by
stimulating the association of DnaB with the origin, which increases the recruitment of
helicase to the origin and thus likely speeds up the timing of the next replication initiation
event the. Consistent with this interpretation, thymidine incorporation experiments show
that overexpression of dnaN cause an increase in the amount of DNA replication after
release of dnaB134ts cells into synchronous replication (data not shown).
154
In contrast to helicase loader (DnaB) and helicase, our results indicate that the
association of DnaA with the origin is not increased by overexpression of dnaN (Fig.3E).
We tested the association of DnaA with the origin before and after replication initiation
and observed no differences in the binding of DnaA to the origin (Fig.3E) in cells
overexpressing dnaN as compared to control cells. These results indicate that P-clamp
may increase replication initiation by regulating a step after association of DnaA but
before association of DnaB with oriC. P-clamp could regulate the interaction of DnaA
with itself or other replication initiation proteins, such as DnaD (22).
In summary, our analysis revealed that, as expected, DnaA is required for the
association of replication proteins with the origin of replication. Unexpectedly, we
observed that the association of DNA polymerase holenzyme, primase, and SSB with the
origin was independent of helicase (Fig. 4). Our observations raise an interesting
question: How is the replication machinery recruited to the origin of replication in the
absence of helicase? In vitro, DnaA and SSB are not sufficient to recruit E. coli primase
and DNA polymerase to the origin of replication (5). Since we observe association of
DNA polymerase holoenzyme subunits with the origin in the absence of replicative
helicase, it is possible that there are different requirements for DNA polymerase binding
to the origin in B. subtilis and E.coli. Alternatively, there may be a significant difference
between what occurs at the origin in vivo versus in vitro. It is also possible that the ChIP
method we employed is more sensitive than the size-exclusion method used in the E. coli
in vitro studies (5), and thus we detect association of replication components with the
origin before helicase. Although P-clamp is likely to be loaded at an RNA primer
(reviewed in (23)), E. coli T-subunit (DnaX) can bind DNA (24), and P-clamp can be
155
Melting of the origin
----+
Replisome association
-
Helicase loading
f3-clamp
~
t
DnaA
Primase
Figure 4
156
.......
SSB
Helicase
Figure 4. Model for the order of association of replisome components to the origin of
replication in B. subtilis. Our evidence indicates that after DnaA performs an essential
function for DNA replication, likely origin unwinding, SSB, primase, P-clamp, T-clamp
loader/leading and lagging strand dimerisation subunit, and the catalytic a-subunit of
DNA polymerase associate with the origin of replication. We propose that the association
of P-clamp stimulates the association of helicase loader and helicase with the origin of
replication, and after helicase is loaded, replication elongation proceeds. The two salient
points of this model are that the association of many replisome components with the
origin does not require helicase, and that helicase loading is regulated by components of
the replisome. We speculate that loading of P-clamp at a primer or at a dsDNA-ssDNA
structure by the T-clamp loader is central for the recruitment of the catalytic DNA
polymerase subunit to the origin. The interactions that bring primase to the origin in the
absence of helicase are unknown.
157
loaded at ssDNA-dsDNA forked structure (25), which is generated during origin melting
(9). These biochemical properties of the DNA polymerase holoenzyme may allow it to
associate with the origin in the absence of replicative helicase. Alternatively, primase,
which we also detect at the origin prior to helicase, may be capable of limited primer
synthesis, thus allowing the loading of P-clamp and the stable association of the
holoenzyme.
Our experiments also demonstrate that P-clamp (DnaN) regulates the initiation of
DNA replication at the DnaA-dependent origin of replication, oriC.P-clamp appears to
activate a step after DnaA association with the origin, but before or at the time of helicase
loader (DnaB) association with the origin of replication. One attractive mediator of this
regulation may be YabA, which regulates replication initiation in B. subtilis and interacts
directly with both DnaA and P-clamp (26, 27).
Exposed ssDNA that is not rapidly duplicated by the DNA replication machinery
can lead to recombination events, DNA breaks, induced mutagenesis, and genomic
instability. Mechanisms that couple the functions of the helicase and DNA polymerase
may promote genomic stability by limiting the production of ssDNA in the absence of
DNA synthesis. During DNA replication, DNA unwinding is coupled to DNA synthesis
by interactions between helicase and the t-subunit of DNA polymerase holoenzyme (28).
Here we propose that a mechanism exists to prevent the extensive production of ssDNA
at the origin, in which helicase loading is stimulated by the presence of the DNA
polymerase holoenzyme.
Our results indicate that in bacteria a macromolecular complex, including DNA
polymerase holoenzyme, assembles early during initiation, before assembly of helicase
158
and not as the last step before elongation starts. It appears that in both bacteria and
eukaryotes, components of the replisome can associate with the origin of replication and
regulate subsequent events that are required for replisome function and the initiation of
DNA replication. These multiple layers of regulation likely serve to increase genomic
stability, thereby allowing proper cell cycle completion and successful generation of
viable progeny.
Acknowledgements
We thank M. M. Berkmen, K. Lemon, L. Simmons, and J.D. Wang for generous
provisions of plasmids and strains. We thank T. Baker, S. P. Bell, F. Solomon, A. Wright,
C. Lee, J. Wang, and J. Auchtung for suggestions and comments on the manuscript.
159
References
1.
S. P. Bell, A. Dutta, Annu Rev Biochem 71, 333-74 (2002).
2.
H. Masumoto, A. Sugino, H. Araki, Mol Cell Biol 20, 2809-17 (Apr, 2000).
3.
S. Hiraga, A. Hagihara-Hayashi, T. Ohya, A. Sugino, Genes Cells 10, 297-309
(Apr, 2005).
4.
M. Izumi et al., Nucleic Acids Res 28, 4769-77 (Dec 1, 2000).
5.
L. Fang, M. J. Davey, M. O'Donnell, Mol Cell 4, 541-53 (Oct, 1999).
6.
B. E. Funnell, T. A. Baker, A. Kornberg, JBiol Chem 261, 5616-24 (Apr 25,
1986).
7.
A. Kornberg, T. A. Baker, DNA Replication (W.H. Freeman and Company, New
York, ed. Second, 1992).
8.
W. Messer, FEMS MicrobiolRev 26, 355-74 (Nov, 2002).
9.
W. Messer et al., Biochimie 83, 5-12 (Jan, 2001).
10.
A. Yuzhakov, J. Turner, M. O'Donnell, Cell 86, 877-86 (Sep 20, 1996).
11.
S. Moriya, K. Kato, H. Yoshikawa, N. Ogasawara, Embo J9, 2905-10 (Sep,
1990).
12.
E. Le Chatelier et al., J Biol Chem 279, 1757-67 (Jan 16, 2004).
13.
M. E. Rokop, J. M. Auchtung, A. D. Grossman, Mol Microbiol 52, 1757-67 (Jun,
2004).
14.
M. Velten et al., Mol Cell 11, 1009-20 (Apr, 2003).
15.
D. Karamata, J. D. Gross, Mol Gen Genet 108, 277-87 (1970).
16.
S. Marsin, S. McGovern, S. D. Ehrlich, C. Bruand, P. Polard, JBiol Chem 276,
45818-25 (Dec 7, 2001).
17.
T. A. Baker, K. Sekimizu, B. E. Funnell, A. Kornberg, Cell 45, 53-64 (Apr 11,
1986).
18.
R. Kadoya, A. K. Hassan, Y. Kasahara, N. Ogasawara, S. Moriya, Mol Microbiol
45, 73-87 (Jul, 2002).
19.
A. K. Hassan et al., J Bacteriol 179, 2494-502 (Apr, 1997).
20.
L. A. Simmons, A. M. Breier, N. R. Cozzarelli, J. M. Kaguni, Mol Microbiol 51,
349-58 (Jan, 2004).
21.
Y. Ogura, Y. Imai, N. Ogasawara, S. Moriya, JBacteriol 183, 3833-41 (Jul,
2001).
22.
D. Ishigo-Oka, N. Ogasawara, S. Moriya, JBacteriol183, 2148-50 (Mar, 2001).
23.
A. Johnson, M. O'Donnell, Annu Rev Biochem 74, 283-315 (2005).
24.
F. P. Leu, R. Georgescu, M. O'Donnell, Mol Cell 11, 315-27 (Feb, 2003).
25.
N. Yao, F. P. Leu, J. Anjelkovic, J. Turner, M. O'Donnell, JBiol Chem 275,
11440-50 (Apr 14, 2000).
26.
M. F. Noirot-Gros et al., Proc NatlAcadSci USA 99, 8342-7 (Jun 11, 2002).
27.
M. F. Noirot-Gros et al., ProcNatl Acad Sci USA 103, 2368-73 (Feb 14, 2006).
28.
S. Kim, H. G. Dallmann, C. S. McHenry, K. J. Marians, Cell 84, 643-50 (Feb 23,
1996).
160
Appendix A
Supplementary Materials and Methods for Chapter 4:
Early association of replisome components with the origin of replication
provides a mechanism for regulating the initiation of DNA replication
161
Genetic methods and strain construction. B. subtilis strains are listed in Supplementary
Table 1. Genetic manipulations were performed using standard protocols (1).
The C-terminal fusion of dnaN to GFP was generated by cloning the 3' 300bp of
dnaN without the stop codon into plasmid pLS31 (gift from Lyle Simmons) containing
monomeric GFP (A206--K) (2) and a 23 amino acid linker. The resulting plasmid
(pAIG47) was integrated at the dnaN locus on the chromosome through a single
crossover. To generate tagged version of primase DnaG, the entire dnaG ORF was cloned
into pEA 18 (3) generating a translational fusion of GFP and DnaG in plasmid pJW 120,
(gift from J. Wang). The gfp-dnaG gene was regulated by the xylose inducible promoter
PxylA. The resulting plasmid was integrated at the amyE locus by double crossover in
strains carrying various temperature sensitive alleles. The N-terminally tagged protein
was at least partially functional as it complemented a temperature sensitive allele of
primase, dnaG20 (data not shown). polC-myc strain were constructed by integrating
through a single cross-over the plasmid pKL 159 (a gift from Katherine Lemon)
containing the 3' -300bp ofpolC fused to 3x c-myc tag.
To generate a construct where the only copy of DnaN is regulated by PxylA, we
amplified a region of the genome of MMB26 (4) that included the PxylA promoter and
the 5' end of the dnaN gene. The amplified fragment was cloned into plasmid pGEMcat
resulting in plasmid pAIG28. pAIG28 was integrated in the genome of JH642 by a single
crossover to generate strain AIG260. The dnaAl allele was previously described (5).
dnaB134, dnaB19, dnaD23, dnal2 are standard temperature sensitive alleles for these
genes (6, 7). Other strains and alleles are listed in Table 1.
162
Chromatin immunoprecipitation. Strains were grown in defined minimal medium with
50gg/ml spectinomycin (to maintain selection for fusion proteins generated by a single
cross-over into the chromosome) at 300 C to mid-exponential phase (OD600 = 0.3). Cells
were shifted to 450 C (for dnaBts, dnaDts, and dnalts strains) or 500C (for dnaAts strains)
for 60 min to prevent initiation of replication and to allow most ongoing rounds of
replication to finish. In experiments where dnaN was overexpressed, inducer was added
to cultures upon shift to non-permissive temperature, to limit the effects of the
overexpression only to the replication cycles after synchronization. Cells were then
rapidly shifted back to 300 C (or 35 OC in the case of dnaAlts experiments) by adding an
equal volume of medium that was at 200 C. Samples were taken at various times and
protein and DNA were cross-linked with 1% formaldehyde for 5 min at room
temperature. Glycine was added (120 mM), and samples were incubated for 5 min at
room temperature to stop the cross-linking. Further processing, cell lysis and
immunoprecipitations were done essentially as described previously (6) with the
following exceptions. Immunoprecipitations for myc-tagged and GFP-tagged proteins
and DnaB were performed overnight at 40 C using mouse anti-c-Myc antibody (Zymed)
at 6 mg per immunoprecipitation, 1:500 dilution of a rabbit polyclonal anti-GFP
antibody, and 1:1 000 dilution of affinity purified, polyclonal rabbit anti-DnaB antibody,
respectively. DnaA was immunopercipiteted for 1h at room temperature with 1:10 000
dilution of chicken anti-DnaA antibody and was then incubated for h with 1:500 dilution
of a donkey anti-chicken secondary antibody. Samples for all IP reactions were then
incubated with 30p1 50% Protein A-Sepharose bead slurry (Pharmacia Biotech) for I h at
40 C (room temperature for DnaA IP). Beads were washed six times with lml 50mM Tris-
163
HCI (pH 7), 150mM NaCI, 5mM EDTA, 1% Triton X-100, and then washed two times
with 1ml TE, three minutes per wash for all washes. Protein was eluted from the beads by
incubation of the beads in l00gl 10mM EDTA, 1% SDS, 50mM Tris-HCI (pH 8) for 10
min at 650 C. Beads were further washed with 150 p.l TE/0.67%SDS. The wash was
combined with the 100 p.l eluate. Cross-links were reversed overnight at 650 C, and DNA
was purified essentially as described previously (6).
DNA in the immunoprecipitates was analyzed by PCR using primers to the oriC
region and to an origin-distal site (dnaD, which is located -1860 kbp from oriC). PCR
was done on two-fold serial dilutions of template DNA to obtain products in the linear
range of PCR reactions. PCR done with total DNA as a template (obtained from cell
lysates before antibody addition) allowed us to control for primer efficiency and copy
number of tested loci. PCR was performed for 27 to 31 cycles depending on amount of
0 C, 1 min extension step at 720
DNA precipitated, with 1 min annealing step at 55
C, and
30 sec denaturation step at 950 C for each cycle. Products were analyzed on 2% agarose
gels stained with ethidium bromide using the Alpha Immager gel documentation system
and the images were analyzed using Gellmager5.2 software. This allowed us to determine
the relative amounts of DNA from the origin region compared with the origin-distal site.
Fold enrichments were calculated by obtaining the fraction DNA precipitated for the
origin region and dividing it by the fraction precipitated for the distal region [(oriC IP/
oriC total)/( dnaD IP/ dnaD total)]. Similar results were obtained with other origin-distal
control loci.
Media and growth conditions. For all experiments, cells were grown with vigorous
shaking at 300 C or 370 C in S7 defined minimal medium with MOPS buffer at a
164
concentration of 50 mM rather than 100 mM (8); the medium was supplemented with
0.1%glutamate, the required amino acids (at 40 ýtg/ml), and 1% glucose (or 1%
arabinose in experiments where PxylA-dnaN constructs). Strains containing single
crossover constructs were routinely grown in 100p~g/ml spectinomycin (Sigma), or in
50gg/ml spectinomycin when grown for ChIP.
Genomic microarrays to study DNA replication. DNA microarrays were prepared
using PCR products from >99% of the annotated B. subtilis open reading frames spotted
onto Corning GAPS slides, essentially as described previously (4). dnaN overexpression
was induced in exponentially growing, asynchronous cultures with 0.5% xylose and
samples were collected at various times before and after induction by mixing samples
with an equal volume of ice cold methanol. Samples were processed as previously
described (4). Test and reference samples (DNA obtained from cells with a single
unreplicated chromosome) were coupled to Cy5 and Cy3 dyes respectively, mixed, and
hybridized to a microarray as previously described (4), to obtain the ratios of test sample
to reference for each chromosomal locus. Different test samples were then compared by
obtaining the ratio of these ratios. Scanning, analysis, and normalization was preformed
as previously described (4)
DNA/protein ratio determination. The ratio of DNA to protein was determined as
previously described (9, 10). Briefly, 25ml of exponentially growing cells were collected
at an OD600 50.6. The ratio of the amounts of DNA and protein recovered for each
culture was calculated. The ratios for all strains were normalized to WT (WT= 1.0)
165
grown on the same day and under the same conditions. The average of three biological
replicates is presented with error bars representing standard deviation. The regulation of
initiation of DNA replication at oriN differs from that at oriC,and as a result oriN+ oriCS strains overreplicate in minimal media as compared to oriC strain.
166
Association with the origin in dnaD23ts cells
Clamp loader/t-subunit (DnaX)
helicase (DnaC)
Omin lmin
[Template] ....
Smin lOmin
I....I~ I~
oriC
IP
Distal
Region
Omin lmin
[Te~late]
....
Omin lmin
Smin lOmin
I....J~ I~
Smin lOmin
.... I....I~ I~
Input
IEnrichment 7.3
6.2
3.5
0.9
2.0
Figure 1
167
19.3 19.0 5.2
I
Figure 1. Association of clamp loader/k-subunit (DnaX) and helicase (DnaC) with
the origin of initiation in dnaD23ts cells. Cells were synchronized with dnaD23ts and
data are shown for association of helicase or clamp loader/t-subunit with the origin at
various times after release of replication by temperature downshift at 0 min. Calculated
fold enrichments are indicated.
168
Table 1. B. subtilis strains used
Strains
Relevant Genotype (reference)
JH642
trpC2pheAl (11)
KPL502
trpC2 pheAl dnaBl34SQTn917mls dnaX:.:dnaX-GFP-spc(12)
MER489
trpC2 pheAl dnaB134QTn917mlsdnaC::dnaC-myc-spc(6)
MMB26
trpC2 pheAl amyE::PxylA-dnaN-cat(4)
MMB 170 pheAl spoIIIJ..oriN-kan oriC-S (ypjG-hepT)122
AIG260
trpC2 pheA I dnaN::PxylA-dnaN-cm (pAIG28)
AIG278
pheAl spoll1J:.:oriN-kanoriC-S amyE.:.PxylA-dnaN-spc (ypjG-hepT)122
AIG380
trpC2 pheAl dnaB134QTn917mlsdnaC.::dnaC-myc-spcamyE:.:PxylA-dnaN-cat
AIG394
trpC2 pheA 1 dnaB134QTn917mlspolC:.'polC-myc-spc
AIG396
trpC2 pheA 1 dnaB134QTn917mlsdnaN:.:dnaN-GFP-spc
AIG456
trpC2 pheA I dnaA lQTn917mls polC::polC-myc-spc
AIG458
trpC2 pheA 1 dnaA IQTn91 7mls dnaC.":dnaC-myc-spc
AIG460
trpC2 pheA 1 dnaA1QTn917mls dnaN: :dnaN-GFP-spc
AIG462
IrpC2 pheA 1 dnaA I2Tn917mls dnaX.::.dnaX-GFP-spc
AIG475
IrpC2 pheA 1 dnaD23QTn917mlspolC:.polC-myc-spc
AIG477
trpC2 pheA 1 dnaD23QTn917mlsdnaC::dnaC-myc-spc
AIG479
trpC2pheA 1 dnaD23QTn917mlsdnaN..::dnaN-GFP-spc
AIG481
trpC2pheA 1 dnaD232Tn917mls dnaX: :dnaX-GFP-spc
A[G511
trpC2pheAl dnaI2QTn917mls dnaC.:dnaC-myc-spc
AIG513
trpC2pheA I dnal2•2Tn917mls polC::polC-myc-spc
169
AIG515
trpC2 pheA 1 dnaI2QTn917mls dnaN:.:dnaN-GFP-spc
AIG517
trpC2 pheAl dnaI2QTn917mls dnaX::dnaX-GFP-spc
AIG537
trpC2 pheA 1 dnaBl9QTn917mls dnaC:.:dnaC-myc-spc
AIG538
trpC2 pheAl dnaBl9QTn917mlspolC::polC-myc-spc
AIG540
trpC2 pheA 1 dnaBl9QTn917mls dnaN:.dnaN-GFP-spc
AIG542
trpC2 pheA 1 dnaBl9QTn917mls dnaX::dnaX-GFP-spc
AIG550
trpC2 pheA I dnaA l2Tn917mls amyE.:.PxylA-GFP-dnaG-cat
AIG552
trpC2 pheA dnaB1340Tn917mls amyE.::PxylA-GFP-dnaG-cat
AIG554
trpC2pheAl dnaAlQTn917mls lacA::Pssb-ssb-GFP-tet
AIG556
trpC2 pheAl dnaBl342Tn917mls lacA:.:Pssb-ssb-GFP-tet
170
References
1.
C. R. Harwood, S. M. Cutting, Molecular BiologicalMethods for Bacillus (John
Wiley & Sons, Chichester, England, 1990).
2.
D. A. Zacharias, J. D. Violin, A. C. Newton, R. Y. Tsien, Science 296, 913-6
(May 3, 2002).
3.
F. J. Gueiros-Filho, R. Losick, Genes Dev 16, 2544-56 (Oct 1, 2002).
4.
A. I. Goranov, L. Katz, A. M. Breier, C. B. Burge, A. D. Grossman, Proc Natl
Acad Sci USA 102, 12932-7 (Sep 6, 2005).
5.
S. Moriya, K. Kato, H. Yoshikawa, N. Ogasawara, Embo J 9, 2905-10 (Sep,
1990).
6.
M. E. Rokop, J. M. Auchtung, A. D. Grossman, Mol Microbiol52, 1757-67 (Jun,
2004).
7.
D. Karamata, J. D. Gross, Mol Gen Genet 108, 277-87 (1970).
8.
K. J. Jaacks, J. Healy, R. Losick, A. D. Grossman, JBacteriol 171, 4121-9 (Aug,
1989).
9.
P. S. Lee, A. D. Grossman, Mol Microbiol 60, 853-69 (May, 2006).
R. Kadoya, A. K. Hassan, Y. Kasahara, N. Ogasawara, S. Moriya, Mol Microbiol
10.
45, 73-87 (Jul, 2002).
11.
M. Perego, G. B. Spiegelman, J. A. Hoch, Mol Microbiol2, 689-99 (Nov, 1988).
12.
K. P. Lemon, A. D. Grossman, Mol Cell 6, 1321-30 (Dec, 2000).
171
Chapter 5
P-clamp and YabA regulate the initiation of DNA replication in
Bacillus subtilis
172
Abstract
Proper regulation of DNA replication is essential for successful completion of the
cell cycle. In the gram-positive bacterium Bacillus subtilis, the initiation of DNA
replication is affected by the P-clamp (DnaN) and a small protein, YabA. YabA is
thought to regulate the replication initiation protein DnaA. Both P-clamp and YabA,
when fused to GFP, form foci that co-localize with the replisome. Here we show that Pclamp and YabA act in the same regulatory pathway, and that the formation of YabA foci
depends on 0-clamp. In contrast, the localization of YabA to the replisome does not
require DnaA or DnaA-dependent replication. We also show that YabA and P-clamp do
not affect DnaA-dependent transcriptional regulation, indicating that YabA and p-clamp
do not regulate the initiation of DNA replication by affecting the level of free DnaA. Our
results are consistent with a model in which 3-clamp and YabA specifically regulate a
step downstream of DnaA binding to the origin of replication.
173
Introduction
During each cell cycle, cells grow, replicate their genome, and divide to form two
new cells. These processes are coordinated, such that the frequency of chromosome
replication is proportional to the rate of cell growth, and that cell division occurs after the
chromosomes have replicated. Replication is typically coordinated with cell mass, such
that cells begin replicating their genome after they have reached some critical mass for a
given growth condition (16, 35, 48). Since the availability of nutrients determines how
fast organisms grow and reach their critical mass, the initiation of DNA replication is
regulated by nutrient availability in many different organisms. In bacteria, nutrient
availability can also regulate replication elongation (3, 12, 34), although growth rate
appears to regulate DNA replication largely at the level of initiation (59). How cell mass
regulates the initiation of DNA replication is not well understood.
The regulator of initiation of DNA replication in bacteria is DnaA, a protein that is
highly conserved among bacteria and is analogous to ORC components in eukaryotes
(38). DnaA is a member of the AAA+ class of proteins, binds ATP or ADP, and has a
weak ATPase activity. DnaA is active for initiation only when in the ATP-bound form
(52). DnaA is also a transcription factor, and appears to coordinate replication status with
cell-cycle and developmental processes (8, 13, 14, 40). During replication initiation,
DnaA binds to the chromosomal origin of replication (oriC) in a sequence specific
manner and can melt the origin to expose ssDNA (38, 39). The exposed ssDNA is
thought to serve as a landing pad for other replication proteins (38, 39).
Besides melting the origin, DnaA also functions in loading helicase at oriC,in both
Escherichiacoli and Bacillussubtilis. Helicase is a component of the replisome that
174
unwinds the chromosome and exposes ssDNA template for DNA polymerase (4, 28). In
E. coli, DnaA and helicase interact directly (11, 36, 51, 55), whereas in B. subtilis DnaA
interacts with DnaD, which in complex with two other proteins mediates helicase loading
(20).
The function of DnaA is regulated by at least three mechanisms in the gramnegative bacterium E. coli. The expression of dnaA is regulated in a cell cycle-dependent
manner through auto-repression and through repression by SeqA when the DNA around
the dnaA locus is hemi-methylated during replication (1, 5, 10, 19, 46). DnaA is also
sequestered by a locus on the chromosome, datA, that has high affinity for DnaA, such
that deletion of datA affects replication by freeing up DnaA molecules (41, 47). The
activity of DnaA is also regulated by conversion between the ATP/ADP bound states of
DnaA through the action of P-clamp and Hda. P-clamp is a component of the replisome
which encircles DNA forms a topological link between DNA polymerase and DNA,
stimulating processive replication (22). Hda is a protein homologous to DnaA which
when interacting with P-clamp loaded onto DNA stimulates the ATPase activity of DnaA
(25, 26, 30, 43, 54). Once replication starts, P-clamp, and Hda stimulate the ATPase
activity of DnaA, converting DnaA into the inactive, ADP-bound form, which prevents
DnaA from re-initiating DNA replication during the came cell-cycle (24, 30, 43).
The initiation of DNA replication in the gram-positive bacterium B. subtilis is
regulated by growth rate and by DnaA. As in E. coli, the expression of dnaA is autorepressed and overexpression of DnaA causes excessive replication and reduces the size
of cells at the time of initiation (48). These observations indicate that the regulation of
DnaA may help coordinate replication and growth. However, DNA methylation does not
175
appear to regulate replication in B. subtilis as it does in E. coli, since B. subtilis does not
have a recognizable SeqA homologue and has very limited DNA methylation (29). There
is also no Hda homologue in B. subtilis, other than DnaA, indicating that conversion
between the ATP/ADP states of DnaA may be regulated by a different mechanism and
possibly different adaptor proteins.
Replication initiation in B. subtilis is regulated by YabA, whose homologues are
restricted to gram-positive bacteria (18, 44, 45). YabA interacts with DnaA and P-clamp
(44, 45). YabA forms foci that localize to the replisome throughout the cell-cycle (18,
45). The mechanism by which YabA regulates DNA replication is not known, although
mutational analysis of YabA has led to the hypothesis that YabA regulates initiation by
sequestering DnaA away from the origin thus affecting the rate of replication initiation
(45). In order to understand the requirements for localization and YabA function, mutants
of YabA that appear defective in interaction with either DnaA or P-clamp were isolated
using a yeast-two-hybrid screen (45). Both types of mutants had yabA-null replication
phenotype and did not form foci when fused to GFP (45). Based on these observations it
was concluded that YabA interacts with both DnaA and P-clamp in order to form foci
and regulate replication (45). Recently, we showed that P-clamp regulates DnaAdependent initiation of replication by affecting a step after association of DnaA with oriC
but before helicase loading (Chapter 4).
Here, investigate the regulation of the initiation of DNA replication by YabA and
P-clamp. We found that YabA and P-clamp act in the same regulatory pathway, and, as
with P-clamp, YabA regulates DnaA- and oriC-dependent initiation of replication. Our
results indicate that localization of YabA with the replisome did not require DnaA or
176
DnaA-dependent replication. The association of YabA with the replisome is likely
mediated by P-clamp. Neither f-clamp nor YabA affected the expression of DnaA
regulated genes. These results are not consistent with the hypothesis that YabA and fclamp affect the level of DnaA. Instead, our observations are consistent with a model
where YabA and P-clamp specifically affect the activity of DnaA at the origin of
replication.
Materials and Methods
Genetic methods and strain construction. B. subtilis strains are listed in Table 1.
Genetic manipulations were performed using standard protocols (17). The yabA deletion
(AIG109) removes the entire yabA ORF and was generated by the long-flanking
homology PCR method (58). The deletion starts at the Ist codon (TTG) and ends 50bp
downstream the translational stop, removing a total of 407bp. The deletion stops 13bp
upstream of the next gene, yabB. The yabA ORF is substituted with the 994bp
chloramphenicol resistance cassette from pGEMcat. The cassette contains the cat ORF,
322bp upstream to include the promoter, and 20bp downstream of the stop codon, which
does not include the transcriptional terminators. The expression of yabB and other
downstream genes in AIGI09 is wild type level as assessed by microarray analysis (data
not shown). AIG80 was constructed by cloning the entire ORF of yabA with its
endogenous ribosome binding site into a plasmid containing the Pspank-(hy) promoter
(pDR66, a gift from David Rudner) thus generating plasmid pAIGIO0. The Pspank-(hy)yabA construct was integrated into the genome of JH642 through a double crossover at
the amyE locus to generate strain AIG80. To generate a construct where the only copy of
177
13-clamp is regulated by PxylA, we amplified a region of the genome of MMB26 that
included the PxylA promoter and the 5' end of the dnaN gene. The amplified fragment
was cloned into plasmid pGEMcat resulting in plasmid pAIG28. pAIG28 was integrated
in the genome of JH642 by a single crossover to generate strain AIG260. GFP-YabA
construct was obtained by cloning the entire yabA ORF in frame with GFP in the pAEI 8
plasmid (15). The resulting plasmid (pAIG58) was integrated into the chromosome of
strains through double cross over at the amyE locus. The GFP-YabA fusion protein was
functional as it complemented phenotypic characteristics ofyabA-null (data not shown)
(45). Other strains and alleles are listed in Table 1.
Media and growth conditions. For all experiments, cells were grown with
vigorous shaking at 370 C in S7 defined minimal medium with MOPS
(morpholinepropanesulfonic acid) buffer at a concentration of 50 mM rather than 100
mM (21); the medium was supplemented with 0.1% glutamate, the required amino acids
(at 40 [tg/ml), and 1% glucose or 1% arabinose in experiments where PxylA-dnaN and
PxylA-GFP-yabA constructs were used. For fast growth conditions, strains were grown
in LB at 370 C. dnaN and GFP-yabA were induced with xylose at 0.5% final
concentration, and yabA and dnaB371 were induced with 1mM IPTG. Strains containing
single crossover constructs were routinely grown in 100gg/ml spectinomycin (Sigma).
DNA/protein ratio determination. The ratio of DNA to protein was determined
as previously described (23, 31). Briefly, 25ml of exponentially growing cells were
collected at an OD600 at or bellow 0.6. The ratio of the amounts of DNA and protein
recovered for each culture was calculated. The ratios for all strains were normalized to
178
WT (WT = 1.0) grown on the same day and under the same conditions. The average of
three biological replicates is presented with error bars representing standard deviation.
Western blotting. Immunoprecipitated protein samples were obtained from cells
grown to 0D600 -0.5 in minimal medium at 370 C. Samples were processed and western
blots were analyzed as described previously (50), with the exception that membranes
were probed with chicken primary anti-DnaA, antibody diluted 1:10 000 for 1h at room
temperature, washed 3 times as previously described (50), and then probed with antichicken, HRP-conjugated antibody (Invitorgen) 1:500 dilution.
Use of DNA microarrays for gene expression profiling. DNA microarrays were
prepared either using PCR products from >99% of the annotated B. subtilis open reading
frames spotted onto Coming GAPS slides, or 65-mer oligonucleotide library representing
all of the annotated ORF of the B. subtilis genome (Sigma-Genosys) essentially as
described previously (2, 6, 14). Oligonucleotide microarrays were used only for the yabA
set of experiments.
Exponentially growing cultures were treated with HPUra to arrest replication, or with
IPTG or xylose to induce yabA or dnaN expression respectively, samples were collected
immediately before and 15 min, 30 min, and 60 min after treatment and processed as
previously described to generate labeled cDNA (2, 14). For comparisons of yabA with
wild type cultures, samples were collected form exponentially growing cultures of WT
(JH642) and yabA-null cultures (AIG109). A similarly processed reference sample was
hybridized with each experimental sample for normalization. The reference sample
contained pooled total RNA from cell cultures grown in defined minimal medium and
cultures treated with DNA damaging agents, thus ensuring that all genes expressed under
179
Table 1. B. subtilis strains used
Strains
Relevant Genotype (reference)
JH642
trpC2pheAl (49)
IRN444
trpC2pheA I recA260::Tn917-mis-cat(33)
MMB26
trpC2pheAl amyE: :PxylA-dnaN-cat (14)
MMB 170 pheAl spolllJ::.oriN-kanoriC-S (ypjG-hepT)122 (14)
MER616
trpC2pheA1 amyE::Pspank-DnaA-spc (50)
AIG80
trpC2pheA 1 amyE::.Pspank-hy-yabA-spc
AIG 109
trpC2pheAl AyabA::cat
AIG 181
trpC2pheAl recA260::Tn917-mls-catAyabA::cat
AIG185
pheAl spolllJ::oriN-kanoriC-S AyabA::cat (ypjG-hepT)122
AIG 189
trpC2pheAl AyabA::cat amyE..:Pspank-hy-dnaB371-spc
AIG208
pheAl spolllJ..::oriN-kanoriC-S amyE.':Pspank-hy-yabA-spc (ypjG-hepT)122
AIG244
trpC2pheA 1 AyabA::cat amyE..::PxylA-dnaN-cat.::tet
AIG260
trpC2pheAl dnaN::PxylA-dnaN-cm (pAIG28)
AIG267
trpC2pheA I amyE::Pspank-DnaA-spc,dnaN: :PxylA-dnaN-cat
AIG388
trpC2pheAl amyE:.:Pspank-DnaA-spc,dnaN.':PxylA-dnaN-mis, AyabA::cat
AIG470
trpC2pheA amyE::PxylA-GFP-yabA-cm (pAIG58)
AIG505
pheAl spolIlJ:.:oriN-kanoriC-S amyE.::PxylA-GFP-yabA-cm (ypjG-hepT)122
AIG593
pheAl spoIIJ::oriN-kanoriC-S amyE::PxylA-GFP-yabA-cm:.:mls(ypjGhepT)122 dnaA.:.:Pspac-hy-dnaN-cm(dnaA-null)
AIG595
pheAl spolllJ.:.:oriN-kanoriC-S amyE.::PxylA-GFP-yabA-cm::mls(ypjGhepT)122 dnaA .:Pspac-hy-dnaA-dnaN-cm
180
those conditions are represented in the sample. Our microarray analysis includes every
spot that has 280% of the pixels at least one standard deviation over background in one or
both Cy3 or Cy5 channels.
All microarray experiments were done with at least three independent replicates and
evaluated using Significance Analysis of Microarrays (SAM)(57). For all experiments,
we picked the most stringent criteria that resulted in a predicted number of false positives
of one or less.
Microscopy. Microscopy was performed essentially as described (32). Briefly,
cells were placed on 1% agarose pads, and images were captured with a Nikon E800
microscope equipped with a Hamamatsu digital camera. Improvision OpenLabs 2.0
software was used to process images.
Results
P-clamp and YabA are likely on the same pathway that regulates the initiation
of replication. It was previously shown by microscopy and flow cytometry that YabA is
a negative regulator of replication (18, 44, 45). We verified these results by monitoring
DNA to protein ratios in cells deleted for yabA or cells overexpressing yabA. As
predicted, in a yabA-null mutant, the DNA to protein ratio increased - 2-fold, whereas in
cells overexpressing yabA, the DNA to protein ratio decreased about 40% (Fig. lA). As
with overexpression of P-clamp, which increases the DNA to protein ratio, the effect of
yabA deletion was more severe in minimal medium than in rich medium (Fig.1A, IC,
1D) (44).
181
Deletion of yabA causes overreplication, which leads to the induction of genes
involved in recombination and in DNA damage repair, including the expression of the
recombination protein RecA (data not shown). In E.coli, DNA damage can cause RecAdependent replication of the chromosome, termed stable inducible DNA replication
(siDNA replication), which is independent of the endogenous origin of replication (27).
To address whether siDNA replication is responsible for the over-replication in yabA-null
cells, we tested if deletion of yabA affects replication in a strain deleted for recA. The
effects of yabA deletion on DNA/protein ratio were still manifested in the absence of
recA (Fig. IA), indicating that the effects are not through siDNA replication. Consistent
with these observations, we found that for yabA to affect DNA/protein ratio, the strains
need to initiation replication from oriC.We constructed oriC inactivated strains by
integrating a different origin of replication, oriN (23), close to the location of the
endogenous origin and deleting part of oriCto generate the oriC-S allele. Neither deletion
nor overexpression of yabA had any effect in oriN' oriC-S strains (Fig. 1B).
Since YabA and P-clamp affect the initiation step of DNA replication and both
require DnaA-dependent initiation of replication from oriC to manifest their effects, it is
plausible that P-clamp and YabA affect the initiation of DNA replication via the same
regulatory pathway. Indeed, yeast-two-hybrid assays indicate that YabA and P-clamp
interact directly (44, 45), (Goranov and Grossman, unpublished observations). To test
whether YabA and P-clamp are on the same pathway, we generated a strain that contains
a deletion of yabA and overexpresses P-clamp (PxylA-dnaN). Each single allele causes
overreplication as determined by DNA to protein ratios (Fig. lA, IC). If P-clamp and
YabA are on the same pathway, it is expected that the two mutations will not have an
182
B
,
2.l
2.5
oriN oriC-S
2
o
I-
r
1.5
1.5
61.
0.•
e
0.,
0.5-
..
WT
yabbA-null
Piptg -ytabA
recA260
recA260,
yiabA-null
ycabA-null
vahbA+
Piptg -yab.4
2.5
2
v.
'U
1.
CI
0.
.
z
0
WT
PxylA-dnaN vabA-null yab.4-null yabA-null Pipt.-dnaB371
PxvylA-dna1N Piptg-dnaB371
WT
yabA-null
PxylA-dnaN
Figure 1
183
Figure 1. Effects of P-clamp on the initiation of DNA replication are likely through
YabA. Indicated strains were grown in minimal medium in the presence of an inducer for
at least 4 generations before exponentially growing cells were collected for analysis of
total DNA and protein. dnaNwas induced with xylose, dnaB3 71 and yabA were induced
with IPTG. Results normalized to WT. A) YabA inhibits DNA replication in a recAindependent manner. B) YabA inhibits DNA replication in an oriC-dependentmanner. C)
dnaN overexpression and yabA deletion have no additive effects, though yabA deletion
and dnaB371 do have additive effects. D) Effects of YabA and n--clamp are diminished
under fast growth conditions. Strains were grown in rich medium (LB) at 370 C until midexponential growth phase and were collected for DNA/protein ratio analysis.
184
additive effect, and indeed, the yabA-null, PxylA-dnaN strain overreplicated only as
much as the yabA-null alone, which has the more severe phenotype (Fig. 1C). To
demonstrate that the potential of the cells to initiate replication is not saturated and that
higher DNA to protein ratios can be observed, we made a yabA-null, Piptg-dnaB371
double mutant which had an additive effect (Fig. 1C). Our results strongly indicate that
YabA and P-clamp are on the same regulatory pathway. Our working model is that YabA
inhibits DnaA activity and P-clamp inhibits YabA.
It has been previously shown that overexpression of dnaA-dnaN operon in B.
subtilis increases the rate of initiation of DNA replication (48).To test if the effects were
due to overproduction of DnaA alone or whether DnaN contributed to the observed
overreplication, we wanted to test if the effects of P-clamp and DnaA overexpression
were additive. We observed that overexpressing DnaA in addition to P-clamp caused
more overreplication than overexpression of either DnaA or P-clamp alone (Fig. 2).
Overexpressing DnaA and P-clamp in yabA-null cells also increased the DNA to protein
ratio as compared to yabA-null cells overexpressing P-clamp (data not shown). These
results strongly indicate that DnaA regulates replication by a parallel or down-stream
pathway in relation to P-clamp and YabA. These data agree with the expectation that
DnaA is limiting for origin melting, and that the initiation of DNA replication is
regulated in part by controlling the level of DnaA (42). Our observations indicate that the
initiation of DNA replication is regulated by multiple pathways in B. subtilis.
DnaA and DnaA-dependent replication are not required for YabA focus
formation. Monitoring GFP-YabA localization in the cells has revealed that GFP-YabA
forms foci and that the intracellular distribution of the foci is similar to that of the
185
..
...
2.5
... ....
....... ..........- -................. .................
...........
-- ~~-I-
2.0
Piptg-dnaA·_·
PxyIA-dnaN
WT Piptg-dnaA
1.5
1.0
0.5 -
0-
!
I
WT
Piptg-dnaA
-I
PxylA-dnaN
Piptg-dnaA
PxylA-dnaN
Figure 2
186
Figure 2. DnaA and f3-clamp overexpression have additive effects on DNA
replication. Indicated strains were grown in minimal medium in the presence of an
inducer for at least 4 generations before exponentially growing cells were collected for
analysis of total DNA and protein. dnaN was induced with xylose, dnaA was induced
with IPTG. Results normalized to WT.
187
replisome (18, 45). Based on YabA mutational analysis, it was concluded that YabA
needs interact with both DnaA and P-clamp in order to form foci and regulate replication
(45).
To gain further insight into the role of DnaA in YabA localization, we investigated
whether DnaA or replication from the DnaA-dependent origin of replication, oriC,are
required for the formation of GFP-YabA foci. We expressed a GFP-YabA fusion protein
in an oriN+ AoriC-S mutant. We also inactivated dnaA in oriN+AoriC-S cells by
integrating a plasmid to disrupt dnaA and place dnaN under the control of the IPTG
inducible promoter Pspac-(hy). We verified the inactivation of DnaA by transcriptional
analysis and Western blotting (data not shown). GFP-YabA still formed foci in the oriN+
JoriC-Scells dnaA-inactivated strains (Fig. 3G, 31). These results are inconsistent with
the notion that formation of GFP-YabA focireqires DnaA (45). These results indicate that
neither DnaA nor initiation from oriC are required for GFP-YabA focus formation.
YabA likely recruited to the replisome by f-clamp. We addressed whether p-clamp is responsible for the localization pattern of YabA. Since n-clamp is part of the
replisome, P-clamp-GFP forms foci with intracellular distribution similar to that of other
replisome components (37). In cells treated with the DNA polymerase inhibitor HPUra,
foci of many replisome components, including DNA polymerase (PolC), clamp loader
HolB subunit, and clamp loader/r-subunit (DnaX), persist for more than 60 min after
replication arrest (Fig. 3C, 3D, and data not shown). On the other hand, P-clamp foci
disappear completely within 10 min of HPUra addition (Fig. 3A, 3B). We speculate that
P-clamp is largely dissociated from the replisome after replication arrest. If n-clamp
were recruiting YabA to the replisome, then GFP-YabA foci should also disappear after
188
A.
+ HPUra
No HPUra
B.
c.
D.
E.
F.
G.
H.
I.
J.
Figure 3
189
Figure 3. GFP-YabA focus formation does not depend on DnaA or DnaAdependent replication and is disturbed replication arrest. Exponentially growing cells
expressing various GFP fusions were treated with HPUra and prepared for microscopy
within 10 min of treatment (B, D, F, H, J) in parallel with untreated cells expressing the
same GFP fusion (A, C, E, G, I). Cells were grown in minimal medium supplemented
with glucose, or with arabinose and xylose in the case of strains expressing GFP-YabA.
A-B) P-clamp-GFP (AIG371), C-D) DnaX-GFP (KPL 382), E-F) GFP-YabA (AIG470),
G-H) GFP-YabA in oriC-S, dnaA-null cells (AIG593), I-J) GFP-YabA in oriC-S,dnaA
overexpresing cells (AIG595).
190
HPUra treatment. Indeed in WT, oriN+ AoriC-S, and oriN+ AoriC-S dnaA-null cells
HPUra caused YabA foci to disappear from virtually all cells within 15 min of treatment
(Fig 3E-H, and data not shown). Our data is consistent with the notion that P-clamp
recruits YabA to the replisome.
P-clamp and YabA do not affect the expression of DnaA regulated genes. The
expression of DnaA regulated genes is sensitive to the levels of DnaA (48). If YabA and
P-clamp regulated the availability of DnaA for replication by sequestering it at the
replisome (45), then altering this regulatory pathway should also affect the expression of
DnaA regulated genes as the free levels of DnaA would be changed. To investigate this
hypothesis we deleted yabA, or overexpressed YabA or p-clamp and monitored the
expression of DnaA regulated genes using microarrays. Our data indicated that
overexpressing YabA or P--clamp for 60 min did not affect any of the known DnaA
targets (Fig 4). Furthermore, deleting yabA did not affect the expression of DnaA
regulated genes with the exception of a single operon, the pyr operon (Fig. 4). The
regulation of this operon is rather complex (7, 56), and the change in pyr gene expression
is likely an indirect effect of replication in the yabA-null strain. Deletion ofyabA did
induce the expression of DNA damage responsive genes (data not shown), consistent
with the idea that excessive replication causes stalled forks and DNA damage (53).
We also tested whether YabA overexpression interferes with the ability of DnaA to
induce a transcriptional response after replication arrest. We performed these experiments
in a DnaBts mutant which allowed us to arrest either the elongation of DNA replication
with the drug HPUra (14), or the initiation of DNA replication by shifting to a nonpermissive temperature in the same strain. When yabA was not overexpressed, both
191
dnaA-dnaN
Iron res pons ive
genes;
Fur regulon
r"B-ylxA-ftsL -pbpB
Pyrimidine
biosynthesis
ymaA-nrdE -nrdFymaB
Amino acid
bios ynthes is
sda
Uncknown
function
WT
yabA-null
HPUra
HPUra
yabA-null
yabAT60
yabA T90
dnaN T60
Figure 4.
192
Figure 4. yabA and dnaN do not affect the expression of DnaA-regulated genes. The
relative mRNA levels in cells treated with HPUra, or overexpressing dnaN or yabA with
respect to untreated cells or wild type were determined using microarrays. Samples of
parallel treated or untreated cell cultures, or mutant and wild type, were taken at various
times for a direct comparison. Samples were taken during mid-exponential growth and
when treated with HPUra, data are shown for samples taken 60 min after treatment. The
average from three biological replicates is presented.
Data are presented as colored boxes, with each box representing a gene. The
brightest green represents a 2 4-fold decrease and the brightest red represents a 2 4.0-fold
increase in relative levels of mRNA. Gray cells indicate no data for that particular gene
in the particular experiment. Black indicates no change. Relevant strain properties and
treatments are indicated. Results are shown for the 56 genes that were previously
proposed as DnaA regulated (14). Genes are ordered according to their position in the
chromosome.
Column WT HPUra; wild type strain (JH642) treated with HPUra compared to the
same strain untreated. column vabA-null HPUra; yabA-null strain (AIG109) treated
with HPUra compared to the same strain untreated. columns vabA T60 and T90; yabA
overexpressing strain (AIG80) 60min or 90 min after addition of inducer compared to the
same stain uninduced. column dnaN T60; dnaN overexpressing strain (MMB 26)
60min after addition of inducer compared to the same stain uninduced.
193
HPUra and temperature shift affected most of the operons previously proposed to be
DnaA regulated (Chapter 2) (Fig. 4). A comparison between DnaBts cultures grown at
non-permissive temperature (inhibited replication initiation) either overexpressing yabA
or not, revealed that all DnaA regulated genes were affected to a similar extent in yabA+
and yabA overexpressing cultures (Fig. 4). Similar results were observed in HPUra
treated cultures, where virtually all DnaA regulated genes were affected similarly in
yabA+ and yabA overexpressing cultures (Fig. 4). Two operons (dnaAN and yclNOP)
were repressed to a lesser extent (-2 fold less) in yabA overexpressing cultures (Fig. 3).
These data suggest that YabA and P-clamp do not affect the expression of the majority of
DnaA regulated genes.
Discussion
To understand how the initiation of DNA replication is regulated in the bacterium
B. subtilis, we investigated the effects of YabA and P-clamp on initiation and their
effects on gene expression. Our results indicate that YabA and 3-clamp act in the same
pathway that regulates initiation. We show that the ability of YabA to form foci is
independent of DnaA and DnaA-dependent replication, but likely depends on P-clamp.
On the other hand, we showed that the ability of YabA to regulate replication requires
DnaA-dependent initiation. We also demonstrate that YabA and fr-clamp do not affect
the expression of DnaA regulated genes. Our data support a model in which YabA
negatively regulates DnaA at the origin of replication, and 3-clamp is a negative
regulator of YabA.
Regulation of the initiation of DNA replication by YabA and P-clamp. Our
experiments indicate that the regulation of DNA replication by P-clamp (DnaN) and
194
YabA requires the DnaA-dependent origin of replication, oriC. P-clamp appears to
activate a step after DnaA association with the origin, but before or at the time of helicase
loader (DnaB) association with the origin of replication (Chapter 4). We speculate that
the interaction of DnaA with other proteins at the origin, such as itself or DnaD, may be
the point of regulation by YabA and P-clamp.
YabA has been previously demonstrated to localize within the cell in a manner
similar to that of the replication machinery through its interactions with P-clamp and
DnaA. This observation led to the proposition that YabA functions with P-clamp to
sequester DnaA at the replication forks with the replisome, removing DnaA away from
the origin (45). In this case we would predict that replication would respond similarly to
changes in P-clamp or YabA expression, i.e. P-clamp and YabA should have negative
effects on replication. We demonstrate that unlike YabA, P-clamp is a positive regulator
of replication (Fig. 2) (Chapter 4). It is formally possible that the overexpression of dnaN
removes of YabA from the replisome thus increasing the levels of DnaA that would be
available to bind to the origin and regulate replication. However we have previously
shown that P-clamp (DnaN) overexpression did not affect the binding of DnaA to the
origin (Chapter 4). The effects of YabA in B. subtilis bear some resemblance to a
regulatory pathway in E. coli, in which the DnaA interacting protein, Hda, regulates the
initiation of DNA replication. Hda regulates the ratio of ATP to ADP bound DnaA
thereby affecting both replication initiation and gene expression in E. coli(1 3, 39, 43).
However YabA and Hda share no homology, and furthermore, deletion or overexpression
of yabA does not affect the expression of DnaA regulated genes in B. subtilis. These
observations imply that YabA is likely not functioning analogously to Hda. We also show
195
that overexpression P-clamp do not affect the expression of DnaA regulated genes (Fig.
3). We favor the hypothesis that YabA and P-clamp function to specifically regulate
DnaA activity at the origin of replication.
Role of YabA focus formation. GFP-YabA molecules form foci that appear to colocalize with the replisome. Previous observations suggested that YabA focus formation
is intricately related to the ability of YabA to regulate replication. Virtually all reported
mutants of YabA that had decreased ability to regulate replication initiation also had
decreased ability to form foci, including mutants that still interacted with either 0-clamp
(YabA-Aim) or DnaA (YabA-Nim) by yeast-two-hybrid assay (45). These results led to
the hypothesis that YabA has to interact with both DnaA and 3-clamp to form foci.
Although P-clamp is required for YabA focus formation, we found that DnaA is not
required. This discrepancy may stem from the inability of the YabA-Nim mutants to
interact with P-clamp in vivo, despite their ability to interact in a yeast-two-hybrid assay.
We speculate that the foci formed by YabA are a manifestation of the ability of YabA to
interact with P-clamp. However, since YabA negatively regulates the initiation of DNA
replication at oriC,the location of YabA foci may not be relevant to the regulatory
function of YabA.
One precedent for a protein that forms foci at the replisome but regulates
replication at the origin of replication is the SeqA protein from E.coli. SeqA forms foci
that co-localize with the replisome due to the interactions of SeqA with hemi-methylated
DNA generated during replication, but functions at the origin of replication to bind and
occlude it from access by replication proteins for parts of the cell-cycle (9).
196
Our data are consistent with a model in which YabA inhibits DnaA at the origin of
replication. When P-clamp associates with the origin, which happens before helicase is
loaded (Chapter 4), it inactivates YabA, allowing DnaA to stimulate helicase loading
likely through interacting with DnaD. Next, after P-clamp and the replisome replicate the
origin and move away, YabA is able to inhibit DnaA again and prevent additional
replication cycles. More work will be needed to verify this model.
Multiple layers of regulation of the initiation of DNA replication. The initiation
of replication in bacteria is coordinated with growth rate and responds to environmental
stimuli. In addition, when multiple replication origins are present in a single bacterial
cell, the initiation of replication occurs synchronously. This complex coordination of
replication initiation is achieved in B. subtilis and E. coli by multiple layers of regulatory
mechanisms. DnaA is the key regulator of replication in both bacterial species, and the
activity of DnaA is highly regulated (38, 48). Here we analyzed one regulatory
mechanism involving P-clamp and YabA that appears to affect DnaA activity
specifically at the origin of replication in B. subtilis. DnaA is also likely regulated by
other mechanisms, as overexpression of DnaA and P-clamp revealed additive effects
(Fig. 2). Replication in B. subtilis is also affected by the proteins required for helicase
loading, DnaB and DnaD (50). DnaD and DnaB are thought to interact only transiently
during the initiation of DNA replication, and expression of mutants that interact
constitutively, such as dnaB371, affects replication initiation (50). Since the effects of
dnaB371 are additive with those of yabA deletion (Fig. 3C), and with DnaA
overexpression, modulation of DnaB and DnaD interactions may constitute yet another
mechanism to regulate replication initiation(50). The activity of the ParA and ParB
197
homologues, Soj and SpoOJ respectively, which act in chromosome partitioning also
affect replication in B. subtilis, although the mechanism is not known (31, 32). It will be a
major task in the future to understand the interdependence of these regulatory
mechanisms and how they modulate replication in response to internal and external
stimuli.
Acknowledgements
I wish to thank T. Baker, S. P. Bell, F. Solomon, A. Wright, C. Lee, J. Wang, and J.
Auchtung for suggestions and comments on the manuscript.
198
References
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
Atlung, T., E. S. Clausen, and F. G. Hansen. 1985. Autoregulation of the dnaA
gene of Escherichia coli K12. Mol Gen Genet 200:442-50.
Au, N., E. Kuester-Schoeck, V. Mandava, L. E. Bothwell, S. P. Canny, K.
Chachu, S. A. Colavito, S. N. Fuller, E. S. Groban, L. A. Hensley, T. C.
O'Brien, A. Shah, J. T. Tierney, L. L. Tomm, T. M. O'Gara, A. I. Goranov,
A. D. Grossman, and C. M. Lovett. 2005. Genetic composition of the Bacillus
subtilis SOS system. J Bacteriol 187:7655-66.
Autret, S., A. Levine, F. Vannier, Y. Fujita, and S. J. Seror. 1999. The
replication checkpoint control in Bacillus subtilis: identification of a novel RTPbinding sequence essential for the replication fork arrest after induction of the
stringent response. Mol Microbiol 31:1665-79.
Baker, T. A., and S. P. Bell. 1998. Polymerases and the replisome: machines
within machines. Cell 92:295-305.
Braun, R. E., K. O'Day, and A. Wright. 1985. Autoregulation of the DNA
replication gene dnaA in E. coli K-12. Cell 40:159-69.
Britton, R. A., P. Eichenberger, J. E. Gonzalez-Pastor, P. Fawcett, R.
Monson, R. Losick, and A. D. Grossman. 2002. Genome-wide analysis of the
stationary-phase sigma factor (sigma-H) regulon of Bacillus subtilis. J Bacteriol
184:4881-90.
Chander, P., K. M. Halbig, J. K. Miller, C. J. Fields, H. K. Bonner, G. K.
Grabner, R. L. Switzer, and J. L. Smith. 2005. Structure of the nucleotide
complex of PyrR, the pyr attenuation protein from Bacillus caldolyticus, suggests
dual regulation by pyrimidine and purine nucleotides. J Bacteriol 187:1773-82.
Collier, J., S. R. Murray, and L. Shapiro. 2006. DnaA couples DNA replication
and the expression of two cell cycle master regulators. Embo J 25:346-56.
Donachie, W. D. 2001. Co-ordinate regulation of the Escherichia coli cell cycle
or The cloud of unknowing. Mol Microbiol 40:779-85.
Donachie, W. D., and G. W. Blakely. 2003. Coupling the initiation of
chromosome replication to cell size in Escherichia coli. Curr Opin Microbiol
6:146-50.
Felczak, M. M., and J. M. Kaguni. 2004. The box VII motif of Escherichia coli
DnaA protein is required for DnaA oligomerization at the E. coli replication
origin. J Biol Chem 279:51156-62.
Gautam, A., and D. Bastia. 2001. A replication terminus located at or near a
replication checkpoint of Bacillus subtilis functions independently of stringent
control. J Biol Chem 276:8771-7.
Gon, S., J. E. Camara, H. K. Klungsoyr, E. Crooke, K. Skarstad, and J.
Beckwith. 2006. A novel regulatory mechanism couples deoxyribonucleotide
synthesis and DNA replication in Escherichia coli. Embo J 25:1137-47.
Goranov, A. I., L. Katz, A. M. Breier, C. B. Burge, and A. D. Grossman.
2005. A transcriptional response to replication status mediated by the conserved
bacterial replication protein DnaA. Proc Natl Acad Sci U S A 102:12932-7.
199
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
Gueiros-Filho, F. J., and R. Losick. 2002. A widely conserved bacterial cell
division protein that promotes assembly of the tubulin-like protein FtsZ. Genes
Dev 16:2544-56.
Hartwell, L. H., and M. W. Unger. 1977. Unequal division in Saccharomyces
cerevisiae and its implications for the control of cell division. J Cell Biol 75:42235.
Harwood, C. R., and S. M. Cutting. 1990. Molecular Biological Methods for
Bacillus.John Wiley & Sons, Chichester, England.
Hayashi, M., Y. Ogura, E. J. Harry, N. Ogasawara, and S. Moriya. 2005.
Bacillus subtilis YabA is involved in determining the timing and synchrony of
replication initiation. FEMS Microbiol Lett 247:73-9.
Herrick, J., M. Kohiyama, T. Atlung, and F. G. Hansen. 1996. The initiation
mess? Mol Microbiol 19:659-66.
Ishigo-Oka, D., N. Ogasawara, and S. Moriya. 2001. DnaD protein of Bacillus
subtilis interacts with DnaA, the initiator protein of replication. J Bacteriol
183:2148-50.
Jaacks, K. J., J. Healy, R. Losick, and A. D. Grossman. 1989. Identification
and characterization of genes controlled by the sporulation-regulatory gene spo0H
in Bacillus subtilis. J Bacteriol 171:4121-9.
Johnson, A., and M. O'Donnell. 2005. Cellular DNA replicases: components
and dynamics at the replication fork. Annu Rev Biochem 74:283-315.
Kadoya, R., A. K. Hassan, Y. Kasahara, N. Ogasawara, and S. Moriya. 2002.
Two separate DNA sequences within oriC participate in accurate chromosome
segregation in Bacillus subtilis. Mol Microbiol 45:73-87.
Katayama, T., T. Kubota, K. Kurokawa, E. Crooke, and K. Sekimizu. 1998.
The initiator function of DnaA protein is negatively regulated by the sliding
clamp of the E. coli chromosomal replicase. Cell 94:61-71.
Kato, J., and T. Katayama. 2001. Hda, a novel DnaA-related protein, regulates
the replication cycle in Escherichia coli. Embo J 20:4253-62.
Kawakami, H., M. Su'etsugu, and T. Katayama. 2006. An isolated Hda-clamp
complex is functional in the regulatory inactivation of DnaA and DNA
replication. J Struct Biol.
Kogoma, T. 1997. Stable DNA replication: interplay between DNA replication,
homologous recombination, and transcription. Microbiol Mol Biol Rev 61:21238.
Kornberg, A., and T. A. Baker. 1992. DNA Replication, Second ed. W.H.
Freeman and Company, New York.
Kunst, F., N. Ogasawara, I. Moszer, A. M. Albertini, G. Alloni, V. Azevedo,
M. G. Bertero, P. Bessieres, A. Bolotin, S. Borchert, R. Borriss, L. Boursier,
A. Brans, M. Braun, S. C. Brignell, S. Bron, S. Brouillet, C. V. Bruschi, B.
Caldwell, V. Capuano, N. M. Carter, S. K. Choi, J. J. Codani, I. F.
Connerton, A. Danchin, and et al. 1997. The complete genome sequence of the
gram-positive bacterium Bacillus subtilis. Nature 390:249-56.
Kurokawa, K., S. Nishida, A. Emoto, K. Sekimizu, and T. Katayama. 1999.
Replication cycle-coordinated change of the adenine nucleotide-bound forms of
DnaA protein in Escherichia coli. Embo J 18:6642-52.
200
31.
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
42.
43.
44.
Lee, P. S., and A. D. Grossman. 2006. The chromosome partitioning proteins
Soj (ParA) and SpoOJ (ParB) contribute to accurate chromosome partitioning,
separation of replicated sister origins, and regulation of replication initiation in
Bacillus subtilis. Mol Microbiol 60:853-69.
Lee, P. S., D. C. Lin, S. Moriya, and A. D. Grossman. 2003. Effects of the
chromosome partitioning protein SpoOJ (ParB) on oriC positioning and
replication initiation in Bacillus subtilis. J Bacteriol 185:1326-37.
Lemon, K. P., I. Kurtser, and A. D. Grossman. 2001. Effects of replication
termination mutants on chromosome partitioning in Bacillus subtilis. Proc Natl
Acad Sci U S A 98:212-7.
Levine, A., S. Autret, and S. J. Seror. 1995. A checkpoint involving RTP, the
replication terminator protein, arrests replication downstream of the origin during
the Stringent Response in Bacillus subtilis. Mol Microbiol 15:287-95.
Lobner-Olesen, A., K. Skarstad, F. G. Hansen, K. von Meyenburg, and E.
Boye. 1989. The DnaA protein determines the initiation mass of Escherichia coli
K-12. Cell 57:881-9.
Marszalek, J., and J. M. Kaguni. 1994. DnaA protein directs the binding of
DnaB protein in initiation of DNA replication in Escherichia coli. J Biol Chem
269:4883-90.
Meile, J. C., L. J. Wu, S. D. Ehrlich, J. Errington, and P. Noirot. 2006.
Systematic localisation of proteins fused to the green fluorescent protein in
Bacillus subtilis: identification of new proteins at the DNA replication factory.
Proteomics 6:2135-46.
Messer, W. 2002. The bacterial replication initiator DnaA. DnaA and oriC, the
bacterial mode to initiate DNA replication. FEMS Microbiol Rev 26:355-74.
Messer, W., F. Blaesing, D. Jakimowicz, M. Krause, J. Majka, J. Nardmann,
S. Schaper, H. Seitz, C. Speck, C. Weigel, G. Wegrzyn, M. Welzeck, and J.
Zakrzewska-Czerwinska. 2001. Bacterial replication initiator DnaA. Rules for
DnaA binding and roles of DnaA in origin unwinding and helicase loading.
Biochimie 83:5-12.
Messer, W., and C. Weigel. 1997. DnaA initiator--also a transcription factor.
Mol Microbiol 24:1-6.
Morigen, E. Boye, K. Skarstad, and A. Lobner-Olesen. 2001. Regulation of
chromosomal replication by DnaA protein availability in Escherichia coli: effects
of the datA region. Biochim Biophys Acta 1521:73-80.
Moriya, S., Y. Imai, A. K. Hassan, and N. Ogasawara. 1999. Regulation of
initiation of Bacillus subtilis chromosome replication. Plasmid 41:17-29.
Nishida, S., K. Fujimitsu, K. Sekimizu, T. Ohmura, T. Ueda, and T.
Katayama. 2002. A nucleotide switch in the Escherichia coli DnaA protein
initiates chromosomal replication: evidnece from a mutant DnaA protein
defective in regulatory ATP hydrolysis in vitro and in vivo. J Biol Chem
277:14986-95.
Noirot-Gros, M. F., E. Dervyn, L. J. Wu, P. Mervelet, J. Errington, S. D.
Ehrlich, and P. Noirot. 2002. An expanded view of bacterial DNA replication.
Proc Natl Acad Sci U S A 99:8342-7.
201
45.
46.
47.
48.
49.
50.
51.
52.
53.
54.
55.
56.
57.
58.
59.
Noirot-Gros, M. F., M. Velten, M. Yoshimura, S. McGovern, T. Morimoto, S.
D. Ehrlich, N. Ogasawara, P. Polard, and P. Noirot. 2006. Functional
dissection of YabA, a negative regulator of DNA replication initiation in Bacillus
subtilis. Proc Natl Acad Sci U S A 103:2368-73.
Norris, V., J. Fralick, and A. Danchin. 2000. A SeqA hyperstructure and its
interactions direct the replication and sequestration of DNA. Mol Microbiol
37:696-702.
Ogawa, T., Y. Yamada, T. Kuroda, T. Kishi, and S. Moriya. 2002. The datA
locus predominantly contributes to the initiator titration mechanism in the control
of replication initiation in Escherichia coli. Mol Microbiol 44:1367-75.
Ogura, Y., Y. Imai, N. Ogasawara, and S. Moriya. 2001. Autoregulation of the
dnaA-dnaN operon and effects of DnaA protein levels on replication initiation in
Bacillus subtilis. J Bacteriol 183:3833-41.
Perego, M., G. B. Spiegelman, and J. A. Hoch. 1988. Structure of the gene for
the transition state regulator, abrB: regulator synthesis is controlled by the spo0A
sporulation gene in Bacillus subtilis. Mol Microbiol 2:689-99.
Rokop, M. E., J. M. Auchtung, and A. D. Grossman. 2004. Control of DNA
replication initiation by recruitment of an essential initiation protein to the
membrane of Bacillus subtilis. Mol Microbiol 52:1757-67.
Seitz, H., C. Weigel, and W. Messer. 2000. The interaction domains of the
DnaA and DnaB replication proteins of Escherichia coli. Mol Microbiol 37:12709.
Sekimizu, K., D. Bramhill, and A. Kornberg. 1987. ATP activates dnaA protein
in initiating replication of plasmids bearing the origin of the E. coli chromosome.
Cell 50:259-65.
Simmons, L. A., A. M. Breier, N. R. Cozzarelli, and J. M. Kaguni. 2004.
Hyperinitiation of DNA replication in Escherichia coli leads to replication fork
collapse and inviability. Mol Microbiol 51:349-58.
Su'etsugu, M., T. R. Shimuta, T. Ishida, H. Kawakami, and T. Katayama.
2005. Protein associations in DnaA-ATP hydrolysis mediated by the Hdareplicase clamp complex. J Biol Chem 280:6528-36.
Sutton, M. D., K. M. Carr, M. Vicente, and J. M. Kaguni. 1998. Escherichia
coli DnaA protein. The N-terminal domain and loading of DnaB helicase at the E.
coli chromosomal origin. J Biol Chem 273:34255-62.
Switzer, R. L., R. J. Turner, and Y. Lu. 1999. Regulation of the Bacillus
subtilis pyrimidine biosynthetic operon by transcriptional attenuation: control of
gene expression by an mRNA-binding protein. Prog Nucleic Acid Res Mol Biol
62:329-67.
Tusher, V. G., R. Tibshirani, and G. Chu. 2001. Significance analysis of
microarrays applied to the ionizing radiation response. Proc Natl Acad Sci U S A
98:5116-21.
Wach, A. 1996. PCR-synthesis of marker cassettes with long flanking homology
regions for gene disruptions in S. cerevisiae. Yeast 12:259-65.
Zyskind, J. W., and D. W. Smith. 1992. DNA replication, the bacterial cell
cycle, and cell growth. Cell 69:5-8.
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Chapter 6
Discussion
203
My research has focused on understanding how the initiation of DNA replication is
regulated, and how cells respond to perturbations in replication. I investigated these
questions in the bacterium B. subtilis. I found that a component of the DNA polymerase
holoenzyme, P-clamp, regulates the initiation of DNA replication. The regulation by Pclamp is likely mediated by the P-clamp interacting protein YabA. This pathway
regulates a step before helicase loading. Interestingly, I also observed that P-clamp
arrives at the origin of replication before helicase, thus its association with the origin is
likely playing a role in the regulation of helicase loading.
My investigation of the responses to DNA damage and replication arrest showed
that most of the transcriptional response is mediated by the recombination protein, RecA,
but that also there is a substantial recA-independent response. Part of the recAindependent response is mediated by the replication initiation protein DnaA. I showed
that at lest one of the DnaA-regulated genes,ftsL, affects cell viability after replication
arrest by coordinating DNA replication and cell-division. I discuss these findings as well
as possible future directions in more detail below.
Regulation of replication by P-clamp and YabA. My work adds support to the
idea that the initiation of DNA replication is regulated at many steps in B. subtilis. One of
the first hints for multiple layers of regulation was the observation that DnaA levels peak
-15min before replication initiates (34), thus indicating that threshold levels of DnaA are
insufficient to initiate replication in B. subtilis, unlike what is currently hypothesized for
E. coli (8, 48). It is possible that initiation cannot occur until DnaA molecules at the
origin are rearranged by protein chaperones, as is the case with some plasmid replication
proteins, and/or steps after DnaA binding to the origin of replication are regulated. The
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work of Megan Rokop uncovered evidence for regulation of the initiation of DNA
replication after DnaA binding and origin melting. She demonstrated that helicase loader
proteins, which associate with the origin after it has been melted, only interact transiently,
and mutant proteins that interact constitutively alter the frequency of DNA replication
(44).
I demonstrated that overexpressing P-clamp (DnaN) stimulates the initiation of
DNA replication likely through inactivating the replication inhibitor YabA (Chapters 4
and 5). The overexpression of P-clamp did not affect the binding of DnaA to the origin
but affected the association of helicase loader and helicase with the origin (Chapter 4).
These results suggest that P-clamp affects a step after DnaA binding to the origin and
before helicase loader association with the origin. Since the target of the P-clamp and
YabA regulation is likely DnaA, either origin melting by DnaA, DnaA interactions with
helicase loader (e.g. DnaD), or some other DnaA-dependent process is affected. More
work is necessary to distinguish between these possibilities, although I favor the
hypothesis that YabA affects the ability of DnaA to interact with DnaD and/or other
helicase loading components, and that P-clamp, once associated with the origin, removes
the inhibition by YabA.
Although I favor the hypothesis that YabA and P-clamp affect replication
specifically at the origin, their proposed involvement in affecting the levels of free DnaA
(37) or the DnaA-ATP/ DnaA-ADP ratio (36) is not ruled out. I have shown that deletion
of YabA and overexpression of YabA or P-clamp do not affect the transcription of genes
regulated by DnaA (Chapter 5), suggesting that the levels of free DnaA are not grossly
perturbed under these conditions. However, minor effects of YabA and P-clamp on DnaA
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levels, which conceivably could regulate replication without affecting transcription, have
not been excluded. In B. subtilis, it is unclear whether gene expression and replication are
equally sensitive to changes in DnaA levels. Also, in B. subtilis, it has not been
determined whether the nucleotide bound state of DnaA plays a role in the ability of
DnaA to regulate gene expression. I have generated many useful constructs that will aid
in the further investigation of these unknown parameters in B. subtilis.
The mechanism by which P-clamp inactivates YabA is not clear. I have performed
P-clamp depletion experiments that support the hypothesis that P-clamp is limiting for the
initiation of replication, besides being essential for replication elongation (Chapter 4). It
is plausible that under conditions where P-clamp is overexpressed, the excess P-clamp
sequesters YabA away from the origin to the cytoplasm. I speculate that during the
normal cell cycle, when P-clamp associates with the origin of replication it sequesters or
removes YabA away from DnaA, thus stimulating replication initiation. To test this
hypothesis, the association of YabA with the origin of replication needs to be carefully
investigated. My preliminary results show that YabA associates with the origin in a DnaA
dependent manner, but I have not been able to rule out the possibility that YabA is
brought to the origin by P-clamp, instead of, as predicted by my hypothesis, that YabA
associates with the origin of replication before P-clamp.
Deletion of yabA or overexpression of P-clamp causes over-initiation of DNA
replication (Chapter 4, 5) and, at least in the case of YabA, loss of synchrony (19). How
this regulatory pathway is linked to the cell cycle is not clear, but one possibility is that
the expected cycling of P-clamp levels during the cell cycle (38, 48) ties into this
regulation. Alternatively, the ability of the P-clamp and DNA polymerase to associate
206
with the origin after it has been melted by DnaA may be regulated by some yet
undiscovered mechanism that is regulated by the cell cycle.
Since the role of protein chaperones in regulating replication has been
demonstrated in the case of plasmids (26, 40), and a parallel has been drawn between
these plasmids systems and DnaA in B. subtilis (35), it might be interesting to directly
test the effects of the deletion/depletion of protein chaperons on DNA replication. Should
protein chaperones have an effect on DNA replication it will be interesting to understand
what aspects of replication they affect.
It will also be interesting to determine if other mutations which regulate the
initiation of DNA replication, such as deletions of soj and spoOJ(30, 31), affect
replication independently of YabA and P-clamp. I have generated strains that will allow
this to be tested. Results obtained by Wang, Berkmen, and Grossman suggest that when
oriC is moved from the 00 region on the chromosome to 2700, under slow growth
conditions, the origin initiates replication more often as compared to when oriC is at 00.
This may indicate that something specific to the 00 region inhibits replication. Since
SpoOJ binds to several loci on the chromosome near the 00 region, it is possible that
SpoOJ is that inhibitory factor. The overreplication phenotype of a spoOJ mutant agrees
with this hypothesis.
DNA polymerase holoenzyme association with the origin. My results
demonstrated that, in B. subtilis, primase and the DNA polymerase holoenzyme can
associate with the origin of replication before helicase (Chapter 4). These observations
are in contrast to in vitro observations with purified E. coli replication components (10,
27). There are several reasons that may explain this discrepancy. It is possible that the
207
events during replication initiation are different in B. subtilis and E. coli. Alternatively,
the in vitro system may be missing replication components that would allow DNA
polymerase association with the origin before helicase. Also, the in vitro E. coli system
utilizes a plasmid that carries the chromosomal origin of replication (oriC) (10). While
this plasmid based system has most of the characteristics of the chromosomally located
origin of replication, there are some differences, for example the activity of origin
proximal promoters is essential for replication in the plasmid system, but has only modest
effects on replication on the chromosome (5).
Another ill defined parameter of replication is how much of the chromosomal
origin of replication is melted during initiation. In vitro and in vivo studies with the
plasmid borne oriC in E. coli, have suggested that DnaA unwinds only about 30bp within
the origin region (10, 13, 28). Additional in vitro experiments have shown that for
primase to associate with the origin of replication, it requires at least -70bp to be
unwound by helicase (10). Thus it has been suggested that the size of the ssDNA exposed
at the origin is one of the limiting factors for association of replication components. It
needs to be emphasized that we still do not know how much of the chromosomal origin
region is melted by DnaA in vivo in either E. coli or B. subtilis. SSB ChIP-chip
experiments hybridizing the immuno-precipitated DNA to high density microarrays could
shed some light on the issue.
One of the cornerstones in the current hypotheses of replication initiation in
bacteria is that primase is brought to the origin through interacting with helicase (27), and
therefore helicase has to be at the origin first. However, primase also interacts with the ycomplex of the DNA polymerase holoenzyme, at least in E. coli (25). Whether this
208
interaction alone is sufficient to bring the primase to the replication fork is not known,
but it seems like an enticing possibility in light of my observations that primase, in B.
subtilis at least, associates with the origin of replication before helicase. It should be
mentioned that such interactions between primase and DNA polymerase holoenzyme in
B. subtilis have not yet been uncovered.
One of the most interesting questions posed by my observations is what recruits the
DNA polymerase holoenzyme to the origin in the absence of helicase there. To begin
addressing this issue, we need to be able to remove/ inactivate components of DNA
polymerase and then test the association of the other subunits with the origin. There are
temperature sensitive mutant alleles for most replication proteins (23). Unfortunately,
most of these mutants are very poorly characterized and for most of them the exact
biochemical function inactivated is not known, making experiments hard to interpret. A
more fruitful approach might be to generate unstable, rapidly degraded proteins, such that
accumulation of normal protein levels requires a high level of transcription. Such
unstable mutant alleles could be placed under repressible promoters, such that
transcription could be repressed leading to rapid net protein loss. Such mutants could be
synchronized for replication either through germinating spores, or through using a well
understood replication ts mutant such as DnaBts. These experiments could allow us to
test the role of primase, P-clamp, clamp loader, DNA polymerase, and any other
replication protein in the association of components of DNA polymerase holoenzyme
with the origin before helicase. It is likely that ssDNA at the melted origin and maybe
interactions with DnaA are the initial factors that recruit the DNA polymerase
holoenzyme to the origin, but a more detailed investigation of the replisome subunit
209
requirements for association will lead to a better understanding of replication initiation.
An alternative approach to ChIP in understanding events during the initiation of DNA
replication may be to express fragments of replication proteins, e.g. P-clamp, and ask is
they can perturb the regulation by YabA. Expression of P-clamp form that cannot be
loaded onto DNA but still interacts with YabA could help us understand how P-clamp
inactivated YabA. If P-clamp has to be loaded onto DNA to inactivate YabA, than
expressing a form that cannot be loaded onto DNA should not affect DNA replication,
and if it does it suggests that P-clamp can inactivate YabA by recruiting it away from the
origin of replication.
It may be interesting to also test the order of association of replication components
at a DnaA-independent, plasmid origin of replication, oriN (22), in vivo in B. subtilis.
These experiments may shed light on the requirement of DnaA for DNA polymerase
association, and how general the observations at oriC are.
Initiation of replication during spore outgrowth. For some of my experiments I
used germinating spores as a method for synchronizing DNA replication (17, 49). To
understand how the initiation of DNA replication under these conditions differs from
replication during exponential growth I read old literature concerning DNA replication
after spore germination. Observations more than 30 years ago suggested that replication
regulates protein synthesis during spore germination and outgrowth (14, 16, 42, 45).
Although the general consensus currently is that progression through the cell cycle does
not impinge on growth (18, 20, 21, 24), spore germination is a clear example where the
inability to progress through the cell cycle inhibits growth. These observations imply that
replication during spore outgrowth begins prior to renewed protein synthesis, and that all
210
necessary replication components are already present and ready for replication. Two
basic questions then arise. Is the replisome pre-loaded at the origin of replication during
the early steps of spore formation, or does the replisome load during germination of the
mature spore, when it encounters favorable growth conditions? How does replication
during early germination stimulate protein synthesis?
The first of these questions can be addressed by performing ChIP experiments with
germinating spores, to assay whether replisome components are associated with the
origin before or after germination. I performed preliminary ChIP experiments with
outgrowing spores and showed that after 90min of germination and outgrowth all tested
replication components were associated with the origin of replication. This result lays the
groundwork for more detailed analysis of the timing of association, during sporulation
and germination. If the replisome is loaded at the origin of replication during sporulation,
i.e. the replisome is at the origin in dormant spores, it would interesting to know how this
association is stably maintained for extended periods of time until the spore germinates.
If the replisome associates during spore germination, then it is interesting to understand
what signals the initiation of replication, and /or what inhibits it from happening while
the spores are dormant.
How replication stimulates protein synthesis may be explained by the ability of
replication to regulate gene expression (Chapter 2, 3). Although the changes in gene
expression during spore outgrowth have not been investigated, replication regulates the
expression of many genes during exponential growth (see below). It is possible that the
early replication during spore germination and outgrowth affects the expression of factors
that regulate global transcription and translation.
211
DnaA as a transcriptional regulator. My work implicated DnaA in the regulation
of more than 15 operons in addition to the two known DnaA targets in B. subtilis
(Chapter 2). The criteria I used to define the potential DnaA regulon was to find operons
with 2 or more putative DnaA binding sites in the promoter region that were affected by
conditions where the expression of the known DnaA operons was affected. I used ChIP to
demonstrate that DnaA bound to six of these promoters. The role of DnaA in regulating
the rest of the operons needs to be more rigorously investigated. I have generated several
strains that carry dnaA-null mutations, or that overexpress dnaA without affecting
replication. Such strains should prove useful in approaching the question of which of the
proposed DnaA targets are directly regulated by DnaA.
The genes regulated by DnaA were affected after inhibition of either initiation or
elongation of replication. Some genes were up-regulated and some were down-regulated
after replication arrest, indicating that DnaA could be either a transcriptional activator or
a repressor. One of the DnaA targets, sda, was previously demonstrated to couple
replication status and sporulation, such that if there were perturbations in replication,
sporulation was inhibited (7). DnaA induces the transcription of sda during replication
arrest, and Sda inhibited early steps of sporulation (7). I demonstrated that the regulation
of another DnaA target gene,ftsL, contributes to the inhibition of cell division after
replication arrest (Chapter 2). After replication arrest,ftsL expression is repressed, and
since FtsL is a very unstable protein (9), protein levels likely decrease thus inhibiting cell
division. Repression offtsL by replication arrest is important for cell survival. WhenftsL
is expressed constitutively after replication arrest, cells divide more often, and lose
212
viability (Chapter 2). This regulation was in addition to other known regulatory pathways
that coordinate replication and cell division.
Other proposed DnaA targets include the highly conserved, putative essential
translation factor ywlC, the GTP-binding protein yqeH, and highly conserved proteins of
unknown function such as yydA. It will be interesting to understand how the regulation of
these proteins helps the cells to survive perturbations in replication. A potentially very
exciting finding would be if it were demonstrated that by regulating YwlC, cells adjust
the translation rate of a large set of mRNAs, and thus adjust their metabolism to changing
conditions.
It is still not clear how DnaA senses the inhibition of DNA replication. My results
demonstrated that DnaA regulated genes are affected upon inhibition of either replication
elongation or initiation. In both cases, due to the manner of arrest, most components of
the replisome are associated with the chromosome, except that when initiation is arrested
helicase is not loaded. I infer that helicase is not needed for DnaA to detect inhibition of
replication. I hypothesize that active replication somehow inactivates DnaA, and when
replication is inhibited DnaA is able to affect gene expression. There are at least two
models for how replication could be inactivating DnaA. One possibility is that the
passing replication forks remove DnaA from the promoters that DnaA regulates, thus
temporarily alleviating the regulation by DnaA. Alternatively, replication could regulate
the transcriptional activity of DnaA by affecting the nucleotide binding state of DnaA, as
appears to be the case in E. coli. A crucial set of experiments will be to determine the
ATP-DnaA/ADP-DnaA ratios in growing B. subtilis cultures before and after replication
arrest. The occupancy by DnaA during the cell cycle and after replication arrest could be
213
monitored by using ChIP-chip. These two approaches should help to understand how
replication regulates DnaA transcriptional activity. If replication stimulates the ATPase
activity of DnaA, as in E. coli, then determining which replisome factors contribute to
this stimulation would be the next step. I have generated a DnaA mutant (Lys318 to Ala),
which should be unable to hydrolyze ATP to ADP. The same mutation in the E.coli
DnaA protein renders DnaA hyper active for replication initiation (29). My preliminary
analysis with a strain that expresses both wt and mutant DnaA show that replication is
inhibited, rather than stimulated. Microarray analysis of this strain also suggests that the
mutant DnaA protein is a more potent transcription factor. Further analysis of the DnaA
mutant will be necessary in order to understand how it affects replication and
transcription. Generating strains where the mutant version of DnaA is the only DnaA in
the cell, as well as strains that carry the mutant version of DnaA and replicate in a DnaAindependent manner should be a priority. Some biochemical characterization of the
mutant DnaA will also be necessary to show that other measurable functions of DnaA are
not affected by the mutation.
Since 13-clamp is implicated in regulating DnaA transcriptional activity in E. coli
(15, 39), 1performed some preliminary experiments testing the possible role of P-clamp
in B. subtilis. I used temperature sensitive mutants of 13-clamp to test whether the
mutations, which inhibit replication elongation, could prevent the induction of the DnaAmediated response. Unfortunately, my experiments revealed that inhibiting replication by
inactivating P-clamp with two different ts alleles (dnaN5 and dnaN34) still elicited a
robust effect on DnaA regulated genes. These observations do not disprove the
involvement of 3-clamp in regulating DnaA, as we do not understand how the ts alleles
214
inactivate the P-clamp protein. So far, in all my experiments where I inhibited DNA
replication directly (inhibiting helicase, 3-clamp, or DNA polymerase), I observed the
DnaA-mediated response. I have not tried inhibiting topoisomerase or gyrase, but I
suspect that the same observations would be made.
It should also be emphasized that not all types of replication perturbations induce
the DnaA mediated transcriptional response. My observations indicate that prolonged and
nearly complete inhibition of replication is required to generate detectable effects on key
DnaA targets (dnaA, dnaN, sda,ftsL, ywlC). Treatment of B. subtilis cells with the DNA
damaging agents Mitomycin C (MMC) or with UV light does not affect any of these
genes, in wt cells (Chapter 3). This is likely due to incomplete inhibition of replication or
transient inhibition, respectively. However, treatment of a strain incapable of repairing
most DNA damage lesions (recA-null) with UV light but not with MMC affected the
DnaA regulated targets. These observations suggest that unrepaired UV damage stalls
replication very efficiently, while MMC damage is still bypassed by replication forks
likely leaving daughter strand ssDNA regions. Indeed, my analysis shows that MMC
slows down without completely abolishing replication (Chapter 3).
It is worth noting that the genes regulated by DnaA in response to replication arrest
may also be regulated by DnaA during the normal cell cycle, such that their transcription
would be cyclical. The regulation of dnaA and nrdA in E. coli is cyclic (3, 48), and DnaA
is thought to couple the expression of these genes to the time of the cell cycle when they
are needed. For example, the nrdA gene (ribonucleotide reductase) is expressed in E. coli
during replication when deoxy-ribonucelotides are needed for replication (15, 46, 47).
One potential candidate of such a cell cycle regulated gene in B. subtilis is sda. This
215
protein inhibits entry into sporulation, and there is evidence that cells can enter into
sporulation only within the first one-half of the replication cycle, but not later (33). It
should be interesting to investigate if the inability of cells to enter sporulation later in the
cell cycle corresponds to increased sda expression. It may be worth identifying additional
DnaA and cell cycle regulated genes by monitoring the global transcript levels of genes
in synchronized cultures.
Other transcriptional responses to replication arrest and DNA damage. One of
the best characterized responses to DNA damage is the SOS response which is mediated
by the recombination protein RecA (11). In B. subtilis,E. coli, V cholerae and other
bacteria, in the presence of DNA damage, RecA induces the expression of genes
repressed by LexA, as well as the expression of phage and mobile element genes
[Chapter 3, (1, 2, 6, 11, 41)]. RecA induces gene expression by inactivating LexA and
phage repressors (11, 43). Since many genes repressed by LexA have been demonstrated
to have functions in DNA damage repair and recombination, it has been of great interest
to identify the complete set of LexA targets.
In collaboration with Elke Kuester-Schoeck and Jade Wang, I performed global
transcriptional analysis of cells treated with DNA damaging reagents or inhibited for
replication (Chapter 3). We also analyzed similarly treated recA and lexA mutants. This
analysis allowed me to define a set of over 60 genes as likely direct targets of LexA.
About 40 of those genes have been previously proposed to be regulated by LexA,
although for many of them, their biochemical function or role in coping with DNA
damage is not known. We also demonstrated that many of the transcriptional effects
caused by DNA damage were due to activation of phage and mobile elements and the
216
indirect effects of phage production. Interestingly, phage gene expression could not be
induced in cells that could not inactivate LexA after DNA damage. This suggested to us
that one or more LexA repressed genes is required for phage induction. We still do not
know what gene that is, but I can speculate that recA is a good candidate, as recA is
repressed by LexA and is induced after DNA damage. Alternatively, at least in the case
of PBSX phage, LexA represses a gene (xkdA), which is homologous to the anti-repressor
of ICEBsl (immA). This may suggest that inactivation of LexA induces an anti-repressor
that inhibits the PBSX phage repressor Xre. It must be emphasized that induction of xkdA
and all other lexA repressed genes is insufficient for inducing PBSX gene expression,
suggesting that DNA damage is needed in parallel to the expression of LexA repressed
genes. Understanding further how phage are regulated is one possible future direction of
my initial observation.
Our analysis revealed that in recA-null cells there is a substantial transcriptional
response of more than 100 genes after replication arrest (Chapter 2). The aforementioned
DnaA mediated response falls in that category. Although DnaA affects the expression of
its targets in recA-null cells, the magnitude of the response is diminished. This indicated
to me that recA or recA-regulated genes affect the ability of DnaA to affect gene
expression. Many of the recA/lexA-regulated genes affect events at the replication fork
and may modulate the generation of the signal(s) that regulate DnaA.
In a recA-null mutant, there are still more than 50 genes affected by replication
arrest that are not likely to be regulated by DnaA. These genes are also affected only by
inhibition of replication elongation but not by inhibition of initiation. I observed the
effects on these genes by inhibiting the DNA polymerase with a drug as well as by
217
inactivating helicase with a temperature sensitive mutation (Chapter 2). Most of them
have been demonstrated to respond to oxidative stress in the cell or the concentration of
iron in the media. Both sets of genes respond to the repressor PerR (4, 12). Preliminary
results have indicated that in a constitutively active PerR repressor, these genes are not
affected by replication elongation arrest. These observations indicate that replication
elongation arrest inactivates PerR causing the induction of PerR repressed genes, which
include the repressor Fur, which inhibits iron responsive genes. Since reactive oxygen
species and iron could cause severe DNA damage, the observed responses could be
rationalized by suggesting that the cells are preventing further DNA damage. It is still
unclear how replication arrest regulates PerR function, as PerR so far has only been
demonstrated to sense elevated oxidation levels in the cell. Unless it is demonstrated that
DNA damage causes elevated oxidation levels, this cellular response appears to be a
preventative measure.
I have shown that replication arrest can inhibit cell division in a recA-independent
manner (Chapter 2). A poorly characterized temperature sensitive mutant, tsi-23, was
isolated as an inducer of the recA-dependent SOS response, but can also induce a recAindependent cell division arrest (32). These observations are most easily explained by
suggesting that tsi-23 causes replication arrest. The tsi-23 mutation maps close to the air
gene (alanine-racemase) on the bacterial chromosome (32). It may be interesting to
identify the gene that carries the tsi-23 and understand how it induces the SOS response
and/or inhibits replication. Examination of the genes close to air locus, revealed a
potential candidate for a gene that carries the tsi-23 mutation: ydbR, a gene with unknown
218
function with high homology to ATP-dependent RNA helicases. More work will be
necessary to show whether ydbR is indeed the gene that carries the tsi-23 mutation.
Concluding remarks. My studies have generated the foundations for many future
experiments investigating the coordination of DNA replication and the cell cycle. If I
were to continue working on any of these projects, my emphasis would be on
understanding the significance and mechanism of the YabA and 3-clamp regulatory
pathway and how it is linked to the cell cycle. In my opinion this one of the most
fundamental biological questions that is still poorly understood. These regulatory
mechanisms likely act in trans, as under fast growing conditions, replication origins
located far from each other initiate replication simultaneously. It is still unclear how this
precision in timing is achieved.
I hope that my work will help to establish B. subtilis as a model organism for
understanding bacterial replication, and will stimulate an interest in approaching and
reevaluating the question of how of DNA replication is regulated. So far bacterial DNA
replication has been well understood only in E. coli. Since E. coli is unique in many
respects, it remains unclear how much of what is known about E. coli can be correctly
generalized for other bacteria and organisms. It will be interesting to see the outcome of
the projects and questions I discussed above, and I only hope that any future scientists
that work on these subjects find them as exciting as I do.
Acknowledgements
I wish to thank A. Grossman, C. Lee, T. Baker, S. Bell, F. Solomon, and A. Wright for
helpful suggestions and comments.
219
References
1.
2.
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
Au, N., E. Kuester-Schoeck, V. Mandava, L. E. Bothwell, S. P. Canny, K.
Chachu, S. A. Colavito, S. N. Fuller, E. S. Groban, L. A. Hensley, T. C.
O'Brien, A. Shah, J. T. Tierney, L. L. Tomm, T. M. O'Gara,A. I. Goranov,
A. D. Grossman, and C. M. Lovett. 2005. Genetic composition of the Bacillus
subtilis SOS system. J Bacteriol 187:7655-66.
Auchtung, J. M., C. A. Lee, R. E. Monson, A. P. Lehman, and A. D.
Grossman. 2005. Regulation of a Bacillus subtilis mobile genetic element by
intercellular signaling and the global DNA damage response. Proc Natl Acad Sci
U S A 102:12554-9.
Augustin, L. B., B. A. Jacobson, and J. A. Fuchs. 1994. Escherichia coli Fis
and DnaA proteins bind specifically to the nrd promoter region and affect
expression of an nrd-lac fusion. J Bacteriol 176:378-87.
Baichoo, N., T. Wang, R. Ye, and J. D. Helmann. 2002. Global analysis of the
Bacillus subtilis Fur regulon and the iron starvation stimulon. Mol Microbiol
45:1613-29.
Bates, D. B., E. Boye, T. Asai, and T. Kogoma. 1997. The absence of effect of
gid or mioC transcription on the initiation of chromosomal replication in
Escherichia coli. Proc Natl Acad Sci U S A 94:12497-502.
Beaber, J. W., B. Hochhut, and M. K. Waldor. 2004. SOS response promotes
horizontal dissemination of antibiotic resistance genes. Nature 427:72-4.
Burkholder, W. F., I. Kurtser, and A. D. Grossman. 2001. Replication
initiation proteins regulate a developmental checkpoint in Bacillus subtilis. Cell
104:269-79.
Campbell, J. L., and N. Kleckner. 1990. E. coli oriC and the dnaA gene
promoter are sequestered from dam methyltransferase following the passage of
the chromosomal replication fork. Cell 62:967-79.
Daniel, R. A., and J. Errington. 2000. Intrinsic instability of the essential cell
division protein FtsL of Bacillus subtilis and a role for DivIB protein in FtsL
turnover. Mol Microbiol 36:278-89.
Fang, L., M. J. Davey, and M. O'Donnell. 1999. Replisome assembly at oriC,
the replication origin of E. coli, reveals an explanation for initiation sites outside
an origin. Mol Cell 4:541-53.
Friedberg, E. C., Graham C. Walker, and Wolfram Siede. 1995. DNA Repair
and Mutagenesis. ASM Press.
Fuangthong, M., A. F. Herbig, N. Bsat, and J. D. Helmann. 2002. Regulation
of the Bacillus subtilis fur and perR genes by PerR: not all members of the PerR
regulon are peroxide inducible. J Bacteriol 184:3276-86.
Gille, H., and W. Messer. 1991. Localized DNA melting and structural
pertubations in the origin of replication, oriC, of Escherichia coli in vitro and in
vivo. Embo J 10:1579-84.
Ginsberg, D., and A. Keynan. 1978. Independence of Bacillus subtilis spore
outgrowth from DNA synthesis. J Bacteriol 136:111-6.
220
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
Gon, S., J. E. Camara, H. K. Klungsoyr, E. Crooke, K. Skarstad, and J.
Beckwith. 2006. A novel regulatory mechanism couples deoxyribonucleotide
synthesis and DNA replication in Escherichia coli. Embo J 25:1137-47.
Gottfried, M., C. Orrego, A. Keynan, and H. O. Halvorson. 1979. Specific
inhibition of outgrowth of Bacillus subtilis spores by novobiocin. J Bacteriol
138:314-9.
Harry, E. J., J. Rodwell, and R. G. Wake. 1999. Co-ordinating DNA replication
with cell division in bacteria: a link between the early stages of a round of
replication and mid-cell Z ring assembly. Mol Microbiol 33:33-40.
Hartwell, L. H., and M. W. Unger. 1977. Unequal division in Saccharomyces
cerevisiae and its implications for the control of cell division. J Cell Biol 75:42235.
Hayashi, M., Y. Ogura, E. J. Harry, N. Ogasawara, and S. Moriya. 2005.
Bacillus subtilis YabA is involved in determining the timing and synchrony of
replication initiation. FEMS Microbiol Lett 247:73-9.
Johnston, G. C., J. R. Pringle, and L. H. Hartwell. 1977. Coordination of
growth with cell division in the yeast Saccharomyces cerevisiae. Exp Cell Res
105:79-98.
Jorgensen, P., and M. Tyers. 2004. How cells coordinate growth and division.
Curr Biol 14:R1014-27.
Kadoya, R., A. K. Hassan, Y. Kasahara, N. Ogasawara, and S. Moriya. 2002.
Two separate DNA sequences within oriC participate in accurate chromosome
segregation in Bacillus subtilis. Mol Microbiol 45:73-87.
Karamata, D., and J. D. Gross. 1970. Isolation and genetic analysis of
temperature-sensitive mutants of B. subtilis defective in DNA synthesis. Mol Gen
Genet 108:277-87.
Kawai, Y., S. Moriya, and N. Ogasawara. 2003. Identification of a protein,
YneA, responsible for cell division suppression during the SOS response in
Bacillus subtilis. Mol Microbiol 47:1113-22.
Kelman, Z., A. Yuzhakov, J. Andjelkovic, and M. O'Donnell. 1998. Devoted
to the lagging strand-the subunit of DNA polymerase III holoenzyme contacts
SSB to promote processive elongation and sliding clamp assembly. Embo J
17:2436-49.
Konieczny, I., and D. R. Helinski. 1997. The replication initiation protein of the
broad-host-range plasmid RK2 is activated by the ClpX chaperone. Proc Natl
Acad Sci U S A 94:14378-82.
Kornberg, A., and T. A. Baker. 1992. DNA Replication, Second ed. W.H.
Freeman and Company, New York.
Krause, M., B. Ruckert, R. Lurz, and W. Messer. 1997. Complexes at the
replication origin of Bacillus subtilis with homologous and heterologous DnaA
protein. J Mol Biol 274:365-80.
Kurokawa, K., S. Nishida, A. Emoto, K. Sekimizu, and T. Katayama. 1999.
Replication cycle-coordinated change of the adenine nucleotide-bound forms of
DnaA protein in Escherichia coli. Embo J 18:6642-52.
Lee, P. S., and A. D. Grossman. 2006. The chromosome partitioning proteins
Soj (ParA) and SpoOJ (ParB) contribute to accurate chromosome partitioning,
221
31.
32.
33.
34.
35.
36.
37.
38.
39.
40.
41.
42.
43.
44.
45.
46.
separation of replicated sister origins, and regulation of replication initiation in
Bacillus subtilis. Mol Microbiol 60:853-69.
Lee, P. S., D. C. Lin, S. Moriya, and A. D. Grossman. 2003. Effects of the
chromosome partitioning protein SpoOJ (ParB) on oriC positioning and
replication initiation in Bacillus subtilis. J Bacteriol 185:1326-37.
Love, P. E., and R. E. Yasbin. 1984. Genetic characterization of the inducible
SOS-like system of Bacillus subtilis. J Bacteriol 160:910-20.
Mandelstam, J., and S. A. Higgs. 1974. Induction of sporulation during
synchronized chromosome replication in Bacillus subtilis. J Bacteriol 120:38-42.
Moriya, S., Y. Imai, A. K. Hassan, and N. Ogasawara. 1999. Regulation of
initiation of Bacillus subtilis chromosome replication. Plasmid 41:17-29.
Mukhopadhyay, S., and D. K. Chattoraj. 2000. Replication-induced
transcription of an autorepressed gene: the replication initiator gene of plasmid
Pl. Proc Natl Acad Sci U S A 97:7142-7.
Noirot-Gros, M. F., E. Dervyn, L. J. Wu, P. Mervelet, J. Errington, S. D.
Ehrlich, and P. Noirot. 2002. An expanded view of bacterial DNA replication.
Proc Natl Acad Sci U S A 99:8342-7.
Noirot-Gros, M. F., M. Velten, M. Yoshimura, S. McGovern, T. Morimoto, S.
D. Ehrlich, N. Ogasawara, P. Polard, and P. Noirot. 2006. Functional
dissection of YabA, a negative regulator of DNA replication initiation in Bacillus
subtilis. Proc Natl Acad Sci U S A 103:2368-73.
Ogura, Y., Y. Imai, N. Ogasawara, and S. Moriya. 2001. Autoregulation of the
dnaA-dnaN operon and effects of DnaA protein levels on replication initiation in
Bacillus subtilis. J Bacteriol 183:3833-41.
Ortenberg, R., S. Gon, A. Porat, and J. Beckwith. 2004. Interactions of
glutaredoxins, ribonucleotide reductase, and components of the DNA replication
system of Escherichia coli. Proc Natl Acad Sci U S A 101:7439-44.
Pak, M., and S. Wickner. 1997. Mechanism of protein remodeling by ClpA
chaperone. Proc Natl Acad Sci U S A 94:4901-6.
Quinones, M., H. H. Kimsey, and M. K. Waldor. 2005. LexA cleavage is
required for CTX prophage induction. Mol Cell 17:291-300.
Rana, R. S., and H. O. Halvorson. 1972. Nature of deoxyribonucleic acid
synthesis and its relationship to protein synthesis during outgrowth of Bacillus
cereus T. J Bacteriol 109:606-15.
Roberts, J. W., and R. Devoret. 1983. Lysogenic Induction, p. 123-144. In R.
W. Hendrix, J. W. Roberts, F. W. Stahl, and R. A. Weisberg (ed.), Lambda II.
Cold Spring Harbor Laboratory, Cold Spring Harbor, New York.
Rokop, M. E., J. M. Auchtung, and A. D. Grossman. 2004. Control of DNA
replication initiation by recruitment of an essential initiation protein to the
membrane of Bacillus subtilis. Mol Microbiol 52:1757-67.
Setlow, P. 1973. Deoxyribonucleic acid synthesis and deoxynucleotide
metabolism during bacterial spore germination. J Bacteriol 114:1099-107.
Sun, L., and J. A. Fuchs. 1992. Escherichia coli ribonucleotide reductase
expression is cell cycle regulated. Mol Biol Cell 3:1095-105.
222
47.
48.
49.
Sun, L., B. A. Jacobson, B. S. Dien, F. Srienc, and J. A. Fuchs. 1994. Cell
cycle regulation of the Escherichia coli nrd operon: requirement for a cis-acting
upstream AT-rich sequence. J Bacteriol 176:2415-26.
Theisen, P. W., J. E. Grimwade, A. C. Leonard, J. A. Bogan, and C. E.
Helmnstetter. 1993. Correlation of gene transcription with the time of initiation of
chromosome replication in Escherichia coli. Mol Microbiol 10:575-84.
Wu, L. J., A. H. Franks, and R. G. Wake. 1995. Replication through the
terminus region of the Bacillus subtilis chromosome is not essential for the
formation of a division septum that partitions the DNA. J Bacteriol 177:5711-5.
223