Co-Orientation of Replication and Transcription Preserves Genome Integrity

Article (PDF Available)inPLoS Genetics 6(1):e1000810 · January 2010with35 Reads
DOI: 10.1371/journal.pgen.1000810 · Source: PubMed
In many bacteria, there is a genome-wide bias towards co-orientation of replication and transcription, with essential and/or highly-expressed genes further enriched co-directionally. We previously found that reversing this bias in the bacterium Bacillus subtilis slows replication elongation, and we proposed that this effect contributes to the evolutionary pressure selecting the transcription-replication co-orientation bias. This selection might have been based purely on selection for speedy replication; alternatively, the slowed replication might actually represent an average of individual replication-disruption events, each of which is counter-selected independently because genome integrity is selected. To differentiate these possibilities and define the precise forces driving this aspect of genome organization, we generated new strains with inversions either over approximately 1/4 of the chromosome or at ribosomal RNA (rRNA) operons. Applying mathematical analysis to genomic microarray snapshots, we found that replication rates vary dramatically within the inverted genome. Replication is moderately impeded throughout the inverted region, which results in a small but significant competitive disadvantage in minimal medium. Importantly, replication is strongly obstructed at inverted rRNA loci in rich medium. This obstruction results in disruption of DNA replication, activation of DNA damage responses, loss of genome integrity, and cell death. Our results strongly suggest that preservation of genome integrity drives the evolution of co-orientation of replication and transcription, a conserved feature of genome organization.
Co-Orientation of Replication and Transcription
Preserves Genome Integrity
Anjana Srivatsan
, Ashley Tehranchi
, David M. MacAlpine
, Jue D. Wang
1 Department of Molecular and Human Genetics, Baylor College of Medicine, Houston, Texas, United States of America, 2 Department of Pharmacology and Cancer
Biology, Duke University Medical Center, Durham, North Carolina, United States of America
In many bacteria, there is a genome-wide bias towards co-orientation of replication and transcription, with essential and/or
highly-expressed genes further enriched co-directionally. We previously found that reversing this bias in the bacterium
Bacillus subtilis slows replication elongation, and we proposed that this effect contributes to the evolutionary pressure
selecting the transcription-replication co-orientation bias. This selection might have been based purely on selection for
speedy replication; alternatively, the slowed replication might actually represent an average of individual replication-
disruption events, each of which is counter-selected independently because genome integrity is selected. To differentiate
these possibilities and define the precise forces driving this aspect of genome organization, we generated new strains with
inversions either over ,1/4 of the chromosome or at ribosomal RNA (rRNA) operons. Applying mathematical analysis to
genomic microarray snapshots, we found that replication rates vary dramatically within the inverted genome. Replication is
moderately impeded throughout the inverted region, which results in a small but significant competitive disadvantage in
minimal medium. Importantly, replication is strongly obstructed at inverted rRNA loci in rich medium. This obstruction
results in disruption of DNA replication, activation of DNA damage responses, loss of genome integrity, and cell death. Our
results strongly suggest that preservation of genome integrity drives the evolution of co-orientation of replication and
transcription, a conserved feature of genome organization.
Citation: Srivatsan A, Tehranchi A, MacAlpine DM, Wang JD (2010) Co-Orientation of Replication and Transcription Preserves Genome Integrity. PLoS Genet 6(1):
e1000810. doi:10.1371/journal.pgen.1000810
Editor: Nancy A. Moran, University of Arizona, United States of America
Received September 21, 2009; Accepted December 10, 2009; Published January 15, 2010
Copyright: ß 2010 Srivatsan et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: JDW is supported by a Welch Research Grant (Q-1698) and the NIH Director’s New Innovator Award Program through grant number 1-DP2-OD004433-
01. DMM is supported by a Whitehead Foundation Scholar Award and the NIH Grant HG004279. The funders had no role in study design, data collection and
analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail:
The fundamental processes of replication and transcription take
place on the same template efficiently and accurately, requiring
them to be coordinated with each other to avoid potential
conflicts. In cells growing rapidly, both replication and transcrip-
tion of ribosomal RNA (rRNA) genes, and many other genes,
are initiated more frequently, further elevating this potential
conflict [1–4]. Due to the asymmetry of the replisome and the
transcription complex, the outcome of their encounter should
depend strongly on their relative directionality. RNA polymerase
(RNAP) is dislodged by replication in either direction [5,6]. On the
other hand, replication is affected mostly by head-on transcription
Preventing or resolving this conflict not only requires numerous
protein factors [13–16] but may also underlie several non-random
aspects of genome organization [17,18]. First, the highly-expressed
rRNA and tRNA genes are transcribed almost exclusively co-
directionally with replication across numerous species [19,20].
Chromosomes of the bacteria Bacillus subtilis and Escherichia coli are
replicated by bi-directional replication forks initiated from a single
origin (oriC), and all rRNA operons are oriented away from oriC
[21–25]. In yeast, replication fork barriers at the end of ribosomal
DNA operons prevent replication from entering head-on into
these strongly-transcribed regions [26]. Second, other highly-
transcribed genes are also significantly enriched in the leading
strand of replication in bacteria, ensuring that their transcription is
co-oriented with replication [27]. This feature may be conserved
in certain regions of the human genome [28]. Third, longer
transcription units are enriched in the leading strand [27,29].
Fourth, essential genes are enriched to a greater extent than non-
essential genes in the leading strand [19]. Finally, there is a general
bias for co-directionality of replication and transcription. In B.
subtilis and E. coli, this bias is 75% and 55% of all genes,
respectively [22–24].
Despite a general theme of avoiding head-on transcription and
replication, the precise evolutio nary forces shaping these inter-
connected aspects of genome organization are not understood.
The effect of head-on repl ication on tran scription is pro posed
to impact fitness negatively by interrupting the expr ession
of highly-transcribe d gene s [27], or in the case of essentia l
genes, by leading to the formation of incomplete transcripts,
which subsequent ly resu lts in toxic truncated polypeptides [19].
However, t he effects on replication are also del eterious. In E. coli,
replication rate is largely unaffected by co-directional transcrip-
tion, b ut is significantly slowed when it occurs head-on to a strong
transcription unit [5,30]. In addition, reversing transcription bias
over an extended segment of the B. subtilis genome leads to a
significant (30%) decrease of replication rate, extending the time
required to replicate the chromosome and potentially impe ding
PLoS Genetics | 1 January 2010 | Volume 6 | Issue 1 | e1000810
the cell cycle [31]. Head-on orientation of replication and
which can be due to obstructed replication or disrupted
transcription [ 32–35] . It is proposed that the transcription of
essential genes is preferentially co-oriented to lower their rate of
mutagenesis [30]. Finally, apart from effects on replication and
transcription, the t ranscription bias is also proposed to promote
chromosome segregation [36,37]. Is there a single evolutionary
advantage associated with the co-directional bias? Alternatively,
is the orientation of each gene selected in its own right? One
challenge in unde rstanding t he evol utionary bases of orientat ion
biases is dissecting how differen t aspects o f genome organ ization
are importa nt in different circumstances and how they im pact
cellular fitness.
Here we report that t he ex tent of the impact of head-on
transcription on replication differs b etw een genes within the same
organism B. subtilis. This was dissected by creating new inversions
of either an extensive region of the genome, or a localized region
containing strongly-transcribed rRNA genes. Using quantitative
genomic approaches, we observed differential rates of replication
throughout the genome of the inversion str ains—normal
replication in intact genomic positions, impedance of replication
elongation by ,30% within the head-on region, an d strong
blockage of repli cation at inverted rRNA operons. We further
characterized the fitness cost and found that inversion of the oriC-
proximal half of a replichore results in a small decrease in growth
rate in minimal medium, but is sufficient to confer a significant
com petitive disadvantage. On the other hand, the replication
block at rRNA operons leads to major disruption of replication,
induction of the DNA damage respo nse and cell death. We also
observed that the rate of mut ation of the gene rpoB is increased
when it is transcribed head-on to replication within an extended
chromosomal inversion, specif ically i n rich medium. Our results
strongly suggest that preservation of genom e integrity has
contributed to evolution of the genome-wide co-directional bias
and its further enrichmen t in highly-expressed and essential
Inversion of ,1/4 of the Bacillus subtilis chromosome to
reverse its transcription bias
We previously moved the origin of replication (oriC) away from
its endogenous position at 0u (Figure 1A) to 257u (not shown) or
94u (Figure 1B) to reverse the genomic transcription bias in an
extended region of the chromosome, and observed that replication
elongation was slowed moderately between 0u and the ectopic
oriC position due to transcription [31]. This raised the intriguing
question: what would be the potential impact of reversed
transcription bias on cellular fitness and genome integrity?
However, this question cannot be answered using these strains,
because other aspects of their genome organization were also
altered, including location of oriC and symmetry of the replichores
(one spanning L of the chromosome, the other J of the
chromosome). Such alterations have been shown to strongly
impact cellular fitness in both E. coli and B. subtilis [38–40].
To examine exclusively the biological impact of reversed
transcription bias, we constructed several new strains. We took
advantage of the fact that the strain with repositioned oriC
(Figure 1B) has oriC-flanking sequences present both at 0u and the
ectopic location (Figure 1C). We reasoned that homologous
recombination might occur at these repeats, and screened for such
progenies (Figure 1D and 1E). Homologous recombination of
repeats upstream of oriC (Figure 1C- repeats marked L, ,400 bp)
repositioned oriC to 0u, with concurrent inversion of the 0u–94u
portion of the chromosome. The resulting strain had
transcription (HT) between 0u and 94u and equal replichore
lengths (Figure 1D). Using the same strategy, we also obtained
strains in which homologous recombination had taken place
between repeats downstream of oriC (Figure 1C- repeats marked
R, ,300 bp). The resulting chromosomes have oriC positioned at
94u and
unequal replichores (UR) but without extended regions of
head-on transcription (Figure 1E). Finally, to minimize the
possibility of reversion of the inversion, we removed a portion of
the remnant homology region at 94u.
Using the HT strain, we evaluated the impact of head-on
transcription on fitness by first examining its exponential growth
(Figure 1F). The HT strain grew slowly in rich medium (LB) with a
doubling time of 70 minutes, compared to 20 minutes for the
control. In minimal medium however, the doubling time of the
HT strain was similar to that of the control (53 and 52 minutes,
respectively). In contrast, the UR strain was sicker than the control
in both LB and minimal media (doubling times of 52 and
77 minutes, respectively), indicating a general growth defect due to
differing replichore lengths and/or ectopic positioning of oriC.
Therefore, unlike the general growth defect introduced by uneven
replichores, the growth defect caused by inverting transcription
bias over J of the chromosome is nutrient-dependent.
Head-on transcription decreases replication fork speed
on a genomic scale
We next examined whether head-on transcription has an effect
on replication in the HT strain. We monitored synchronized fork
progression in this strain using genomic microarrays (Figure 2A),
in minimal medium where no significant loss of growth rate was
observed (Figure 1F). Cells were synchronized for their replication
cycles using a temperature-sensitive allele of the replication protein
DnaB [41,42]. The gene dosage profile obtained 30 minutes after
replication initiation indicates that ,50% of cells initiated
replication. The average position of replication forks can be
estimated as the midpoint of the transition between replicated and
unreplicated genomic positions [43]. This position is ,0.68 Mbp
Author Summary
An important feature of genome organization is that
transcription and replication are selectively co-oriented.
This feature helps to avoid conflicts between head-on
replication and transcription. The precise consequences of
the conflict and how it affects genome organization
remain to be understood. We previously found that
reversing the transcription bias slows replication in the
Bacillus subtilis genome. Here we engineered new
inversions to avoid changes in other aspects of genome
organization. We found that the reversed transcription bias
is sufficient to decrease replication speed, and it results in
lowered fitness of the inversion strains and a competitive
disadvantage relative to wild-type cells in minimal
medium. Further, by analyzing genomic copy-number
snapshots to obtain replication speed as a function of
genome position, we found that inversion of the strongly-
transcribed rRNA genes obstructs replication during
growth in rich medium. This confers a strong growth
disadvantage to cells in rich medium, turns on DNA
damage responses, and leads to cell death in a subpop-
ulation of cells, while the surviving cells are more sensitive
to genotoxic agents. Our results strongly support the
hypothesis that evolution has favored co-orientation of
transcription with replication, mainly to avoid these
Head-On Transcription Disrupts Replication
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from oriC on the left replichore and ,0.56 Mbp on the inverted
right replichore (Figure 2A, blue arrows), indicating that
replication forks move slower within the inverted region. We
inhibited transcription initiation by adding the drug rifampicin
4 minutes after synchronized replication began, and found that in
rifampicin-treated cells replication forks progressed further in the
inverted region (,0.72 Mbp) (Figure 2A, red arrows) compared to
untreated cells (,0.56 Mbp), demonstrating that the reduction in
replication fork speed is due to transcription. This reduction in
fork movement does not lead to proportionally slower growth
likely because B. subtilis has flexible cell division cycles and can
compensate for slower fork progression via multifork replication
Nutrient-dependent changes in the asynchronous gene
dosage profile
We next examined whether the decreased replication rate in the
inverted region varied depending on nutrient status and genomic
position. To this end, we obtained the genomic microarray profile
of the HT strain during exponential growth (Figure 2B). Cells are
not synchronized and hence the positions of the replication forks
would vary from cell to cell (Figure 2B inset). Importantly, by
assuming that these cells are in a steady state and their genomic
profile is time-invariant on a population basis, we can use this
profile to calculate replication speed at every position on the
chromosome. This speed is inversely correlated with the local
slope of log values of the gene dosage with respect to gene positions
(see Materials and Methods).
We first observed that in minimal medium, the genomic
profile was smooth but asymmetric (Figure 2B, blue). The rates
of replication were similar in the unaltered regions of the
chromosome, indicated by similar slopes on the left replichore
(172u to 360u, 0.40860.002/Mbp) and the non-inverted region on
the right replichore (94u to 153u, 0.46260.007/Mbp) (Table 1). In
contrast, within the inverted segment on the right replichore (0u to
94u), the slope was 0.64660.004/Mbp, indicating a ,30%
decrease in fork speed within the head-on region, in agreement
with our previous results using an ectopic oriC [31].
Interestingly, when cells were grown in a relatively rich medium
(minimal medium supplemented with casamino acids, hereafter
referred to as CAA), the gene dosage profile changed sharply at
specific locations on the chromosome (Figure 2B, orange). These
transition points could be differentiated more clearly when we
derived the relative gene dosages between the profiles in CAA vs.
minimal medium (Figure 2C), and clearly corresponded to the
positions of the rRNA operons within the inverted segment
(Figure 2B and 2C, green arrows). The steep slopes at these
transitions suggest that replication progression is strongly impeded
and even stalled at these locations. Other than at rRNA loci, the
genomic profile in CAA was very similar to that of minimal
medium including at the intact rRNA operon on the left replichore
(Figure 2C, orange arrow). Our results indicate that the genome-
wide impedance of replication by reversed transcription bias other
than at rRNA loci is largely uniform and unaltered by nutrient
conditions. Importantly, we identify rRNA loci as positions of
strong, nutrient-dependent obstruction of fork movement when
they are transcribed head-on.
Head-on transcription of rRNA disrupts replication and
induces the DNA damage response
We next examined whether the forks obstructed by head-on
transcription during growth in CAA were also disrupted. The
Figure 1. Alteration of different aspects of genome organization results in different growth defects. (A) Schematic diagram of the wild
type B. subtilis chromosome (black circle). oriC at 0u and terC at 172u represent the origin and terminus of DNA replication, respectively. Orange
arrows: replication; grey arrows: predominant direction of transcription; blue arrowheads: rRNA operons. (B) Schematic diagram of a mutant
chromosome with oriC relocated to 94u [31], resulting in head-on transcription and unequal replichores (HT+UR). Green arrow: replication head-on to
transcription between 0u and 94u. (C) Schematic diagrams of homologous recombination events leading to inversions of J of the chromosome. L
and R: sequences flanking oriC that are repeated at 0u and 94u; spc: spectinomycin resistance gene, which was inserted to replace the endogenous
oriC at 0 u [31]. Recombination at L (dashed lines) and R (dotted lines) give rise to the chromosomes shown schematically in (D,E), respectively. (D)
Schematic diagram of the chromosome with an inversion between 0u and 94u resulting in head-on transcription (HT). (E) Schematic diagram of the
chromosome with an inversion between 0u and 94u resulting in unequal replichores (UR). (F) Doubling times at 37uC in liquid LB (red bars), minimal
medium with (CAA, grey bars) or without (Min, black bars) casamino acids, calculated by measuring OD
. Data shown are for strains in the JH642
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Figure 2. Genomic microarray profiles of the HT inversion strain. (A) Synchronized replication profile of the HT strain. Cells containing the
dnaB134ts allele were grown in Min at 30uC and their replication cycle was synchronized by shifting to 45uC for 60 minutes, and then back to 30uCto
allow replication initiation. Genomic profiles were obtained 30 minutes after initiation, relative to pre-initiation reference DNA. Profiles were obtained
without (blue) or with (red) addition of rifampicin (rif) 4 minutes after initiation to inhibit transcription. Elevation of the baseline of the gene dosage
profile on the right replichore is likely due to incomplete replication of the pre-initiation reference DNA of the HT strain, despite the 60 minutes
incubation at 45uC. Grey shaded region: inversion; blue and red arrows: average positions of replication forks in the absence (blue) or presence (red)
of rif. Insets: schematic diagram of chromosomes of the HT strain undergoing synchronized replication with (top), and without (bottom) transcription
(grey arrows). Purple ovals: replisomes. (B) Overlay of the asynchronous genomic profiles of the HT strain grown in Min (blue), and CAA (orange).
Profiles were obtained from asynchronous cultures grown at 37uCtoOD
,0.5. Average gene dosage ratios (log
) are plotted relative to gene
positions adjusted according to deletion of the phage SPb and the integrative and conjugative element ICEBs1 clusters. The prophage-like skin
element is not removed in this strain, which likely explains the small peak at 21.5 Mbp (black arrow). Green arrows: sharp changes in slope at rRNA
loci in the CAA profile. Inset: schematic diagram of chromosomes of the HT strain undergoing asynchronous replication. (C) Expanded view of the
ratio of gene dosage (log
) in CAA versus Min within the region outlined in B (21.2 Mbp to 1.2 Mbp). Green arrows: positions of inverted rRNA
operons; orange arrow: position of co-directional rrnB; grey shaded region: inversion.
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recombination protein RecA localizes to stalled replication forks as
foci only when the forks are disrupted [44]. Hence, we examined
the sub-cellular localization of RecA using a recA-gfp fusion
construct [45]. We visualized microscopically cells carrying this
allele grown in CAA, and found that 97% of the cells of the HT
strain had RecA foci compared to 27% in the control (Figure 3A
and 3B), indicating that replication forks are disrupted by head-on
We also noticed that the HT strain had abnormal nucleoid
morphology, which appeared filamented and even fragmented in
some cases (Figure 3A, lower panels). Cell lengths also significantly
increased (not shown) and chain lengths doubled (Table 2), which
might explain the ,2-fold decrease in the number of colony-
forming units in strain HT compared to its control (Table 3).
In addition to RecA foci formation, a subpopulation of cells
exhibits the SOS DNA damage response (Figure 3C). Using a
GFP-fusion reporter of tagC, a member of the SOS regulon [46],
we found that the SOS response is induced in greater than 6% of
single cells of the HT strain in LB medium, but in less than 1% in
minimal medium (Figure 3C). The increase in SOS response in
rich medium is accompanied by increased cell death. We
performed live/dead staining in which the nucleoids of dead cells
with permeable membranes stain with propidium iodide (red),
while live cells stain with SYTO9 (cyan) (Figure 3D). There was a
marked increase in the fraction of dead cells in the HT strain
relative to the isogenic control, again specifically in rich medium
(12% versus 1% of cells in LB) (Figure 3E). This suggests that
failure to repair replication forks disrupted by strong head-on
transcription might lead to failure to complete replication and cell
death. In agreement with this hypothesis, we found that HT cells
show higher sensitivity to the genotoxic agent mitomycin C in rich
medium (Figure 3F), suggesting that exogenous DNA damage adds
further demand on their already overwhelmed DNA repair
capacity, and dramatically elevates cell death in the population.
Increased mutation rate of a gene transcribed head-on to
Having obtained evidence of disruption of replication by
head-on transcription, we next examined whether it also has a
consequence on genome stability by measuring the rates of
mutations conferring resistance to the drug rifampicin (rif
). In B.
subtilis, rif
mutations map to rpoB [47,48], which is transcribed co-
directionally to replication in the wild-type strain but head-on in
the HT strain (Figure 4A). Since rpoB encodes a sub-unit of RNA
polymerase, its mutation might confer a growth advantage in this
scenario. Hence we analyzed the results of the fluctuation test with
the P
method, which measures the rate of mutation indepen-
dently of its effect on growth rate [49]. We observed that the
mutation rate increased ,3-fold in rich medium in the HT strain
compared to an isogenic control with no inversion (Figure 4C).
This could be due to a global cellular response to disruption of
replication caused by the chromosomal inversion. To examine this
possibility, we also monitored the rif
mutation rate in a strain
with inversion of K of the left replichore, leaving the rpoB region
unaltered (Figure 4B). This strain has the same extent of reversed
transcription bias as the HT strain, exhibits a similar nutrient-
dependent growth defect (Table 4, Figure S1A), and has one
inverted rRNA operon located near oriC after chromosome
inversion, which causes replication blockage in rich medium, as
monitored by microarrays (Figure S1B and S1C). However, there
was no increase in the rif
mutation rate in this strain (Figure 4C).
Therefore the presence of an inversion alone is not sufficient to
cause an increase in rif
mutation rate, rather it is specific to the
strain in which rpoB is within the inverted region. In minimal
medium all strains had similar rif
mutation rates (Figure 4D).
Inversion of ribosomal RNA operons is sufficient to
disrupt replication
The most dramatic reduction of replication speed due to head-
on transcription occurs at the rRNA operons within the inversion
(Figure 2C). This suggests that the nutrient-dependent effect on
replication fork progression is mostly due to inversion of the
strongly-transcribed rRNA operons. We tested this hypothesis by
examining the consequences of specifically inverting rRNA
operons. We inverted the rrnIHG cluster (,17 kbp) that contains
3 rRNA operons and 6 tRNA genes, by inserting two overlapping
halves of the neomycin resistance gene (neo) flanking the cluster,
and selecting for recombination events that created a complete neo
gene, similar to [50] (Figure 5A and 5B). The rrn inversion strain
was inviable in LB and had a strong growth defect in CAA
compared to the pre-inversion control (doubling times of 44 and
28 minutes, respectively), while their doubling times in minimal
medium were similar (44 and 42 minutes, respectively) (Figure 5C).
These results indicate that the growth defect of the HT strain was
mostly due to inverted rRNA genes.
We next examined asynchronous replication in the rrn inversion
strain using genomic microarrays, and observed impedance
specifically at the inverted loci, where the gene dosage profile
Table 1. Slopes of asynchronous gene dosage profiles in the HT inversion strain.
Medium Replichore Boundaries Transcription bias Slope (Mbp
Min Left oriC (0u)-terC (172u) co-directional 0.408 (60.002)
Right oriC (0u)-aprE (94u) head-on 0.646 (60.004)
Right aprE (94u)-pksE (153u) co-directional 0.462 (60.007)
CAA Left oriC (0u)-terC (172u) co-directional 0.450 (60.002)
Right oriC (0u)-rrnD (13u) head-on 0.640 (60.101)
Right rrnD (13u)-rrnE (40u) head-on 0.615 (60.031)
Right rrnE (40u)-rrnGHI (80u) head-on 0.654 (60.015)
Right rrnGHI (80u)-aprE (94u)
head-on 1.51 (60.068)
Right aprE (94u)-pksE (153u) co-directional 0.456 (60.007)
includes four rRNA operons, all transcribed head-on to replication (rrnJW, rrnA and rrnO).
SE standard error.
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showed sharp discontinuity (Figure 5D and 5E). This indicates that
a significant number of replication forks were stalled in this short
segment. This effect was much stronger in cells replicating asyn-
chronously in CAA than in minimal medium. Further, the majority
of cells with the rrn inversion had RecA-GFP foci/filaments when
Figure 3. The HT strain exhibits disruption of DNA replication and loss of genome integrity. (A) RecA localization and nucleoid
morphology in the HT and isogenic control strains in CAA. Upper and lower panels: phase contrast images (blue) overlaid with RecA-GFP (green) or
DAPI (white) fluorescence images, respectively. (B) Percentages of cells with RecA-GFP foci in the indicated media. Cells were grown in CAA or Min
and stained with DAPI and the membrane dye FM4-64 (not shown) to count individual cells. Light grey bars: control; dark grey bars: HT strain. The HT
strain with recA-gfp was extremely sick in LB. (C) Induction of the SOS DNA damage response in the HT and isogenic control strains in the indicated
media, as monitored using a TagC-GFP reporter. The total length of SOS-positive cells was divided by the total length of cells in each field. Light grey
bars: control; dark grey bars: HT strain. (D) Live/dead staining of the HT and control strains in LB and Min. The fluorescent dyes SYTO9 (cyan) and
propidium iodide (red) were added to exponential phase cultures (OD
,0.2–0.6) to label live and dead cells, respectively. (E) Average percentages
of dead cells in the control (light grey bars) and HT (dark grey bars) strains in different growth media (LB, CAA and Min). The cell length that was
stained with propidium iodide was divided by the total length of cells in each field. (F) Mitomycin C (MMC) sensitivity of the HT and isogenic control
strains in LB and Min. Cells were grown to OD
= 0.3 and serial dilutions ranging from 10
to 10
were spotted on LB and Min plates with or
without MMC (0.0625
mg/ml). Plates were scanned after overnight incubation at 37uC. Scale bar in A and D: 5 mm.
Table 2. Increased length of chains of cells in rich medium.
Strain Relevant genotype
Average chain
length (6SE) (mm)
JDW712 Isogenic control for JDW713 23.09 (69.38)
JDW713 Stabilized HT strain 59.19 (66.61)
JDW858 Pre-inversion control for JDW860 15.20 (67.56)
JDW860 rrnIHG inversion 26.99 (62.44)
SE standard error.
Table 3. Colony-formation by inversion strains in different
Strain Relevant genotype
Number of cfu/ml
LB Min
JDW712 Isogenic control for JDW577 0.76 (60.3) 3.20 (60.9)
JDW713 Stabilized HT strain 0.44 (60.15 ) 5.14 (61.6)
JDW858 Pre-inversion control for JDW860 1.9 (60.14) 3.6 (60.283)
JDW860 rrnIHG inversion 0.293 (60.186) 2.63 (61.484)
normalized with respect to OD
JDW860 cannot grow in LB.
cfu colony-forming units.
SD standard deviation.
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grown in CAA (Figure 5F and 5G), indicating that strong head-on
transcription at this cluster is sufficient to disrupt replication forks.
Notably, the nucleoid morphology of the rrn inversion strain was
largely normal (Figure 5F, lower panels), indicating that the gross
nucleoid defects of the HT strain were not due to disruption of
replication by rrn inversion.
Finally, the rrn inversion lowered cell viability. The rrn inversion
strain had a much higher fraction of dead cells relative to the pre-
inversion control in CAA (10% versus 0.7%, respectively)
(Figure 5H and 5I). Thus strong head-on transcription of just
rRNA and tRNA operons drastically impacts cell viability through
disruption of replication.
Figure 4. Increased mutation rate of a gene within the inversion in the HT strain. (A,B) Schematic diagrams of chromosomes (black circles)
with inversion of K of either the right (A) or the left (B) replichore, resulting in head-on transcription (HT) within the inversion. Orange and green
arrows: replication co-directional and head-on to transcription, respectively; grey arrows: predominant direction of transcription. Mutations
conferring resistance to rifampicin (rif
) map to the rpoB gene (red arrow) which is transcribed co-directional to replication in the wild type and left
replichore HT strains, but transcribed head-on in the right replichore HT strain. (C) Spontaneous rif
mutation rates as measured by fluctuation tests in
LB (dark grey bars) in the control, right and left replichore HT strains. *: rif
rate in the right replichore inversion strain is significantly different from
control (P,0.05). (D) Rif
mutation rates in Min (light grey bars).
Table 4. Doubling times at 37uC.
Strain Relevant genotype Doubling times (minutes) (6SD)
JH642 Wild type control 20 (60.8) 24 ( 62.1) 52 (63.1)
Inversion 0u to 94u, with head-on transcription 70 (68.8) 54 ( 64.2) 53 (64)
Inversion 0u to 94u, with unequal replichores 52 (66.2) 43 ( 61.4) 77 (64)
oriC at 94u: head-on transcription and unequal replichores 149 (621) 115 (615.6) 92 (63.4)
Isogenic control for JDW577 25 (61.8) 28 (60.5) 47 (60.5)
Stabilized inversion 0u to 94u, with head-on transcription 86 (63) 50 (63) 53 (61.7)
Inversion 0u to 257u, with head-on transcription 62 (63.6) 39 ( 62.7) 58 (65.7)
Isogenic control for JDW713 21 (61.4) 28 (60.4) 44 (61.8)
Stabilized inversion 0u to 94u, with head-on transcription 72 (64.7) 55 ( 66.9) 44 (60.6)
Pre-inversion control for rrnIHG inversion (ybaN to rrnG-5S)20(60) 28 (60.7) 42 (61)
Pre-inversion control for rrnIHG inversion (ybaJ to rrnG-5S)20(60) 28 (60.7) 42 (61)
rrnIHG inversion (ybaN to rrnG) .160 44 (64.9) 44 (61.0)
rrnIHG inversion (ybaJ to rrnG) .160 54 (65.0) 46 (64.6)
JH642 background.
YB886 (prophage ‘‘cured’’) background.
SD standard deviation.
Head-On Transcription Disrupts Replication
PLoS Genetics | 7 January 2010 | Volume 6 | Issue 1 | e1000810
Genome organization has evolved to enhance fitness, as
evidenced by observations that certain genome rearrangements
are not tolerated, or cause growth defects [17,38,39,51–56], while
others do not and might even be prevalent [57]. One feature of
genome organization is that it precludes extensive head-on
transcription [5,17,18,27,31]. In this study we engineered B.
Figure 5. Inversion of
impedes replication, triggers RecA recruitment, and elevates cell death. (A,B) Schematic diagrams of the
chromosomes (black circles) of the pre-inversion control strain (A) and the rrnIHG (rrn) inversion strain (B). Blue arrowheads: rRNA operons; large
green arrowhead: rrnIHG cluster. (C) Doubling times of the pre-inversion and rrn inversion strains at 37uC in CAA (grey bars) and Min (black bars). (D,E)
Asynchronous gene dosage profiles of the rrn inversion strain in CAA (D) and Min (E). Profiles were obtained from asynchronous cultures grown at
37uC in the indicated media to OD
,0.5. Average ratios of copy number (log
) in the rrn inversion strain relative to fully-replicated control are
plotted against the genomic position. Arrows indicate the position of the rrnIHG inversion. (F) RecA recruitment and nucleoid morphology in the pre-
inversion and rrn inversion strains in CAA, obtained as described in Figure 3A. Blue: phase contrast images; green: RecA foci; white: nucleoids stained
with DAPI. (G) Average percentage of cells with RecA recruitment in the indicated media. Light grey bars: pre-inversion control; dark grey bars: rrn
inversion. (H) Live/dead staining of the pre-inversion and rrn inversion strains grown in CAA and Min. Images were obtained as described in Figure 3D.
Cyan: live cells; red: dead cells. (I) Average percentages of dead cells in the pre-inversion (light grey bars) and rrn inversion (dark grey bars) strains. The
cell length that was stained with propidium iodide was divided by the total length of cells in each field. Scale bar in F and H: 5
Head-On Transcription Disrupts Replication
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subtilis strains with the co-directional bias of replication and
transcription reversed over either an extended segment of the
genome or at a localized rRNA gene cluster. We employed
microarray-based copy number profiling that enabled us to
visualize directly the replication status of the entire genome, to
identify positions at which there are significant perturbations and
to quantify the extent of such perturbations. We found that
replication is affected by head-on transcription in at least two
ways: one is the apparently uniform deceleration throughout the
extended region of reversed transcription bias, and the second and
stronger is the disruption of replication at highly-transcribed
rRNA loci. Disruption of replication at rRNA genes activates
DNA repair pathways and results in sensitivity to genotoxic stress
and loss of viability. Together these observations support the
hypothesis that head-on collisions between transcription and
replication result in loss of genome integrity, and that avoidance
of this consequence contributes to the evolution of co-orientation
bias in genomes.
Other aspects of genome organization
Previously we had engineered B. subtilis strains in which the
replichores were unequally distributed, and a significant portion of
the genome was replicated by forks traveling in the opposite
direction to transcription. We found that DNA replication
elongation was impeded within the region of reversed transcription
bias and these strains had strong growth defects [31]. However it
was not possible to attribute the growth defects to reversed
transcription bias alone since uneven replichores themselves have
been shown in E. coli to lead to strong growth defects and
dependence on recombination and/or DNA translocation ma-
chineries for viability [38,39]. Therefore, we have constructed
several new strains that separate the alteration of replichore
symmetry from that of the transcription-replication bias (Figure 1).
We found that the B. subtilis HT strain with head-on
transcription and replication exhibits a strong growth defect only
in rich medium, and the UR strain with asymmetric replichores
exhibits growth defects regardless of growth medium. The effect of
both aspects of genome alteration on fitness is approximately
multiplicative (Figure 1F and Table S1) [58,59]. There are small
deviations but this is significant only in minimal medium
(p = 0.01). Thus we were able to largely separate two aspects of
genome organization–the impact of unequal replichore size, and
the impact of colliding transcription and replication forks, on
genome integrity and cellular fitness.
We found that the HT strain with the extended inversion also
exhibits strong disruption of replication and induction of DNA
damage response in rich medium. In addition, it has altered
nucleoid morphology and a long and twisted cell shape especially
in rich medium (Figure 3A and 3D). We further inverted only an
rrn cluster, and demonstrated that it is sufficient to cause disruption
of replication, but not the nucleoid morphology defect. The
change in morphology in the HT strain can be either associated
with head-on transcription at locations other than rRNA operons,
or with additional effects of inverting J of the chromosome. The
latter could include the alteration of gene positions relative to
the origin, which affects their dosage [4,60], ectopic localization of
the parS sites which affect chromosome organization [61–63], or
defects in chromosome segregation which is proposed to be
facilitated by transcription [36,37].
Relationship between genome-wide replication rate and
Using synchronized microarrays, we demonstrate that inverting
the transcription bias over J of the chromosome decreases fork
progression rate in a transcription-dependent manner (Figure 2A),
confirming our previous results obtained using an engineered
strain with an ectopic oriC [31]. However, transcription through-
out the genome is not uniform. To examine whether different
inverted transcription units have different impacts on replication,
we obtained asynchronous microarray profiles of exponentially
growing cells (Figure 2B). Using mathematical analysis to obtain
the rate of replication as a function of genomic position, we
confirmed a modest genome-wide impedance of replication
throughout the inverted region and showed that it is mostly
independent of gene position. An important exception is at rRNA
and tRNA operons, evidenced by the punctuated pattern of
replication stalling at these clusters. Further, this stalling is strongly
potentiated by growth in rich medium.
In general, growth in r ich medium results in higher ini tiation
frequencies of both replication and rRNA transcrip tion [1–4],
either of which could elevate the conflict between transcription
and replication, thereby accounting for the observed increase
in replication stalling. However, we observed that in the right
replichore HT inversion strain, replic ation is not initiated
more frequently in rich medium than in minimal medium. In
contrast to wild type cells where gene dosage at oriC is much
higher in rich medium than in poor medium, indicating higher
rate of replication i nitiation [31], gene dosage at oriC in the
HT strain is similar in CAA and minimal media (Figure 2B).
The genomic profiles of the HT strain in the two media are
almost identical except at the inverted rrn loci (Figure 2C). It is
possible that failure of re plication elongation prevents subse-
quent replica tion initiati on; alternatively, replication initiation
frequency could be lower becau se it is couple d to growth [3,4],
which i s slower for the HT strain in rich medium (Figure 1F).
Regardless of the reason, it is clear that in the inversion strains,
the effect on replication obs erved at r RNA operons in rich
medium is not due t o increased replication but is exclusively
due to stronger rRNA transcr iption, which is initiated m ore
frequently because of higher iNTP and lower (p)ppGpp levels
Several models exist to explain why re plication is stalled by
strong head -on transcriptio n of rRN A operons. The replisome
might be capable of bypassing a single head-on RNAP, but the
presence of multiple RNAPs on the long and highly-transcribed
rrn region could make it harder for the replisome to proceed. In
addition, since rRNA operons are highly structured regions,
their transcription might obstruct replication forks, as p roposed
for other unusually structured regions [8,65]. RNAPs might also
stall upon hea d-on rep lication to form backe d-up RNAP arrays.
Backed-up RNAP arrays can create a barrier to replication
[15]. Finally, head-on transcription might create RNA-DNA
duplexes or supercoiling of DNA that poses a barrier to
replication [66–68], and this barrier might strengthen to the
extent of blocking replication when transcription i s sufficiently
Impact of head-on transcription on fitness and genome
Inverting the transcriptional bias of 1/4 of the B. subtilis
chromosome slows replication rate within this region by , 30%.
The growth rate of the HT inversion strain in minimal medium is
not significantly affected (Table 4). However we discovered that
the HT strain indeed has a significant growth disadvantage even in
minimal medium when competing with wild type cells, with its
relative fitness being 0.92 (+/20.07) (after factoring in the marker
effect) (Table S2, Text S1). This selective effect enables the wild
Head-On Transcription Disrupts Replication
PLoS Genetics | 9 January 2010 | Volume 6 | Issue 1 | e1000810
type strain to take over after multiple generations and clearly is
sufficient to shape genome evolution.
The impact of inversions on repl ication and cellular fitness is
rRNA genes in b oth the HT and rrn strains result in repli cation
blocks (Figure 2 and Figure 5) and likely lead to extensively
delayed cel l-cycle progression, which ex plains the dramatic
increase of doubling time in rich medium (Figure 1F and
Figure 5C). Indeed, blockage of replication elongation has bee n
shown to prevent cell proliferation in E. coli, which can only be
reversed upon removal of the barrier [69]. More importa ntly, we
obtained strong e vidence th at th e obstruction creat ed by i nverted
rRNA transcription also leads to disruption of replication. First,
RecA forms foci/filaments in the majority of these cells in rich
medium, indicating g eneration of single-stranded DNA or
double-stranded ends (DSEs). Second, there is a significant
increase in induction of the SOS DNA damage response [70] in
the inversion strains in rich medium (Figure 3C). In B. subtilis the
SOS response is not robustly turned on by DSEs due to efficient
repair by RecN, and our observation of ,6% of cells of the HT
strain showing SOS induction agrees with the reported value
[71]. Third, an increased number of cel l deaths occur i n the
inversion s trains especially in rich medium, likely due to failure t o
repair damaged replication forks. Finally, the inversion renders
cells more sensitive to the genotoxic agent mitomycin C
especially in rich medium, suggesting that the DNA r epair
capacity in these cells is highly compromised due to overwhelm-
ing demand, leading to detrimental consequences upon challenge
by e xternal DNA damag e.
It remains unclear whether replication fork collapse [72]
takes place soon after forks are s talled by head-on transcription,
or only when a s econd round of replication forks collides with a
prior round of stalled replication forks, as was demonstrated
previously at the replication terminator sequence in E. coli [73].
It is possible that both types of collisions would lead to
disruption of replication, with collision of sub sequent replica-
tion forks being more costly. This might explain why inve rting
rrnIHG near the origin of replication results in a particularly
strong growth defect, as the second fork encounters the stalled
first fork soon after it initiates from the origin of replication.
Indeed the oriC-proximal region is the chromosomal location of
highly-expressed genes, especially those involved in macromo-
lecular s ynthesis [ 74], and thus inversions here might be
particularly detrimental.
Disruption of replication has been shown to lead to higher levels
of genome instability especially in the vicinity of the disruption
[73,75,76]. We observed an increase in rpoB mutation rate when
the genomic region encoding rpoB is inverted (Figure 4). This
increase is only observed when cells are grown in rich medium,
implicating a dependence on the level of rpoB transcription or
disruption of replication. It is unlikely to be solely due to a global
cellular response to disruption of replication such as the SOS
response, since inversion of a symmetric half of the other
replichore without affecting rpoB, did not elevate its mutation rate
despite the fact that replication is clearly inhibited by inverted rrnB
transcription in this strain (Figure 4 and Figure S1). One possibility
is that disruption of replication at rpoB results in recruitment of an
error-prone DNA polymerase via RecA. RecA has been shown to
activate directly an error-prone DNA polymerase in E. coli [77].
Further work will be required to differentiate this possibility from
other remaining possibilities, e.g., that the increased mutation rate
is due to altering rpoB location rather than orientation, or due to
disruption of replication at the neighboring rRNA loci, or that the
rpoB mutagenesis is due to a threshold level of the SOS response
that is only met upon inversion of the right, but not the left
The impairment of replication by transcription has been
shown to result in transcription- associated recombination [32]
or deletion [34]. Replication orientation significantly influences
the spectrum of point mutations in yeast [35]. This suggests
that impairment of genome replication might also contribute to
transcription-associated mutagenesis [78–81]. Another major
mechanism which might be at play is the direct activation of
the error-prone translesion polymerase via its interactions with
transcription factors such as NusA [82].
Implications for the evolution of genome organization
Our observations offer st rong support for the hyp othesis that
the effect of transcription on r eplication is an important dri ving
force in the evolu tion of genome organization [18]. In a ddition,
our work suggests that the precise cost might vary depending on
both the gene type and the growth environment. First, inversion
of the strand bias of transcription within an extended segment of
the chromosome results in a small growth defect in n utrient-
poor me dium yet is suf ficient to c onfer a str ong c ompetitive
disadvantage. Second, the inversion of rRNA operons leads
to disruption of DNA replication, which is especially costly if
cells are grown in rich medium (Figure 2, Figure 3, and
Figure 5). This explains why rRNA operons are all oriented co-
directionally. Although the most prominent disruptive effects
we observed were at inverted rRNA operons, it is possible that
these effects can also be extended to other highly-expressed or
mutation rate is higher for the rpoB gene transcribed head-on,
supporting a model that co-orientation of transcription and
replication of essential genes might have evolved to avoid their
mutagenesis [30] (Figure 4). In addition, highly-expressed non-
essential genes are also intolerant of mutagenesis since it could
result i n r eduction of their expressivity. Thus minimizing
mutagenesis may also unde rlie the orientation bias of highly-
expressed genes.
There are considerable differences in transcription orientation
biases among organisms. While low G+C Firmicutes (such as B.
subtilis) and the Mycoplasmas have strong transcription orientation
biases, other bacteria do not [18]. The widely-studied bacterium
E. coli has only 55% co-orientation bias. Interestingly, inversion of
long segments of the E. coli replichores including rRNA operons
results in no growth defect [38]. What causes such a drastic
difference in the penalty of head-on transcription? There are at
least three possible explanations: differences in the composition of
their replication machineries [24], non-replicative helicases [16]
or transcription factors (AT and JDW unpublished). These
considerable differences might underlie the differential abilities
of organisms to cope with conflict between transcription and
replication and thus influence the evolution of their genome
Materials and Methods
Media and growth conditions
Cells were grown in LB or defined minimal medium (50mM
MOPS) [83] with 1% glucose, supplemented with 40
tryptophan, 40
mg/ml methionine, 40 mg/ml phenylalanine,
mg/ml arginine, or casamino acids (CAA) (Difco) (0.5%).
Cells containing inversion of rRNA operons were maintained on
mg/ml kanamycin (kan). Chloramphenicol (cm), erythromycin
(erm), and spectinomycin (spc) were used as described at 5, 0.5 and
mg/ml, respectively.
Head-On Transcription Disrupts Replication
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Strains and plasmids
Standard techniques were used for genetic and molecular
biological manipulations [84]. Strains used are listed in Table S3.
Primer sequences are listed in Table S4. Strains were constructed
in the JH642 background [85], and in the YB886 phage-defective
background [86] because phage excision and duplication during
stressful conditions often create localized gene copy number
alterations. YB886 is cured of phage SPb, defective for phage
PBSX induction [86], and also lacks the transposon-like element
ICEBs1 [87].
Strain JDW704 was created by introducing an ectopic oriC at
aprE (94u) in YB886 and then deleting the endogenous oriC (0u),
using genomic DNA from JDW258 [31] and MMB703 [40],
respectively. Progenies of JDW704 were screened for recombina-
tion events in sequences flanking oriC by PCR using oJW114/
oJW135, oJW115/oJW157, oJW112/oJW75 and oJW113/
oJW146. The stabilized inversion strain JDW713 was obtained
by transforming JDW704 with linearized plasmid pJW247. The
isogenic control was generated by transforming YB886 with
linearized plasmid pJW207. The left replichore inversion strain
JDW605 was created by screening progenies of MMB703 [40] by
PCR using primers oJW112/oJW388, and oJW113/oJW146.
Inversion of rRNA operons was performed similarly to the
method described in [50] and [88]. Briefly, two halves of the
neomycin resistance gene ( neo ) overlapping by 583 bp, were
inserted flanking the rrnIHG region in strain YB886 using plasmids
pJW260 and pJW261, respectively. The strain was plated on
kanamycin to select for cells in which a complete neo gene was
created by recombination between the two halves. The inversion
junctions were tested by PCR using oJW450/oJW442, and
oJW452/oJW436. Genomic coordinates [22] of the rrn inversion
are 159778 to 176408 in JDW860 and 154793 to 176408 in
Plasmid pJW247 was constructed by cloning sequences flanking
the inversion junction into the vector pUC18, on either side of the
cat gene. The sequences were amplified using oJW206/oJW360
and oJW210/oJW211, and cat was amplified from pGEMcat
[89] using oJW208/oJW209. Plasmid pJW207 was constructed
similarly, except that the PCR product of oJW204/oJW205 was
used instead of oJW206/oJW360. Plasmid pJW260 was construct-
ed by cloning sequences flanking the downstream rrn inversion
junction into the vector pUC18erm, on either side of the neo
fragment. The sequences were amplified using oJW434/oJW435
and oJW438/439, and neo was amplified from pBEST502 [90]
using oJW436/oJW437. Plasmid pJW261 was constructed by
cloning sequences flanking the upstream rrn inversion junction into
the vector pBEST501 [90], on either side of the neo fragment. The
sequences were amplified using oJW428/oJW429 and oJW432/
433, and cat was amplified from pGEMcat using oJW485/oJW486.
Microarrays and data analysis
Strains with the dnaB134ts allele [41,42] were grown in minimal
medium at 30uCtoOD
= 0.2. Cells were shifted to 45uC for
60 minutes to prevent new initiation and allow ongoing replication
to complete. The temperature was rapidly shifted down to 30uCto
allow synchronized initiation of replication, and the culture was
split into 2 flasks. 4 minutes after the down-shift, rifampicin was
added to one flask to 0.25mg/ml. Cells were collected 30 minutes
after the down-shift, mixed with an equal volume of 100% ice-cold
methanol and processed for microarray analysis as described [44].
Hybridization was performed according to the Agilent Oligo
aCGH protocol using custom 44K oligonucleotide Agilent
microarrays. Microarrays were scanned using a GenePix 4000B
scanner (Axon Instruments). Cy3 and Cy5 levels were quantified
using Agilent’s Feature Extraction software. Relative DNA content
ratio of Cy3 to Cy5 levels) was plotted against the gene
position on the chromosome, with the origin in the center and the
terminus at each end of the x-axis. For the inversion strains, the
genomic positions are rearranged to reflect genome reorganiza-
tion. For synchronized microarrays, the rolling average of gene
dosage ratios (log
) for every 200 consecutive positions was
calculated from the raw data, and plotted against the mid-point of
these positions.
Calculating replication rate as a function of gene-position
Genomic microarray profiles of cells grown in mid-exponential
phase were obtained by hybridizing against a synchronized
reference, such that the ratios were proportional to the actual
gene dosage. The data for the right replichore were analyzed
based on the following (for the left replichore, the equations are
identical except with negative signs):
1. During exponential growth, the total number of cells
increases exponentially with cell mass doubling. Define
as the
mass doubling time, t as the time of measurement, and
(t) as the
total number of cells at time t, then
2. Define x as the position of the gene, and the total gene dosage
of the population of cells at position x as f(x, t). There are 4.2Mbp
of nucleotides and over 10
cells, so we can approximate f as a
continuous function with continuous variables x and t, despite that
fact that each cell is undergoing discrete events including
replication initiation and cell division.
Assuming that cells are in steady state and their genomic profile
is time-invariant on a population basis, we have
NtzDtðÞ ðE2Þ
Combining E1 and E2, we have
3. The rate of replication fork progression is a function of
genome position x but not of time t during steady state growth.
Therefore we can define the replication rate at position x as v(x). By
this definition, for small Dx, we have:
Combining E3 and E4, we have
fx,tðÞ~fxzD x,tzDx
vxðÞðÞ~f (xzDx,t)|2
Define g~log
(f ), then
which can be rearranged as
v(x)~{T|(g(xzDx,t){g(x,t))=Dx, ðE5Þ
Head-On Transcription Disrupts Replication
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which can also be written in differential form as
1=v(x)~{T|Lg(x,t)=Lx ðE6Þ
Once we obtain g(x, t), equations E5 and E6 give a precise
definition of how to obtain v(x). Our microarray data show that, for
a fixed t, g(x, t) can be approximated by a piecewise linear function
over x. Let a discrete series (x
, x
) be the connecting points of
the piecewise linear function. If x falls between x
and x
, and
(t), where a
is independent of t, then we can
obtain v( x) as:
|T) ðE7Þ
Measurement of mutation rates
Measurement of rifampicin-resistance (rif
) mutation rates
was performed using the fluctuation test as described [49]. 50
parallel c ultures of 1–2 ml each were set up for each strain,
grown at 37u CtoOD
,0.5 and plated on minimal medium
containing rifampicin (5
mg/ml). Serial dilutions were also
plated on non-selective medium to count the numbe r of colony-
forming units. After incubation at 37u C for 36 hours, the
number of plates with no rif
colonies was counted. Rif
mutation rate was calculated using the P
method [49]. Results
from 2 or more independent experiments were averaged. Error
bars represent the range of data for n = 2, and the standard
error for n.2.
Live cell microscopy
The recA::(recA-gfp spc) allele [45] was used to replace the
endogenous recA gene in the inversion and control strains. The
recA-gfp strains were grown at 30uC in CAA and minimal medium.
= 0.2–0.6, 250 ml aliquots were labeled with the
membrane dye FM4-64 (0.05
mg/ml) and/or DAPI (0.1 mg/ml).
Cells were spotted onto thin agarose pads (1% agarose in 16
Spizizen’s salts) on multi-well slides, covered with a cover slip and
imaged in a Zeiss Axiovert microscope using a 1006 oil immersion
objective. Images were captured using a Hamamatsu Digital CCD
camera, and analyzed using the AxioVision software. The number
of cells with RecA-GFP foci or filaments relative to the total
number of cells in each image was counted.
SOS induction was monitored similarly in cells containing the
tagC-gfp reporter grown at 37uC in LB, CAA and minimal
medium. Data were analyzed by calculating SOS induction per
cell length.
For estimating the number of dead cells microscopically, the
Live/Dead BacLight Bacterial Viability Kit (Molecular Probes)
was used, in which live cells are labeled with SYTO9 (green
fluorescence, colored in cyan in Figure 3D and Figure 5H) and
dead cells with propidium iodide (red fluorescence). Data were
analyzed by calculating cell death per cell length.
For all microscopy experiments approximately 1000 cells were
counted for each strain and each growth condition. Results from 2
or more independent experiments were averaged. Error bars
represent the range of data for n = 2, and the standard error for
Measurement of number of colony-forming units (cfu)
Cells were grown at 37uCtoOD
,0.2–0.6 and serial
dilutions were plated on minimal medium. Colonies were counted
after 36 hours of incubation at 37uC. Data from 3 independent
experiments were averaged.
Mitomycin C sensitivity assay
Cells were grown in either LB or minimal medium to
= 0.3. 5 ml of 1:10 serial dilutions ranging from 10
were spotted correspondingly on LB, Min and plates
supplemented with mitomycin C to a concentration of
mg/ml. Plates were incubated overnight at 37uC and
photographed the next day. The experiment was repeated twice
and representative images are shown.
Supporting Information
Figure S1 Inversion of the oriC-proximalhalfoftheleft
replichore (HT-lef t) also leads to impedance of replication
fork progression. (A) HT-Left exhibits a strong growth defect
especially in rich media. Doubling times at 37uC in liquid
LB (red bars), minimal medium with (CAA, grey bars), or
without (Min, black bars) ca samino acids were calculated b y
measuring OD
. (B) Overlay of the asynchronous genomic
profiles of the HT strain grown in Min (blue), and CAA
(orange). Profiles were obtained from asynchronous cultures
grown at 37u CtoOD
, 0.5. Average gene dosage ratios
)areplottedrelativetogenepositions adjusted according
to known deletions of the background strain JH642 [6]. (C)
Ratios of gene dosa ge (log
Left strain, calculated similarly to Figure 2C. Green arrow:
position of the inve rted rRNA operon rrnB; orange arrows:
positions o f co-directional rRNA operons; grey shaded
region: inversion.
Found at: doi:10.1371/journal.pgen.1000810.s001 (0.95 MB TIF)
Table S1 Comparison between the observed and expected
fitness of the HT and UR mutants in the indicated growth media,
under the multiplicative null model.
Found at: doi:10.1371/journal.pgen.1000810.s002 (0.02 MB
Table S2 Relative fitness of the HT strain in the indicated
growth media (W
Found at: doi:10.1371/journal.pgen.1000810.s003 (0.03 MB
Table S3 Strains.
Found at: doi:10.1371/journal.pgen.1000810.s004 (0.05 MB
Table S4 Primers.
Found at: doi:10.1371/journal.pgen.1000810.s005 (0.05 MB
Text S1 Supplemental Materials and Methods.
Found at: doi:10.1371/journal.pgen.1000810.s006 (0.03 MB
We thank Allison Kriel for technical assistance; Alycia Bittner, Elicia
Grace, Phil Hastings, Xiangwe He, Elizabeth Ostrowski, Susan Rosenberg,
and Lizhao Zhang for helpful discussions; and the anonymous reviewers for
critical comments.
Author Contributions
Conceived and designed the experiments: AS JDW. Performed the
experiments: AS AT. Analyzed the data: AS JDW. Contributed reagents/
materials/analysis tools: DMM JDW. Wrote the paper: AS JDW.
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Head-On Transcription Disrupts Replication
PLoS Genetics | 14 January 2010 | Volume 6 | Issue 1 | e1000810
    • "While we do not have any direct indication about the speed of individual replication forks, this result suggests that the fork coming from oriC and going in the native direction must be, overall, slower than the fork coming from oriZ replicating in the wrong orientation from the termination area onwards. The strongest deviations of the replication profiles were consistently observed at the rrn operons [10,13,[46][47][48]. However, all rrn operons are relatively close to oriC. "
    [Show abstract] [Hide abstract] ABSTRACT: Duplication of bacterial chromosomes is initiated via the assembly of two replication forks at a single defined origin. Forks proceed bi-directionally until they fuse in a specialised termination area opposite the origin. This area is flanked by polar replication fork pause sites that allow forks to enter but not to leave. The precise function of this replication fork trap has remained enigmatic, as no obvious phenotypes have been associated with its inactivation. However, the fork trap becomes a serious problem to cells if the second fork is stalled at an impediment, as replication cannot be completed, suggesting that a significant evolu¬tionary advantage for maintaining this chromosomal arrangement must exist. Recently we demonstrated that head-on fusion of replication forks can trigger over-replication of the chromosome. This over-replication is normally prevented by a number of proteins including RecG helicase and 3’ exonucleases. However, even in the absence of these proteins it can be safely contained within the replication fork trap, highlighting that multiple systems might be involved in coordinating replication fork fusions. Here we discuss whether considering the problems associated with head-on replication fork fusion events helps us to better understand the important role of the replication fork trap in cellular metabolism.
    Full-text · Article · Jul 2016
    • "The extra forks established in the irradiated cells would further disrupt the replichore arrangement of the chromosome, leading to unscheduled amplification of the replicated areas and increasing the incidence of fork fusions that might trigger further over-replication of the DNA (Fig. 4b, c). It would also increase conflicts with transcription, which itself is likely to have pathological consequences, especially at highly transcribed genes such as rrn operons (Trautinger et al. 2005; Wang et al. 2007; Guy et al. 2009; Boubakri et al. 2010; Srivatsan et al. 2010; Atkinson et al. 2011; De Septenville et al. 2012; Merrikh et al. 2012; Dimude et al. 2015; Ivanova et al. 2015). Recombination initiated via RecBCD-and RecFOR-mediated loading of RecA might further compound the problem by linking chromosomes and/or partially replicated areas together. "
    [Show abstract] [Hide abstract] ABSTRACT: The RecG protein of Escherichia coli is a double-stranded DNA translocase that unwinds a variety of branched substrates in vitro. Although initially associated with homologous recombination and DNA repair, studies of cells lacking RecG over the past 25 years have led to the suggestion that the protein might be multi-functional and associated with a number of additional cellular processes, including initiation of origin-independent DNA replication, the rescue of stalled or damaged replication forks, replication restart, stationary phase or stress-induced 'adaptive' mutations and most recently, naïve adaptation in CRISPR-Cas immunity. Here we discuss the possibility that many of the phenotypes of recG mutant cells that have led to this conclusion may stem from a single defect, namely the failure to prevent re-replication of the chromosome. We also present data indicating that this failure does indeed contribute substantially to the much-reduced recovery of recombinants in conjugational crosses with strains lacking both RecG and the RuvABC Holliday junction resolvase.
    Full-text · Article · Feb 2016
    • "Refactoring genetic pathways provides an opportunity to increase modularity [163], but cryptic regulation mechanisms and polar effects make it difficult to design a de novo pathway architecture with optimal activity [164,180]. Finally, while genome-scale design rules will continue to be discovered, we already know that it is important to balance the size of replichores in circular chromosomes [166], to co-orient transcription of essential operons with translation [168], to preserve sequences involved in DNA structure [169] and repair [170], and to consider how chromosome size impacts its structural integrity [167].  Barriers to changing the genetic code  biochemical barriers  biological knowledge (ability to design functional genomes)  biotechnology (ability to create the genomes)  Approach to engineer genomes with new biological functions Fig. 1Fig. "
    [Show abstract] [Hide abstract] ABSTRACT: Withstanding 3.5 billion years of genetic drift, the canonical genetic code remains such a fundamental foundation for the complexity of life that it is highly conserved across all three phylogenetic domains. Genome engineering technologies are now making it possible to rationally change the genetic code, offering resistance to viruses, genetic isolation from horizontal gene transfer, and prevention of environmental escape by genetically modified organisms. We discuss the biochemical, genetic, and technological challenges that must be overcome in order to engineer the genetic code.
    Full-text · Article · Sep 2015
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