Journal of General Virology (2015), 96, 1957–1968
DOI 10.1099/vir.0.000102
Defects in RNA polyadenylation impair both
lysogenization by and lytic development of Shiga
toxin-converting bacteriophages
Dariusz Nowicki,13 Sylwia Bloch,13 Bożena Nejman-Faleńczyk,1
Agnieszka Szalewska-Pałasz,1 Alicja We˛grzyn2 and Grzegorz We˛grzyn1
Correspondence
1
Grzegorz We˛grzyn
2
[email protected]
Received 2 October 2014
Accepted 19 February 2015
Department of Molecular Biology, University of Gdańsk, Wita Stwosza 59, 80-308 Gdańsk, Poland
Laboratory of Molecular Biology (affiliated with the University of Gdańsk), Institute of Biochemistry
and Biophysics, Polish Academy of Sciences, Wita Stwosza 59, 80-308 Gdańsk, Poland
In Escherichia coli, the major poly(A) polymerase (PAP I) is encoded by the pcnB gene. In this
report, a significant impairment of lysogenization by Shiga toxin-converting (Stx) bacteriophages
(W24B, 933W, P22, P27 and P32) is demonstrated in host cells with a mutant pcnB gene.
Moreover, lytic development of these phages after both infection and prophage induction was
significantly less efficient in the pcnB mutant than in the WT host. The increase in DNA
accumulation of the Stx phages was lower under conditions of defective RNA polyadenylation.
Although shortly after prophage induction, the levels of mRNAs of most phage-borne early genes
were higher in the pcnB mutant, at subsequent phases of the lytic development, a drastically
decreased abundance of certain mRNAs, including those derived from the N, O and Q genes,
was observed in PAP I-deficient cells. All of these effects observed in the pcnB cells were
significantly more strongly pronounced in the Stx phages than in bacteriophage l. Abundance of
mRNA derived from the pcnB gene was drastically increased shortly (20 min) after prophage
induction by mitomycin C and decreased after the next 20 min, while no such changes were
observed in non-lysogenic cells treated with this antibiotic. This prophage induction-dependent
transient increase in pcnB transcript may explain the polyadenylation-driven regulation of phage
gene expression.
INTRODUCTION
Shiga toxin-converting bacteriophages (or Stx phages) are
classified as lambdoid viruses due to organization of their
genomes, which resembles that of bacteriophage l (Allison,
2007; Łoś et al., 2011). Although there are also homologies
between sequences of some crucial regulatory genes of Stx
phages and l, they rarely share a common virion structure.
Nevertheless, similarities in regulatory mechanisms of viral
development are also evident, including the lysis-versuslysogenization decision, control of phage gene expression
and phage DNA replication; though differences in important details were also reported (Ptashne, 2004; We˛grzyn &
We˛grzyn, 2005; We˛grzyn et al., 2012; Allison, 2007; Nejman
et al., 2009; Łoś et al., 2011; Riley et al., 2012). What clearly
distinguishes all Stx phages from l is the presence of stx
genes, coding for Shiga toxins, in their genomes.
Escherichia coli strains become potentially pathogenic to
humans when they are lysogenized with an Stx phage. This
arises from production of Shiga toxins, which are
3These authors contributed equally to this work.
000102 G 2015 The Authors Printed in Great Britain
dangerous to humans, due to the induction of Stx prophages and during lytic development of the viruses. E. coli
strains bearing such prophages (called STEC for Shiga
toxin-producing E. coli) may cause serious diseases (Gyles,
2007; Łoś et al., 2012). Particularly, a subset of STEC strains,
called enterohaemorrhagic E. coli (EHEC) are especially
dangerous when infecting humans (Hunt, 2010; Mauro &
Koudelka, 2011). The severity of medical problems caused by
STEC strains can be exemplified by the outbreak that
occurred in Germany in 2011. Its fatal results, summarized
as over 4000 symptomatic infections and over 50 cases of
death, besides confirming the importance of the problem,
also underlined our incomplete knowledge about regulatory
mechanisms operating during development of the Stx phages
(Mellmann et al., 2011; Beutin & Martin, 2012; Bloch et al.,
2012; Karch et al., 2012; Werber et al., 2012).
RNA polyadenylation is a post-transcriptional modification of various ribonucleic acids, which proceeds as the
addition of adenosine (A) residues to the 39 ends of RNA
strands, independently of any template. This process occurs
in both eukaryotic and prokaryotic cells, while having
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1957
D. Nowicki and others
different consequences in these two groups of biological
entities (see Mohanty & Kushner, 2011 for a review). In
bacteria, it is generally assumed that polyadenylated RNA
molecules are more susceptible to degradation, thus addition of poly(A) tails destabilizes various transcripts. However,
there are also examples of RNA stabilization upon polyadenylation, which makes the control of RNA metabolism
even more complicated (Mohanty & Kushner, 2011; Régnier
& Hajnsdorf, 2013). Moreover, although it was initially
believed that RNA polyadenylation in bacterial cells concerns
mainly (if not exclusively) mRNA and small regulatory RNA
(sRNA) species, recent reports indicate that this kind of
modification of tRNA molecules may have an important
impact on cell physiology (Mohanty et al., 2012; Mohanty &
Kushner, 2013). In E. coli, the major enzyme responsible for
RNA polyadenylation is poly(A) polymerase I (PAP I),
encoded by the pcnB gene (Cao & Sarkar, 1992; Mohanty &
Kushner, 2011). Transcripts derived from over 90 % of ORFs
undergo polyadenylation by this enzyme to some extent
(Mohanty & Kushner, 2000, 2006).
There is a surprisingly low number of published reports
concerning the role of RNA polyadenylation in development of bacteriophages. The most important discoveries
in this field are summarized below. Short stretches of A or U
residues were detected in the RNA genome of bacteriophage
MS2, however, their biological role is unclear (Klovins et al.,
1997). Polyadenylation of different mRNAs of bacteriophage
T7 was reported, but it appeared that this modification
occurred after nucleolytic cleavage of the primary transcripts
(Johnson et al., 1998). Similar results were described after
analysis of mRNAs encoded by filamentous bacteriophage f1,
where polyadenylation has been suggested to occur at the
later stages of transcript degradation (Goodrich & Steege,
1999). Contrary to reports on T7- and f1-derived transcripts,
no significant polyadenylation of mRNAs of phage T4 could
be found (Yonesaki, 2002). One of the few published
examples of the involvement of RNA polyadenylation in
the development of bacteriophages is the requirement of this
biochemical reaction for maturation of the CI RNA, a small
regulatory transcript of phage P4 (Briani et al., 2002).
Another example is polyadenylation of oop RNA, an antisense
transcript to mRNA of the cII gene of bacteriophage l
(Wróbel et al., 1998). Polyadenylated oop RNA was degraded
more rapidly than its unmodified equivalent (SzalewskaPałasz et al., 1998). However, the only observed effect of the
pcnB gene dysfunction was decreased efficiency of lysogenization by phage l, which was ascribed to more efficient
inhibition of expression of the cII gene by the more stable
antisense RNA (Wróbel et al., 1998).
Because the field studying the role of polyadenylation in the
development of bacteriophages is clearly underexplored and
in the light of mounting evidence for important regulatory
functions of this RNA modification (Mohanty & Kushner,
2011; Régnier & Hajnsdorf, 2013), we aimed to investigate the
effects of mutating the pcnB gene, coding for the major E. coli
poly(A) polymerase, on lytic and lysogenic modes of
development of Shiga toxin-converting bacteriophages.
1958
RESULTS
Impaired lysogenization of the pcnB mutant by
Stx phages
The only effect of severely decreased RNA polyadenylation,
due to mutation of the pcnB gene, on lambdoid bacteriophage
development reported to date was a decreased efficiency of
lysogenization by bacteriophage l (Wróbel et al., 1998).
Therefore, we have measured this parameter for a series of Stx
bacteriophages (W24B, 933W, P22, P27 and P32) in WT and
DpcnB :: kan strains of E. coli. We confirmed that bacteriophage l lysogenizes the mutant host less efficiently and found
that similar but significantly more pronounced effects can be
observed for all tested Stx phages (Table 1). In fact, in all our
experimental systems with different m.o.i., which is known to
influence efficiency of lysogenization (see We˛grzyn &
We˛grzyn, 2005 for a review), formation of lysogenic E. coli
DpcnB :: kan cells was either extremely rare or impossible to
detect (Table 1). Therefore, we concluded that lysogenization
by Stx phages is severely impaired by defects in RNA
polyadenylation.
Decreased efficiency of Stx phage lytic
development in the pcnB mutant after infection or
prophage induction
Previous studies suggested that lytic development of
bacteriophage l is only slightly or minimally, if at all,
affected by pcnB dysfunction (Wróbel et al., 1998). Here,
we have confirmed only minor (less than twofold, on
average) effects of the DpcnB :: kan mutation on the kinetics
and efficiency of formation of l phage progeny, either after
infection of the host cells (Fig. 1) or following prophage
induction with mitomycin C (Fig. 2). In both kinds of
experiments, burst sizes of l were in the range of a few
hundred, as expected. However, while the differences
between WT and DpcnB :: kan hosts reached statistical
significance in experiments with infection (Fig. 1), no such
significance was detected after prophage induction (Fig. 2).
On the other hand, significant differences between Stx
phage lytic development in WT and DpcnB :: kan hosts were
noted, irrespective of the nature of the initiation of phage
lytic development (infection or prophage induction) and
this was true for all tested Shiga toxin-converting phages
(Figs 1 and 2). In all cases, the phages produced five to ten
times less progeny in the pcnB mutant than in the WT host.
Thus, not only lysogenization, but also lytic development
of Stx phages was negatively affected by dysfunction of the
pcnB gene. We conclude that the RNA polyadenylation
process is important in both developmental pathways of
these viruses.
Phage DNA synthesis is negatively influenced by
dysfunction of the pcnB gene
To identify the process(es) in phage development affected
by impaired RNA polyadenylation, we tested the kinetics of
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Journal of General Virology 96
RNA polyadenylation in development of lambdoid phages
Table 1. Efficiency of lysogenization of E. coli WT (MG1655) and otherwise isogenic DpcnB :: kan strains with lambdoid
bacteriophages
Results are presented as mean ±SD from three independent experiments. In all cases, differences between WT and DpcnB :: kan strains were
significant (P,0.05 in Student’s t-test).
Phage
Efficiency of lysogenization
WT
m.o.i.51
lpapa
W24B
933W
P22
P27
P32
0.412±0.024
0.381±0.031
0.400±0.028
0.355±0.016
0.324±0.024
0.250±0.025
DpcnB :: kan
m.o.i.55
m.o.i.510
m.o.i.51
m.o.i.55
m.o.i.510
0.553±0.019
0.467±0.024
0.560±0.054
0.367±0.028
0.480±0.034
0.270±0.018
0.733±0.054
0.662±0.035
0.784±0.044
0.554±0.045
0.640±0.029
0.620±0.037
0.010±0.010
0.001±0.001
0.001±0.001
,0.001*
,0.001*
0.002±0.001
0.022±0.004
0.002±0.001
0.001±0.001
0.001±0.001
0.001±0.001
0.002±0.001
0.040±0.020
0.005±0.001
0.003±0.001
0.002±0.001
0.002±0.001
0.003±0.002
*No lysogens could be detected.
the increase in phage DNA amount (which reflects phage
DNA synthesis) in the host cells. The rationale for testing
this parameter stems from previous observations that
indicated sensitivity of DNA replication of different Stx
phages to environmental or physiological stresses, like
starvation, oxidative stress or temperature variation (Łoś
et al., 2009; Nejman-Faleńczyk et al., 2012; Nowicki et al.,
2013).
Bacteriophage l and three Stx phages (W24B, 933W and
P27) were chosen to test DNA synthesis [P22 and P32 were
omitted since replication regions of their genomes are very
similar to that of P27 (Nejman et al., 2009)]. Following
prophage induction, the kinetics of increases in phage
DNA was monitored in WT and DpcnB :: kan hosts. We
found that although mutation of the pcnB gene influenced
synthesis of phage l DNA only slightly (up to twofold
differences in the amount of phage DNA which, nevertheless, reached statistical significance), considerable impairment (fivefold to tenfold) in the increase in amount
of Stx phages’ genome-specific sequences was evident
(Fig. 3). Again, all Stx phages behave similarly in this
experiment. We concluded that RNA polyadenylation
indirectly influenced the efficiency of DNA synthesis in
the Stx phages.
Expression of phage genes in WT host and the
pcnB mutant
To learn about the molecular mechanism(s) of polyadenylation-mediated control of development of Stx phages,
expression of selected viral genes was investigated after
prophage induction by measuring the levels of particular
transcripts. The following genes were chosen due to their
crucial roles in either lytic or lysogenic development: xis,
cIII, N, cI, cro, cII, oop, O, Q and R. Among the tested
phages only genomes of l, W24B and 933W have been
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completely sequenced and published to date, therefore the
analysis of viral gene expression was possible only in these
cases. As most of the tested genes (xis, cIII, N, cI, cro, cII,
oop, O) are expressed shorty after infection or prophage
induction (so-called early genes) samples for analysis were
withdrawn at time points before the extensive DNA
replication and before progeny virions were assembled.
Results of these analyses are presented as levels of the tested
RNAs at particular times after prophage induction relative
to that before induction [time zero; these values were used
as calibrators in reverse transcriptase (RT)-quantitative
PCR (qPCR)].
A relatively short time after induction with mitomycin C,
the amount of all tested transcripts of phages l and W24B,
and almost all transcripts of phage 933W, was significantly
increased in the pcnB mutant relative to the WT host (Fig.
4). However, opposite results were obtained at later time
points in which the abundance of most of the transcripts
was significantly reduced in polyadenylation-deficient cells.
These differences included genes crucial for lytic development, N and O, coding for an antitermination protein
(responsible for positive regulation of expression of
delayed-early genes required for lytic development) and
the replication initiator, respectively. Importantly, the
levels of N and O transcripts were drastically decreased in
the pcnB mutant after induction of W24B and 933W
prophages, while the differences were moderate in experiments with bacteriophage l (Fig. 4). Such a pattern
corresponds to effects of the pcnB gene mutation on lytic
development of the tested phages, significantly pronounced
for the Stx phages and small or negligible for l (compare
Fig. 4 with Figs 2 and 3). Therefore, we conclude that the
impairment of lytic development of Stx phages in host cells
defective in RNA polyadenylation arises from significantly
lower abundance of transcripts of genes coding for proteins
crucial for this mode of phage growth.
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1959
Fig. 1. Lytic development of bacteriophages lpapa (a), W24B (b), 933W (c), P22 (d), P27 (e) and P32 (f) after infection of otherwise isogenic E. coli WT (open symbols) or
DpcnB :: kan (closed symbols) host cells at time zero. Results are presented as burst size (phages per infected cell). Mean values from three independent experiments with
error bars indicating SD (note that in some cases, the bars are smaller than the size of symbols) are shown. Significant differences (P,0.05; Student’s t-test) were detected
between WT and DpcnB :: kan strains in all experiments with the last five points indicated.
150
Time (min)
100
100
50
150
Time (min)
100
100
50
100
50
100
50
0
0.001
0.01
0.1
10
1
Burst size
(p.f.u. per cell)
100
1000
(a)
1960
Time (min)
150
0.001
0.01
λ papa
0.1
1
10
WT
ΔpcnB
(b)
0
Time (min)
150
0.001
0.01
φ 24B
WT
ΔpcnB
0.1
0
Time (min)
150
0.001
0.01
933W
WT
ΔpcnB
0.1
0
50
WT
ΔpcnB
0.001
1
1
1
0.01
10
10
10
P22
100
100
100
100
0.1
1000
1000
1000
(e)
(d)
(f)
1000
0
Time (min)
150
0.001
0.01
P27
0.1
1
WT
ΔpcnB
10
100
1000
(f)
0
50
P32
WT
ΔpcnB
D. Nowicki and others
Growth rate of lysogenic cells and expression of
the pcnB gene after prophage induction
We asked how can defects in RNA polyadenylation affect
expression of many viral genes in a coordinated manner
which is, however, different at various time points after
prophage induction? It has been demonstrated that the pcnB
gene is expressed at low levels in exponentially growing E. coli
cells (Binns & Masters, 2002; Mohanty et al., 2004), however,
the level of the PAP I protein is growth-rate-dependent,
being significantly higher in slowly growing cells (Jasiecki &
We˛grzyn, 2003). This regulation of pcnB expression during
slow growth is apparently due to the presence of a complicated promoter system upstream of this gene, composed
of three s70- and two sS-dependent promoters (Jasiecki &
We˛grzyn, 2006; Nadratowska-Wesołowska et al., 2010).
Therefore, we have measured growth rates of lysogenic and
non-lysogenic WT and DpcnB :: kan cells under growth
conditions employed in this work. The average generation
time of non-induced lysogenic strains was 28±1 min (Table
2, note that all lysogens gave very similar results irrespective
of the kind of the prophage), while shortly after prophage
induction, the bacterial growth slowed down and the average
generation time measured between 0 and 40 min after
addition of mitomycin C was 35±2 min (Table 2). At later
times (over 40 min) after prophage induction, bacterial
growth temporarily resumed (Table 2). The average
generation time of non-lysogenic strains was 25±1 (there
was no difference between WT and DpcnB :: kan bacteria) and
after the addition of 1 mg, microgram mitomycin C ml21 the
generation time (measured between 0 and 40 min after the
addition) did not change significantly (Table 2). Apparently,
the concentration of mitomycin C employed in these
experiments was too low to cause bacterial growth inhibition,
thus, the effects observed in lysogenic strains were due to
prophage induction.
To test whether the decreased bacterial growth rate after
prophage induction, likely arising from the induction-related
stress conditions, might cause enhanced expression of the pcnB
gene, we measured the transcript levels of this gene at various
times after prophage induction in WT cells lysogenic for l,
W24B or 933W, as well as in non-lysogenic bacteria. pcnB
transcript levels did not change significantly in non-lysogenic
cells after addition of mitomycin C (Fig. 5). However, a
dramatic increase in pcnB transcript levels was observed for all
tested lysogens 20 min after prophage induction. This increase
was transient, as at 40 min and later times after the addition of
mitomycin C the abundance of this transcript dropped to the
level measured in non-lysogenic cells (Fig. 5).
We conclude that prophage induction created conditions
that resulted in a significant increase in pcnB gene expression.
This may explain the mechanism of RNA polyadenylationdependent regulation of phage genes under such conditions.
DISCUSSION
In this work, the effects of defective RNA polyadenylation,
caused by dysfunction of the pcnB gene coding for the
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Journal of General Virology 96
RNA polyadenylation in development of lambdoid phages
(b)
(a)
1010
Relative phage titre p.f.u.
per A600 unit
1010
108
108
106
106
WT
DpcnB
104
102
100
λ papa
WT
DpcnB
104
102
φ24B
100
0
100
200
Time (min)
0
300
(c)
100
200
Time (min)
300
(d)
1010
Relative phage titre p.f.u.
per A600 unit
1010
108
108
106
106
WT
DpcnB
104
WT
DpcnB
104
102
102
933W
100
0
100
200
Time (min)
300
(e)
P22
100
0
100
200
Time (min)
300
(f)
1010
Relative phage titre p.f.u.
per A600 unit
1010
108
108
106
106
WT
DpcnB
104
WT
DpcnB
104
102
102
P27
P32
100
100
0
100
200
Time (min)
300
0
100
200
Time (min)
major poly(A) polymerase [poly(A) polymerase I, or PAP
I], on lysogenization and lytic growth of several Shiga
toxin-converting phages were investigated. Similar to
bacteriophage l that was previously reported to lysogenize
E. coli pcnB mutant cells less efficiently than WT hosts
(Wróbel et al., 1998), we found the formation of prophages
by Stx phages to be impaired in bacteria devoid of poly(A)
polymerase I (Table 1). However, defects in polyadenylation caused significantly more pronounced impairment of
lysogenization by Stx bacteriophages than by l. It was
suggested that less effective lysoganization of pcnB hosts by
l is due to defective polyadenylation of oop RNA, an
antisense transcript acting as a negative regulator of expression of the cII gene, coding for the positive regulator of the
pE promoter, necessary for production of the cI repressor
and establishment of the prophage state (Wróbel et al.,
1998). Since polyadenylated oop RNA is rapidly degraded
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300
Fig. 2. Lytic development of bacteriophages
lpapa (a), W24B (b), 933W (c), P22 (d), P27
(e) and P32 (f) after prophage induction
with 1 mg mitomycin C ml”1 at time zero in
otherwise isogenic E. coli WT (open symbols)
or DpcnB :: kan (closed symbols) host cells.
Results are presented as number of phages
(p.f.u.) per A600 unit of the culture. Mean values
from three independent experiments with error
bars indicating SD (note that in some cases,
the bars are smaller than the size of symbols)
are shown. Significant differences (P,0.05)
between WT and DpcnB :: kan strains were
detected in all Stx phages, but not in l.
in E. coli cells (Szalewska-Pałasz et al., 1998), impaired
modification of this transcript would result in its stabilization, more effective inhibition of cII expression, less
efficient production of the cI repressor and difficulties in
lysogenization. Such a mechanism could also operate in Stx
phages as oop RNA has been reported for W24B, 933W and
some other lambdoid phages (Nejman-Faleńczyk et al.,
2015). However, if modification of oop RNA is the only
cause of polyadenylation-facilitated lysogenization, it must
be more effective than in l.
In bacteriophage l, Wróbel et al. (1998) found that
lysogenization was the only major process of phage
development considerably impaired in PAP I-deficient
cells. In contrast, lytic growth of all Stx phages tested here
and measured as efficiency of production of phage
progeny, was significantly less effective in the pcnB mutant.
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1961
D. Nowicki and others
Realtive phage DNA content (ng per A600 unit)
(a)
(b)
40
40
w 24B
λ papa
20
20
WT
WT
DpcnB
DpcnB
0
0
0
60
120
180
240
60
120
180
240
Time (min)
(c)
Realtive phage DNA content (ng per A600 unit)
0
Time (min)
(d)
40
40
933W
20
P27
20
WT
WT
DpcnB
DpcnB
0
0
0
60
120
Time (min)
180
240
0
60
120
Time (min)
This was the case when the developmental mode had been
initiated by infection or prophage induction, suggesting
that a common mechanism might operate to slow down
the production of phage progeny.
To learn about the specific mechanism of polyadenylationcontrolled Stx phage development, expression patterns of
phage genes crucial for the regulation of virus development
were determined after prophage induction in WT and pcnB
hosts. Levels of mRNAs of xis, cIII, N, cI, cro, cII, O, Q and
R genes, as well as levels of oop antisense RNA, were
determined by RT-qPCR. Note that after prophage
induction xis, cIII and N are transcribed from the pL
promoter (after excision of the prophage and circularization of phage DNA, xis may be expressed from another
promoter, pI, this promoter operates in l but not in some
other lambdoid phages, like W24B), cI is expressed from its
own promoter, pM (though under these conditions this
promoter is repressed by Cro and low levels of cI expression was detected), cro, cII, O and Q mRNAs form a
multicistronic transcript from pR, the R gene is under
control of the pR’ promoter and oop RNA is a short
transcript originating from a promoter located in the cII–O
region but oriented in the opposite direction relative to pR
(see We˛grzyn & We˛grzyn, 2005 for a review). Interestingly,
1962
180
240
Fig. 3. Levels of DNA of bacteriophages
lpapa (a), W24B (b), 933W (c) and P27 (d)
either without prophage induction (circles) or
after prophage induction with 1 mg mitomycin
C ml”1 at time zero (squares) in otherwise
isogenic E. coli pcnB+ (open symbols) or
DpcnB :: kan (closed symbols) host cells.
Results are presented as mean values from
three independent experiments with error bars
indicating SD (note that in some cases, the
bars are smaller than the size of symbols).
Control experiments with strains lacking a
prophage resulted in detection of no phage
DNA. Significant differences (P,0.05) between WT and DpcnB :: kan strains were
detected for all phages.
although several transcripts were tested, some common
features could be established for all investigated phages and
most of the genes assessed. Namely, the majority of tested
RNAs were significantly more abundant in the pcnB mutant
relative to WT bacteria (at a relatively short time after
prophage induction), while PAP I-deficient cells had lower
levels of transcripts at later times. Importantly, the differences
were severe in the Stx phages, revealing impaired lytic growth
in the pcnB host, but moderate in phage l where lytic
development was only slightly affected. This strongly suggests
that changes in the expression of crucial genes of the Stx
bacteriophages are responsible for less efficient lytic growth of
these viruses under conditions of deficient RNA polyadenylation. Such a hypothesis is supported by the fact that levels of
the N and O gene transcripts are significantly less abundant in
the pcnB mutant cells at later times after prophage induction
and these genes code for an antitermination protein (responsible for the positive regulation of other genes required for
lytic development) and the replication initiator (a protein
necessary for phage DNA replication initiation), respectively.
In fact, we found that synthesis of all tested Stx phages’
DNA after prophage induction was significantly less in
pcnB cells than in WT hosts, indicating that this process
can be at least one of the targets of the regulation. Most
probably, this effect was indirect, possibly due to impaired
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Journal of General Virology 96
RNA polyadenylation in development of lambdoid phages
(b)
xis cIII N
cI cro cII oop O
Q
R
λ - 80 min
wt
ΔpcnB
xis cIII N
cI cro cII oop O
Q
R
1300
1200
1100
1000
900
800
700
600
500
400
300
200
100
0
xis cIII N
1400
1300
1200
1100
1000
900
800
700
600
500
400
300
200
100
0
cI cro cII oop O
Q
R
φ24B - 60 min
wt
ΔpcnB
xis cIII N
cI cro cII oop O
Q
R
φ24B - 80 min
wt
ΔpcnB
xis cIII N
cI cro cII oop O
Q
Relative quantification results
(E-Method)
Relative quantification results
(E-Method)
R
λ - 60 min
wt
ΔpcnB
Relative quantification results
(E-Method)
1400
1300
1200
1100
1000
900
800
700
600
500
400
300
200
100
0
Q
φ24B - 40 min
wt
ΔpcnB
R
110
100
90
80
70
60
50
40
30
20
10
0
1300
1200
1100
1000
900
800
700
600
500
400
300
200
100
0
933W - 40 min
wt
ΔpcnB
xis cIII N
1400
1300
1200
1100
1000
900
800
700
600
500
400
300
200
100
0
cI cro cII oop O
Q
R
933W - 60 min
wt
ΔpcnB
Relative quantification results
(E-Method)
cI cro cII oop O
(c)
110
100
90
80
70
60
50
40
30
20
10
0
xis cIII N
cI cro cII oop O
Q
R
933W - 80 min
wt
ΔpcnB
Relative quantification result
(E-Method)
xis cIII N
Relative quantification results
(E-Method)
1300
1200
1100
1000
900
800
700
600
500
400
300
200
100
0
λ - 40 min
wt
ΔpcnB
Relative quantification results
(E-Method)
110
100
90
80
70
60
50
40
30
20
10
0
Relative quantification results
(E-Method)
Relative quantification results
(E-Method)
(a)
xis cIII N
cI cro cII oop O
Q
R
Fig. 4. Levels of transcripts of the indicated genes of l, W24B and 933W bacteriophages, assessed by RT-qPCR analysis, at
various times (40, 60 and 80 min) after prophage induction with 1 mg mitomycin C ml”1, in otherwise isogenic E. coli WT
(open columns) or DpcnB :: kan (closed columns) host cells. The sample at the time point ‘zero’ was used as a calibrator (all
presented quantities are expressed as n-fold difference relative to the calibrator minus an untreated control sample; in fact,
values for untreated samples were close to zero, and all calibrator values were similar and low). Statistical analysis (Student’s
t-test) was performed for results from each time point and indicated significant differences (P,0.05) between expression of
most genes in WT (pcnB+) and DpcnB :: kan hosts for tested phages. For a clear presentation, asterisks representing P
values are not marked. Differences in levels of expression of the following genes of tested phages (WT versus DpcnB :: kan
hosts) did not reach significance in the t-test (P.0.05): l papa at 60 min: cI, O, Q, R; at 80 min: cI, cII, Q; w24B at 40 min: cI;
at 60 min: cI; at 80 min: cI, R; 933W at 60 min: cI, cro, xis, Q, R. Results are presented as mean values from three
independent experiments with error bars indicating SD.
expression of the O gene. Importantly, replication of
phage l DNA was only slightly slowed down in PAP Ideficient cells and abundance of mRNA for the O gene
was similar in WT and DpcnB strains, corroborating the above assumptions.
Investigating the mechanism by which defects in RNA
polyadenylation affect expression of many viral genes in a
coordinated manner at different times after prophage
induction, we have found that growth rates of lysogenic
strains (both WT and DpcnB) temporarily decreased
following induction with mitomycin C. This phenomenon
was not observed in non-lysogenic bacteria. Since expreshttp://vir.sgmjournals.org
sion of pcnB is growth-rate-dependent, being significantly
higher in slow growing cells (Jasiecki & We˛grzyn, 2003), we
supposed that a transient production of PAP I may occur
in lysogenic cells after prophage induction. Testing this
possibility, we found the level of the pcnB mRNA increased
dramatically shortly after prophage induction (up to 20 min)
in WT bacteria, while at the same time this increase
was transient, as it was abolished during the next 20 min.
Again, such a phenomenon was not observed in nonlysogenic cells.
Based on these results, we propose the following hypothesis:
decreased bacterial growth rate, arising from the induction-
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1963
D. Nowicki and others
related stress conditions, enhances pcnB expression and thus,
more efficient polyadenylation of various transcripts. In fact,
it was demonstrated that transcripts derived from over 90 %
of ORFs are polyadenylated in E. coli (O’Hara et al., 1995;
Mohanty & Kushner, 2006). However, among all RNA
molecules appearing due to transcription of a particular gene,
only some are polyadenylated, thus, a fraction as small as 2 %
of total RNA molecules undergoes this modification under
standard laboratory growth conditions (Sarkar et al., 1978).
Therefore, enhanced polyadenylation (occurring shortly after
prophage induction) can lead to more efficient degradation
of transcripts in the WT strain. On the other hand,
this cannot occur in the DpcnB strain, resulting in a higher
abundance of a large fraction of tested (phage-derived)
transcripts in the mutant host relative to WT bacteria. At later
times (over 40 min) after prophage induction, the level of
PAP I in WT cells decreased, thus, most RNA molecules
would not be polyadenylated, while polyadenylation of
specific RNAs could be highly predominant over low level
polyadenylation of many transcripts. There are both
experimental results and theoretical analyses suggesting that
such a specificity or preference might be characteristic to
tRNA and sRNA species (Mohanty et al., 2012; Mohanty &
Kushner, 2013; Régnier & Hajnsdorf, 2013). If putative
sRNAs, involved in the control of phage gene expression,
were preferentially polyadenylated their stability could be
affected resulting in weak negative effects on the abundance
of phage transcripts in WT cells but in more effective
degradation in pcnB mutants. Although oop RNA is the only
polyadenylated short regulatory transcript specifically affecting lambdoid phage development reported to date (Wróbel
et al., 1998), results from a recent study suggest that phageencoded sRNAs may be significantly more abundant.
Namely, global genomic analysis indicated that one of the
prophages present in EHEC encodes 55 potential candidates
for such RNA species (Tree et al., 2014).
METHODS
Bacterial strains and bacteriophages. E. coli MG1655 strain
(Jensen, 1993) and its isogenic DpcnB :: kan derivative (Masters et al.,
1993; Wróbel et al., 1998) were used in this study. The following
bacteriophages were used: l (papa), W24B (Dstx2 :: cat) (Allison et al.,
2003), 933WDtox (Dstx2 :: catGFP), 22Dtox (Dstx2 :: catGFP), 27Dtox
(Dstx2 :: catGFP) and 32Dtox (Dstx2 :: catGFP); derivatives of phages
933W, 22, 27 and 32 were described previously (Gamage et al., 2004),
they are devoid of the toxin genes (due to a safety concern) but have
all other features of the original phages with cat and/or GFP markers;
these are referred to as W24B, 933W, P22, P27 and P32, respectively, in
the text.
Media and growth conditions. Bacteria were routinely cultured in
the Luria-Bertani (LB) medium, supplemented with 10 mM CaCl2
and 10 mM MgSO4. Where appropriate, the following antibiotics
were added: up to 20 mg chloramphenicol ml21 and/or up to 50 mg
kanamycin ml21. The same broth, supplemented with 1.5 % (w/v)
bacteriological agar, was used as a bottom agar for pouring plates.
Top agar consisted of 1 % (w/v) tryptone, 0.5 % (w/v) NaCl and 0.7 %
(w/v) bacteriological agar. Cultures were grown and phage infections
were carried out at 37 uC under aerobic conditions.
1964
Efficiency of lysogenization. To estimate the efficiency of
lysogenization, we used the procedure described in detail previously
(Nowicki et al., 2013), with slight modifications. Briefly, host bacteria
were cultured to A600 1.0 in LB medium supplemented with MgSO4
and CaCl2 (to final concentrations of 10 mM each) at 37 uC with
shaking. Cultures were washed with twice with TMC buffer (10 mM
Tris/HCl pH 7.2, 10 mM MgSO4 and 10 mM CaCl2) and cells were
suspended in the same buffer. Different m.o.i. values (m.o.i of 1, 5 or
10) were used for cell infection. Mixtures of bacteria and phages were
incubated in TMC buffer for 30 min at 30 uC, then one-half of each
mixture was spread on a LB agar plate and the second half on LB agar
plates containing 20 mg chloramphenicol ml21(except for lpapa).
Efficiency of lysogenization was calculated as a fraction of lysogens
(all PCR-tested chloramphenicol-resistant colonies were lysogens)
among all bacterial cells (determined on the basis of number of
colonies appearing on LB agar plates with no antibiotic). The
following primers were used in these tests: catR, 59-GCATTCTGCCGACATGGAAG; catF 59-CACTGGATATACCACCGTTG; gfpR, 59GGTCTGCTAATTGAACGCTTCC and gfpF, 59-CATGGCCAACACTTGTCACTAC (catR and catF are complementary to fragments of
the cat gene and gfpR and gfpF are complementary to fragments of
the GFP gene). The length of the specific reaction product was 599 bp
for cat and 375 bp for GFP. The presence of each prophage was
confirmed by the prophage induction assay. Lysogens of bacteriophage l
were tested by PCR (among 10–30 % of survivors) with the following
primers (complementary to fragments of the cro gene): croF, 59
ATGCGGAAGAGGTAAAGCCC; croR, 59-TGGAATGTGTAAGAGCGGGG (product length 74 bp) and confirmed by the prophage
induction assay.
One-step growth experiments. Lytic development of lambdoid
phages was studied in one step growth experiments in phage-infected
bacteria, as described previously (Nowicki et al., 2013). Briefly,
bacteria were grown in LB medium supplemented with MgSO4 and
CaCl2 (to a final concentration of 10 mM each) at 37 uC to A600 0.2.
The phage was added at an m.o.i. of 0.1. Unadsorbed phages were
removed by double washing in TMC buffer. The number of phageinfected bacterial cells (infection centres) was determined at times
between 0 and 10 min after infection. Culture samples were withdrawn,
0.1 ml of each serial dilution and mixed with 0.3 ml of an overnight
culture of E. coli strain MG1655, 3 ml of top agar (prewarmed to 45 uC)
was added, mixed and poured onto an LB plate. Plates were incubated
overnight at 37 uC and the number of plaques was determined [infected
cells were ‘infection centres’ (ICs) as they were sources of viruses, which
after one lytic developmental cycle could be released from host cells and
after infection of neighbouring cells subsequent lytic cycles could form
plaques on a bacterial lawn]. Samples withdrawn at later times (10 min
and later) were treated with an equal volume of chloroform and titrated
to determine the number of p.f.u. Results were calculated as a ratio of
phage titre to the titre of ICs, thus the numbers in the figures represent
the burst size (number of progeny phages per one infected cell).
Phage lytic development after prophage induction. Prophage
induction experiments were performed as described previously
(Nowicki et al., 2013). Phage lytic development was provoked in
lysogenic bacteria, growing at 37 uC in LB medium at A600 0.2, by
addition of 1 mg mitomycin C ml21. The relative phage titre,
expressed as p.f.u. ml21, was calculated by subtracting the mean value
in the control experiment (without an inducer, which allows
estimation of effects of spontaneous prophage induction) from the
mean value determined in the main experiment.
Estimation of phage DNA synthesis efficiency. Samples of
bacterial culture were withdrawn at indicated times after prophage
induction. Bacteriophage DNA was isolated and separated as described
previously (Nejman et al., 2009; Nowicki et al., 2013) and quantified by
staining with Quant-iT PicoGreen dsDNA (Invitrogen), according to
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Journal of General Virology 96
RNA polyadenylation in development of lambdoid phages
Table 2. Generation times of lysogenic and non-lysogenic WT and DpcnB :: kan strains before and at different times after treatment
with 1 mg mitomycin C ml”1
Bacteria lysogenic for different phages gave very similar results (with no statistically significant differences), therefore, the presented values concern
all lysogens. Results are presented as mean±SD from three independent experiments. Statistically significant (P,0.05 in Student’s t-test) differences
were found only between values indicated by asterisks and other values (but not between both values marked by asterisks).
Strain
Generation time (min)
Before induction
WT
25±1
25±1
28±1
28±1
DpcnB :: kan
WT (lysogen)
DpcnB :: kan (lysogen)
Relative quantification results (E-Method)
2.0
20 min
***
1.8
40 min
1.6
60 min
***
1.4
80 min
1.2
**
1.0
0.8
0.6
0.4
0.2
0.0
MG1655
(WT)
MG1655
(λ)
MG1655
(φ24B)
MG1655
(933W)
Fig. 5. Expression patterns of pcnB gene of WT E. coli strain
MG1655 and its lysogens of bacteriophage l, w24B and 933W,
assessed by RT-qPCR analysis, after treatment of bacterial cells
with 1 mg mitomycin C ml”1. Levels of transcripts corresponding to
pcnB gene were determined at the following times: 20 (white
columns), 40 (black columns), 60 (dark grey columns) and 80 min
(grey columns). The sample at the time point ‘zero’ was used as a
calibrator (all presented quantities are expressed as n-fold
difference relative to the calibrator minus an untreated control
sample; in fact, values for untreated samples were close to zero,
and all calibrator values were similar and low). Statistical analysis
(Student’s t-test) was performed for results from each time point
and indicated significant differences P¡0.01 (**) and P¡0.001
(***) between pcnB gene expression in WT E. coli strain MG1655
and its lysogens. Differences in levels of pcnB gene expression of
tested lysogens (WT versus lysogens) at 40 and 60 min did not
reach significance in the t-test (P.0.05). The results presented
are mean values from three independent experiments with error
bars indicating SD.
manufacturer’s instructions. Concentration of phage DNA (ng ml21)
was calculated relative to l DNA standards of known concentrations and
stained under the same conditions as the tested DNA.
http://vir.sgmjournals.org
Between 0 and 40 min after
induction
Between 40 and 80 min after
induction
24±2
24±2
35±1*
35±2*
24±1
25±2
29±2
29±2
Preparation of RNA and cDNA from bacteria. For the preparation of
RNA, the induction of prophages W24B, 933W and l from E. coli
MG1655 or MG1655DpcnB :: kan was performed with 1 mg mitomycin
C ml21 (Sigma Aldrich) as described above. To inhibit the growth of
bacteria, all samples were treated with 10 mM NaN3 (Sigma Aldrich).
Total RNA was isolated from 16109 bacterial cells using the High
Pure RNA Isolation kit (Roche). The TURBO DNA-free kit (Life
Technologies) was used to remove contaminating DNA from purified
RNA to a level that is mathematically insignificant for RT-qPCR.
RNA preparations were digested with TURBO DNase for 60 min at
37 uC. The total RNA isolated was quantified by staining with Qubit
RNA BR Assay kit (Invitrogen) and Qubit 2.0 Fluorometer (Life
Technologies), which provides an accurate and selective method for
quantification of high-abundance RNA samples. The band patterns of
total RNA were also visualized by electrophoresis. Absence of DNA
from RNA samples was verified by PCR amplification and by RTqPCR amplification of all genes tested. RNA samples were stored at
280 uC. Preparation of cDNA from the total RNA (1.25 mg) was
performed with Transcriptor Reverse Transcriptase and random
hexamer primers (Roche), according to the manufacturer’s instructions. All cDNA reaction mixtures were diluted tenfold for use in
RT-qPCR.
Real-time PCR. Gene expression was determined by RT-qPCR
essentially as described previously (Bloch et al., 2014). All experiments
were performed with the LightCycler 480 Real-Time PCR System
(Roche), using cDNA samples from lysogenic bacteria. Steady-state
levels of transcripts of W24B, 933W and l genes were compared in parallel
to the 16S rRNA, according to a procedure described by Strauch et al.,
(2008), and it was found to be stable. Primers, presented in Table 3, were
developed by Primer3web (version 4) and produced by Sigma Aldrich or
GENOMED. Real-time PCR amplifications were carried out for 55
cycles in a 20 ml reaction using LightCycler 480 SYBR Green I Master
(Roche) as a fluorescent detection dye. All reactions were performed in
Roche 96-well plates. For each reaction, 10 ml of 26 SYBR Green I
Master Mix, 6.25 ng cDNA ml21 and 200 nM of each gene-specific
primer (Table 3) were mixed and supplemented with water to a final
volume of 20 ml. Relative quantification assays were performed with
cDNA relative to 16S rRNA and phage gene multiplex assay. Parameters
for RT-qPCR were as follows: initial incubation at 95 uC for 5 min,
followed by 55 cycles of 95 uC for 10 s, 60 uC for 15 s and 72 uC for 15 s.
Specificity of amplified products was examined by melting curve analysis
immediately after the final PCR cycle and confirmed by gel
electrophoresis. Each experiment was repeated three times.
Real-time PCR data analysis. The relative changes in gene
expression, revealed by quantitative real-time PCR experiments,
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1965
D. Nowicki and others
Table 3. cont.
Table 3. Primers used in real-time PCR
Primer name
pF_W24B_xis
pR_W24B_xis
pF_W24B_cIII
pR_W24B_cIII
pF_W24B_N
pR_W24B_N
pF_W24B_cI
pR_W24B_cI
pF_W24B_cro
pR_W24B_cro
pF_W24B_cII
pR_W24B_cII
pF_W24B_oop
pR_W24B_oop
pF_W24B_O
pR_W24B_O
pF_W24B_Q
pR_W24B_Q
pF_W24B_R
pR_W24B_R
pF_933W_xis
pR_933W_xis
pF_933W_cIII
pR_933W_cIII
pF_933W_N
pR_933W_N
pF_933W_cI
pR_933W_cI
pF_933W_cro
pR_933W_cro
pF_933W_cII
pR_933W_cII
pF_933W_oop
pR_933W_oop
pF_933W_O
pR_933W_O
pF_933W_Q
pR_933W_Q
pF_933W_R
pR_933W_R
pF_l_xis
pR_ l_xis
pF_l_cIII
pR_ l_cIII
pF_l_N
pR_ l_N
pF_l_cI
pR_ l_cI
pF_l_cro
pR_ l_cro
pF_l_cII
pR_ l_cII
pF_l_oop
pR_ l_oop
pF_l_O
pR_ l_O
1966
Sequence (5§ to 3§)
TATCGCGCCGGATGAGTAAG
CGCACAGCTTTGTATAATTTGCG
ATTCTTTGGGACTCCTGGCTG
GTAAATTACGTGACGGATGGAAAC
AGGCGTTTCGTGAGTACCTT
TTACACCGCCCTACTCTAAGC
TGCTGTCTCCTTTCACACGA
GCGATGGGTGGCTCAAAATT
CGAAGGCTTGTGGAGTTAGC
GTCTTAGGGAGGAAGCCGTT
TGATCGCGCAGAAACTGATTTAC
GACAGCCAATCATCTTTGCCA
TGCAAGACCGAGCGTTCT
CGCATATAATTACCTCGTTGGATG
AAGCGAGTTTGCCACGAT
GAACCCGAACTGCTTACCG
GGGAGTGAGGCTTGAGATGG
TACAGAGGTTCTCCCTCCCG
GGGTGGATGGTAAGCCTGT
TAACCCGGTCGCATTTTTC
CTGTCCCTGGCGCGTAATAT
GGTAACACTGGAGCTTTTAATGGC
ATTCTTTGGGACTCCTGGCTG
GTAAATTACGTGACGGATGGAAAC
AGGCGTTTCGTGAGTACCTT
TTACACCGCCCTACTCTAAGC
TGCTGTCTCCTTTCACACGA
GCGATGGGTGGCTCAAAATT
CGAAGGCTTGTGGAGTTAGC
GTCTTAGGGAGGAAGCCGTT
TGATCGCGCTGAAACTGATTTAC
GACAGCCAATCATCTTTGCCA
GCCGGGGACTCTAAGCAA
GATTGATCCAGTAATTCCCTCAGA
AATTCTGGCGAATCCTCTGA
GAATTGCATCCGGTTT
GGGAGTGAGGCTTGAGATGG
TACAGAGGTTCTCCCTCCCG
GGGTGGATGGTAAGCCTGT
TAACCCGGTCGCATTTTTC
TACCGCTGATTCGTGGAACA
GGGTTCGGGAATGCAGGATA
ATTCTTTGGGACTCCTGGCTG
GTAAATTACGTGACGGATGGAAAC
CTCGTGATTTCGGTTTGCGA
AAGCAGCAAATCCCCTGTTG
ACCTCAAGCCAGAATGCAGA
CCAAAGGTGATGCGGAGAGA
ATGCGGAAGAGGTAAAGCCC
TGGAATGTGTAAGAGCGGGG
TCGCAATGCTTGGAACTGAGA
CCCTCTTCCACCTGCTGATC
GCAACCGAGCGTTCTGAA
GATAGATCCAGTAATGACCTCAGA
AATTCTGGCGAATCCTCTGA
GAATTGCATCCGGTTT
Primer name
pF_l_Q
pR_ l_Q
pF_l_R
pR_ l_R
pF_E.coli_pcnB
pR_E.coli_pcnB
pF_E.coli_16SrRNA
pR_ E.coli_16SrRNA
Sequence (5§ to 3§)
TTCTGCGGTAAGCACGAAC
TGCATCAGATAGTTGATAGCCTTT
ATCGACCGTTGCAGCAATA
GCTCGAACTGACCATAACCAG
AGCCAGACGGAAACGGC
ACCACTAACGCCACGCC
CCTTACGACCAGGGCTACAC
TTATGAGGTCCGCTTGCTCT
were analysed using the calibrator normalized relative quantification
method with efficiency correction, the so-called E-Method. The EMethod produces more accurate relative quantification data because
it can compensate for differences in target and reference gene
amplification efficiency either within an experiment or between
experiments. This method provides an efficiency corrected calculation
mode by using the determined PCR efficiency of target (Et) as well
as the efficiency of reference (Er). Relative fold change ratio was
calculated using the following formula, described in the application
manual of Roche LightCycler Real-Time PCR Systems: Normalized
relative ratio5Et CT(t) calibrator2CT(t) sample/Er CT(r) calibrator2CT(r) sample
(where ‘t’ is target and ‘r’ is reference). The sample at the time point
‘zero’ was used as a calibrator. The raw run data for W24B, 933W and l
genes were transferred from the LightCycler 480 to the LinRegPCR 12.5
software using the ‘LC480 Conversion: conversion of raw LC480 data’
software
(available
at
http://www.hartfaalcentrum.nl/index.
php?main=files&sub=0). PCR efficiency was determined for each gene
by LinRegPCR program (Ramakers et al., 2003; Ruijter et al., 2009). This
software was successfully used previously to calculate PCR efficiency
(Čikoš et al., 2007; Feng et al., 2008; Regier & Frey, 2010; Aglawe et al.,
2012, Borges et al., 2012).
Statistical analysis. For each experiment, data from independently
replicated experiments (each experiment was repeated at least three
times) were pooled and analysed using STATISTICA (StatSoft). Variation
among replicates was used as the error term. Data comparisons were
made by using a paired Student’s t-test. Differences were considered
significant when P,0.05.
ACKNOWLEDGEMENTS
The authors thank Dr Katarzyna Potrykus for critical reading of the
manuscript. This work was supported by the National Science Center,
Poland (grant no. N N301 192439 to A. W.) and Foundation for
Polish Science (grant no. VENTURES/2012-9/7 to D. N.). Studies by
D. N., S. B. and B .N. -F. were supported by the European Social Fund
as a part of the project ‘Educators for the elite-integrated training
program for PhD students, post-docs and Professors as academic
teachers at University of Gdansk’ within the framework of the Human
Capital Operational Programme, Action 4.1.1, improving the quality
of educational offer of tertiary education institutions. S. B. acknowledges support from the European Social Fund, the State Budget and
the Pomorskie Voivodeship Budget according to the Operational
Programme Human Capital, Priority VIII, Action 8.2, Underaction
8.2.2: ‘Regional Innovation Strategy’ within the system project of the
Pomorskie Voivodeship ‘InnoDoktorant – Scholarships for PhD
students, VIth edition’. The authors declare no conflict of interest.
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Journal of General Virology 96
RNA polyadenylation in development of lambdoid phages
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