downloaded from · 9 valerie j. carabetta 1, bijoy k. mohanty 2, sidney r. kushner 2, and thomas j....

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The response regulator SprE (RssB) modulates polyadenylation and mRNA stability in 6

Escherichia coli 7

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Valerie J. Carabetta1, Bijoy K. Mohanty2, Sidney R. Kushner2, and Thomas J. Silhavy1* 9

1Department of Molecular Biology, Princeton University, Princeton, New Jersey 08544 10

2Department of Genetics, University of Georgia, Athens, Georgia 30605 11

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Running title: SprE modulates RNA polyadenylation 15

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*Corresponding author. Mailing address: Department of Molecular Biology, Princeton 19

University, Princeton, NJ 08544. Phone: (609) 258-5899. Fax: (609) 258-2957. E-mail: 20

[email protected] 21

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Copyright © 2009, American Society for Microbiology and/or the Listed Authors/Institutions. All Rights Reserved.J. Bacteriol. doi:10.1128/JB.00870-09 JB Accepts, published online ahead of print on 18 September 2009

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Abstract 1

In Escherichia coli, the adaptor protein SprE (RssB) controls the stability of the alternate 2

sigma factor, RpoS (σ38, σS). When nutrients are abundant, SprE binds RpoS and delivers it to 3

ClpXP for degradation; but when carbon sources are depleted, this process is inhibited. It also 4

has been noted that overproduction of SprE is toxic. Here we show that null mutations in pcnB, 5

encoding poly(A) polymerase I (PAP I), and in hfq, encoding the RNA chaperone Hfq, suppress 6

this toxicity. Since PAP I, in conjunction with Hfq, is responsible for targeting RNAs, including 7

mRNAs, for degradation by adding poly(A) tails onto their 3’ ends, these data indicate that SprE 8

helps modulate the polyadenylation pathway in E. coli. Indeed, in exponentially growing cells, 9

sprE deletion mutants exhibit significantly reduced levels of polyadenylation and increased 10

stability of specific mRNAs, similar to what is observed in a PAP I-deficient strain. In stationary 11

phase, we show that SprE changes the intracellular localization of PAP I. Taken together, we 12

propose that SprE plays a multi-functional role in controlling the transcriptome, regulating what 13

is made via its effects on RpoS, and modulating what is degraded via its effects on 14

polyadenylation and turnover of specific mRNAs. 15


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Introduction 1

Natural microbial environments often require bacteria to rapidly adapt to a variety of 2

stresses such as nutrient starvation. Escherichia coli responds to starvation by entering a 3

protective, metabolically less active state, known as stationary phase (20). Cells in stationary 4

phase can survive for weeks without essential nutrients and rapidly resume growth once 5

environmental conditions improve and nutrients become available again. The transition from 6

exponential growth to stationary phase is accompanied by multiple changes that affect all 7

cellular processes. For example, there is an increase in proteolysis, accompanied by a decrease 8

in overall transcription and translation (15, 20). Despite the global decrease in these processes, 9

the expression of a subset of stress-responsive genes is induced as the cells enter stationary 10

phase. These genes are controlled by the stationary phase sigma factor RpoS (σ38, σS) (15, 50). 11

The RpoS regulon consists of genes that specify proteins that combat multiple stresses and 12

catalyze the synthesis of storage compounds, such as glycogen (15, 25). 13

Because of the drastic changes that occur upon entry into stationary phase, RpoS is one of 14

the most highly regulated proteins in E. coli. It is controlled at all levels: transcription, 15

translation, protein stability, and activity (15). During exponential growth, when nutrients are 16

abundant, RpoS levels are kept low by the action of the adaptor protein SprE (RssB) and the 17

ClpXP protease. After SprE binds to RpoS, the complex interacts with the ATP-dependent 18

chaperone ClpX. RpoS is subsequently unfolded by ClpX and degraded by the ClpP protease, 19

releasing SprE, which is recycled (19, 34, 39, 52). Upon entry into stationary phase, this process 20

is inhibited, leading to the accumulation of RpoS and the transcription of its target genes (22, 21

51). 22


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The N-terminal domain of SprE shares strong sequence homology to the receiver domain 1

of the response regulator family of proteins (5). In a typical two domain response regulator, 2

regulation of activity occurs through phosphorylation at a conserved aspartate (D58 in SprE) in 3

the N-terminal domain. However, a mutant form of SprE, in which the conserved aspartate 4

residue was changed to alanine, responds correctly to starvation for various essential nutrients, 5

including carbon. Thus, phosphorylation does not regulate SprE activity in response to 6

starvation (37). The C-terminal output domain of SprE shares no sequence homology to any 7

known family of proteins, making it a novel response regulator. 8

It has recently been discovered that SprE is targeted by a set of anti-adaptor proteins, 9

exemplified by IraP, that bind to SprE and inhibit its activity under certain stress conditions. 10

Each anti-adaptor protein appears to be specific for a particular stress, but not all stresses 11

necessarily elicit the production of an anti-adaptor protein. For example, no SprE-specific anti-12

adaptor protein has been identified during carbon starvation. If SprE indeed senses this 13

particular type of starvation, the mechanism is not understood (7, 8, 12). 14

SprE levels are growth phase regulated. There are low levels of SprE during exponential 15

growth, while entry into stationary phase increases protein levels 2-3 fold (41). SprE acts to 16

modulate RpoS primarily during exponential phase, therefore it is not obvious why SprE levels 17

increase at a time when it no longer functions in this capacity. One possible explanation is that 18

increasing SprE levels in stationary phase is necessary to prime the cells for the rapid destruction 19

of RpoS when cells encounter nutrients and return to exponential growth (41). Another 20

hypothesis is that SprE has a second function in stationary phase cells that requires higher levels 21

of the protein. Interestingly, it has been noted that overproduction of SprE causes a significant 22

growth defect in exponentially growing cells (34, 43). The expected phenotype of the 23


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overproduction of SprE is lower RpoS levels, even in stationary phase. However, rpoS null 1

mutants do not exhibit the growth defect seen with SprE overproduction, thus SprE-mediated 2

degradation of RpoS cannot account for this observed toxicity. 3

Here we have exploited the growth defect associated with SprE overproduction to 4

uncover an additional role for this protein in polyadenylation and the control of mRNA stability. 5

Specifically, in exponentially growing cells the absence of SprE results in a significant reduction 6

in poly(A) levels and a consequent increase in the half-lives of specific mRNAs, which have 7

previously been shown to be dependent on polyadenylation for their decay. In stationary phase 8

cells lacking SprE, the intracellular location of poly(A) polymerase I is altered, suggesting a 9

multi-functional role for SprE in the control of the transcriptome. 10

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Materials and Methods 12

Bacterial strains, media and growth conditions 13

All strains are listed in Table 1. Standard microbial techniques were used for strain 14

constructions (44). Luria broth and M63 liquid medium and agar were prepared as described 15

previously (44), and were supplemented with appropriate antibiotics as needed. Antibiotic 16

concentrations used were as follows: 125 µg/mL ampicillin (Ap), 25 µg/mL tetracycline (Tc), 20 17

µg/mL chloramphenicol (Cm), 50 µg/mL kanamycin (Km), and 20 µg/mL spectinomycin. 18

Bacteria were grown at 37°C with aeration and growth was monitored by measuring the OD600. 19

Plasmids were induced with 0.2% arabinose or 200 ng/mL anhydrotetracycline (ATC), where 20

applicable. 21

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Plasmid constructions 23


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For construction of pZS*11sprE+, colony PCR was performed on MC4100 cells with 1

primers that amplify the sprE open reading frame (ORF). The primers 5sprEEcoRI (5’-2

AGAAACCGAATTCATTAAAGAGGAGAAAGGTACCGCATGACGCAGCCATTGGTCGG3

) and 3sprEXbaI (5’-4

GGCTCTAGACTCAGCTAATTAAGCTCATTCTGCAGACAACATCAAGCGC) were used 5

for cloning. The forward primer contained a non-native ribosome binding site upstream of the 6

sprE start codon. After digestion with EcoRI and XbaI (sites underlined), the resulting PCR 7

product was ligated into the EcoRI and XbaI sites of pZS*11 (23). This plasmid (3-4 copies/cell) 8

placed the sprE gene under the control of the tetracycline promoter (23). The Tet repressor was 9

supplied in trans to repress sprE expression. 10

For the construction of pBADsprE+, colony PCR was performed using 5sprEEcoRI and 11

3sprEpBADHindIII (5’-TCCAAGCTTTGCTCATTCTGCAGACAACATCAAGCGC). After 12

digestion with EcoRI and HindIII, the PCR product was ligated into the EcoRI and HindIII sites 13

on pBAD18. This plasmid (15-30 copies/cell) placed the sprE gene under control of the 14

arabinose inducible promoter, PBAD (13). The ppcnB-GFP plasmid was constructed as follows. 15

The primers 5pcnBStuI (5’-TGGCGGAGGCCTCAGCGTCGAGCAAATCCTTCAG) and 16

3pcnBNheIns (5’-GGATCTGCTAGCTGCTGCTGCTGCGGTACCCTCACGACGTGGT) were 17

used to amplify the 140 base pairs upstream of pcnB, which contained the native pcnB 18

promoters, as well as the entire pcnB ORF. The cloned pcnB fragment replaced the stop codon 19

with three alanine codons to serve as a linker to an in-frame translational fusion with GFP at the 20

C-terminus. The resulting PCR product was digested and ligated into the StuI and NheI 21

restriction sites on pCMW1, kindly provided by Chris Waters (49). This plasmid expressed the 22

PAP I-GFP fusion protein under the control of the native pcnB promoters. The plasmids 23


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pBMK28 [pcnB-(His)6 Cmr] and pDK24 (rnb Apr) have been described previously (32, 33). All 1

oligonucleotides were synthesized by Integrated DNA Technologies. Each construct was 2

verified by DNA sequence analysis by Genewiz Inc. (South Plainfield, NJ). 3

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PCR mutagenesis and the screen for novel sprE mutations 5

Random PCR mutagenesis was performed using the GeneMorph Random Mutagenesis 6

Kit (Stratagene) as per manufacturer’s instructions. Briefly, 500 ng of the plasmid pZS*11sprE+ 7

was subjected to 30 rounds of mutagenic PCR in order to achieve a mutation rate of 0-3 8

mutations/kb. The primers 5sprEEcoRI and 3sprEXbaI were used for the PCR reaction, so that 9

we could clone the mutagenized sprE products as described above. The pool of mutagenized 10

plasmids was transformed into VC30, which contained the rpoS750’-‘lacZ reporter fusion. This 11

fusion is subject to the same regulation as full-length RpoS (45). Colonies were screened for 12

color on LB agar containing 125 µg/mL Ap + 80 µg/mL X-gal after incubation overnight at 13

37°C. Colonies that appeared to have a different color than the parent (VC30), indicating 14

changes in RpoS regulation were analyzed further. Plasmids were mini-prepped following 15

manufacturer’s instructions (Qiagen) and candidates were sequenced by Genewiz Inc. 16

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Suppressors of SprE-mediated toxicity 18

Strains containing pBADsprE+ were streaked on LB agar containing 125 µg/mL Ap + 19

0.2% arabinose. Faster growing colonies were selected from the slow-growing parental lawn 20

and were purified. Plasmid DNA was mini-prepped as described above and was retransformed 21

into the parent strain to check for plasmid linkage (44). All plasmid linked mutations were 22


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discarded. Chromosomal mutations were mapped by using a library of random transposon 1

mutations according to standard procedures (44). 2

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Growth rates 4

Growth was monitored every 20 minutes for 3-4 hours, by measuring the OD600. 5

Doubling times were calculated using data points obtained during mid-exponential growth. At 6

least three independent experiments were performed for each strain analyzed. 7

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Site-directed mutagenesis 9

Plasmids were mutagenized using the Gene Tailor Site-Directed Mutagenesis system 10

(Invitrogen) according to the manufacturers’ instructions. Plasmid DNA was methylated at 11

cytosine residues prior to PCR. The D58A mutation was engineered in trans into the plasmid 12

pZS*11sprE60 (see results). The primers sprED58Afor (5’- 13

CTCCAGACCTGATGATATGTGCTATCGCGATG) and sprED58Arev (5’- 14

ACATATCATCAGGTCTGGAGTGAAACCTCC) were used to introduce a GAT � GCT 15

transversion (underlined). Mutagenized plasmids were transformed into cells that contained the 16

McrBC endonuclease to degrade the methylated, non-mutated parental strands. Plasmids were 17

mini-prepped as described above and were sequenced by Genewiz Inc. 18

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SDS PAGE and western blot analysis 20

Cells were grown to an OD600 of 0.1, followed by induction of plasmid expression for 1 21

hour, where applicable. At mid-exponential phase (OD600 0.4-0.5) the cells were pelleted and 22

were immediately resuspended with sodium dodecyl sulfate (SDS) sample buffer. Samples were 23


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boiled for 10 minutes and then equal volumes were loaded onto a 12% polyacrylamide gel (44). 1

Following electrophoresis, proteins were transferred to a Protran nitrocellulose membrane 2

(Whatman) and were probed with either a 1:6000 dilution of RpoS antibody or a 1:4000 dilution 3

of SprE antibody (laboratory stocks). Donkey anti-rabbit immunoglobulin G-horseradish 4

peroxidase conjugate (GE Healthcare) was used as a secondary antibody at a dilution of 1:6000. 5

Bands were visualized using the ECL antibody detection kit (GE Healthcare) and Hyblot CL film 6

(Denville Scientific Inc.). Quantification of bands was carried out using the ImageJ program (1). 7

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Fluorescence microscopy 9

One mL of overnight cultures or mid-exponential cultures were pelleted and washed 10

twice with one mL of M63liquid medium. Overnight cultures were resuspended in one mL of 11

M63, while mid-exponential cultures were resuspended in 200 µL M63 in order to concentrate 12

the cells. From the resuspended cells, 5 µL was spotted onto a 1% agarose pad (in M63). 13

Images were obtained employing a QImaging Rolera-XR camera on a Nikon 90i microscope 14

equipped with a Nikon Plan Apo 1.4/100× Oil Ph3 phase objective, using the NIS Elements 15

software package. At least two independent experiments were performed per strain, and at least 16

150 cells were counted for determination of localization percentages. 17

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Isolation and analysis of total RNA 19

The total RNA from exponentially growing cultures was isolated using the Catrimide 20

method as described previously (26). All cultures were grown to Klett 50 (no. 42 green filter) in 21

LB at 37˚C with shaking. Total RNA from the stationary phase cultures was isolated after 16-18 22

hrs of growth in LB at 37˚C with shaking using an SDS lysis method as described previously 23


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(26). The strains containing pZS*11sprE60 were grown in presence of 200 ng/mL of ATC. 1

The RNA-DNA dot blots used to determine total intracellular poly(A) levels were carried out as 2

previously described (26). Total RNA for half-life determinations was isolated at various time 3

points after addition of rifampicin to stop the transcription initiation as described before (26). 4

The half-lives of full-length transcripts were determined by Northern analysis by separating the 5

transcripts either by 7% acrylamide/8M urea or 1.5% agarose-glyoxal gels (26). Half-lives were 6

determined using linear regression analysis. 7

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Results 9

Growth rate is decreased in the presence of increased SprE levels 10

It has previously been reported that overexpression of SprE causes significant growth 11

defects (34, 43). To determine the levels of SprE required to cause toxicity, we initially 12

examined a strain carrying an rssA2::cam chromosomal allele, which contains a transposon 13

inserted upstream of the sprE gene such that expression is driven by the constitutive 14

chloramphenicol promoter (41). This allele resulted in a 4-fold increase in SprE levels relative to 15

wild-type (Fig. 1A, lanes 1 and 3, Table 2) with no observable effect on growth rate (Table 2). 16

Next the sprE open reading frame was cloned under the control of the anhydrotetracycline-17

inducible PLtetO-1 promoter into a low copy number (3-4 copies/cell) vector, pZS*11. Induction 18

from this vector led to a 5-fold increase above wild-type SprE levels (Fig. 1A, lane 4, Table 2). 19

Strains containing pZS*11sprE+ had no apparent growth defect on solid medium containing 20

anhydrotetracycline. However, this plasmid did confer a slight growth defect in liquid medium 21

under inducing conditions (40.6 +/- 4.5 minute doubling time, Table 2), when compared to the 22

parent strain carrying the vector control (33.8 +/- 4.3 minute doubling time, Table 2). To further 23


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increase SprE levels, the pBADsprE+ plasmid was constructed, which led to >20-fold 1

overproduction of SprE when induced (Fig. 1A, lane 5). Overproduction of SprE at this level 2

caused a pronounced toxicity, as evidenced by poor growth and the appearance of multiple 3

suppressors growing out of the parental lawn on solid medium. The generation time of the strain 4

harboring pBADsprE+ was >60 minutes (data not shown). 5

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Isolation of sprE mutations that enhance toxicity 7

The overproduction of a given protein can be toxic to cells due to a particular protein’s 8

activity or the formation of toxic aggregates. To help distinguish these two possibilities, we 9

mutagenized pZS*11sprE+, as described in the Materials and Methods, since this plasmid did not 10

confer a significant growth defect (Table 2). We reasoned that if toxicity resulted from an 11

activity of SprE, we could isolate mutants that showed enhanced activity. 12

We identified two mutants that conferred toxicity without changing the intracellular level 13

of the SprE protein (Fig. 1B, compare lanes 1 and 2). Sequencing of these mutants revealed two 14

different alleles, both of which carried multiple mutations: sprEL29F, A97T and sprEL29F, 15

V120I, L124F. Reconstruction experiments using site-directed mutagenesis demonstrated that 16

each of the multiple mutations was required for the mutant phenotype (data not shown). Since 17

both alleles were phenotypically indistinguishable, sprEL29F, V120I, L124F, referred to as 18

sprE60 for simplicity, was used for all studies described below. 19

The SprE60 protein caused a growth defect similar to that seen with pBADsprE+ (61.8 20

+/- 8.6 minutes, Table 2). However, in either case, it should be noted that this growth rate may 21

actually be an under representation of the true growth rate of these mutants. Occasionally mutant 22

cultures had doubling times similar to the wild-type control, indicating that a faster-growing 23


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suppressor took over the culture at an early time point, particularly since a doubling time in 1

excess of 100 minutes was also observed once (data not shown). 2

Expression of sprE from the pBAD plasmid resulted in the production of ~20 times 3

greater SprE levels than wild-type and a generation time of >60 min. The expression of the 4

sprE60 allele from the PLtetO-1 promoter on pZS*11 was much lower, yielding protein levels 5-6 5

times greater than wild type. Nevertheless, expression of the two plasmid constructs resulted in 6

comparably poor growth with similar generation times. These results suggested that the point 7

mutations associated with the sprE60 allele may have increased the inherent toxicity associated 8

with high level overproduction of wild-type SprE. Although not conclusive, these results 9

supported the possibility that it is an activity of SprE that causes toxicity. 10

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Toxicity is not related to any known function of SprE 12

SprE plays an important role in controlling the intracellular level of the RpoS sigma 13

factor. However, strains lacking rpoS, sprE or clpXP, all players in the regulation of RpoS 14

protein levels, are all viable and they exhibit no apparent growth defects. In order to determine if 15

any member of the RpoS pathway was involved with the toxicity associated with overproduction 16

of SprE, chromosomal null alleles of rpoS, sprE and clpXP were introduced into the strains 17

carrying either pZS*11sprE60 or pBADsprE+. Upon induction of expression of either sprE 18

allele, none of these null alleles relieved the toxicity associated with either plasmid (data not 19

shown). These results indicate that toxicity does not depend upon RpoS levels, per se, as the 20

rpoS and clpXP mutations affect the levels of RpoS in a completely opposite fashion. These 21

results also demonstrate that the toxicity we observed was not related to SprE’s known function 22

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As noted above, the N-terminal domain (NTD) of SprE shares homology with the 1

response regulator family of proteins and contains a conserved aspartate that serves as the 2

phospho-acceptor site (6). Surprisingly, mutation of this site, D58A, resulted in an altered SprE 3

protein that is still functional (37), contrary to most other response regulators (16). In order to 4

determine if SprE phosphorylation was associated with toxicity, we cloned the sprED58A allele 5

into pBAD18. Overproduction of the non-phosphorylatable SprE protein still caused toxicity 6

(data not shown); suggesting that overproduction of the wild-type SprE protein does not lead to a 7

titration of phosphate from an essential kinase(s). The D58A mutation was also engineered by 8

site-directed mutagenesis into pZS*11sprE60 to create pZS*11sprE60D58A. Under these 9

circumstances, there was a partial restoration of the doubling time from 61.8 +/- 8.6 minutes to 10

48.0 +/- 5.0 min (Table 2). We conclude that phosphorylation is not required for the toxicity 11

phenotype per se, but it may enhance it. 12

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Suppressors of SprE toxicity 14

In order to gain insight into the cause of the toxicity, we isolated and characterized 15

suppressors of the slow growth phenotype. Specifically, strains carrying pBADsprE+ were 16

plated on LB agar containing 0.2% arabinose, and faster growing colonies that grew above the 17

background lawn were purified. Plasmid DNA was isolated from the faster growing variants to 18

check for plasmid-linked mutations, which were discarded. Standard genetic techniques were 19

used to map one of the chromosomal suppressors as described in Materials and Methods. DNA 20

sequence analysis showed that this suppressor was a null allele of pcnB, which encodes poly(A) 21

polymerase I (PAP I), the enzyme that adds poly(A) tails onto the 3’ ends of full-length mRNAs 22

as well as decay products, facilitating their degradation (21, 27, 36). 23


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PAP I is also involved in the copy number control of ColE1 based plasmids through the 1

polyadenylation of the anti-sense RNA (RNA I), which is involved in replication control (14). In 2

the absence of PAP I, plasmid copy number is significantly decreased, possibly explaining why 3

this mutation suppressed the toxicity of pBADsprE+, a plasmid containing a ColE1 origin of 4

DNA replication (13). If a decrease in plasmid copy number were the primary reason for the 5

suppression of the slow-growth phenotype, then inactivation of PAP I would not be expected to 6

alter the toxicity caused by pZS*11sprE60, since this plasmid contains the pSC101 origin, which 7

is not affected by polyadenylation (24). However, a ∆pcnB allele also suppressed the toxicity 8

associated with pZS*11sprE60 (Table 2). In fact, the presence of the ∆pcnB allele led to a 9

doubling time that was only slightly longer than the wild-type control (Table 2). Moreover, the 10

absence of PAP I did not significantly affect the levels of the SprE60 protein (Fig. 1B, compare 11

lanes 2 and 3, and Table 2). These results suggest that inactivation of PAP I does not relieve the 12

toxicity by lowering gene dosage or the levels of the sprE mRNA. In fact, in a ∆pcnB strain 13

there was a slight increase in SprE levels (Table 2). The absence of PAP I also suppressed the 14

slight growth defect associated with the presence of pZS*11sprE+, restoring the growth rate to 15

32.6 +/- 0.9 minutes, which was equivalent to that of the strain carrying the empty vector control 16

pZS*11 (Table 2). These results provide evidence that the toxicity associated with the SprE60 17

and overproducing wild-type SprE are indeed the same. 18

PAP I requires the RNA-chaperone Hfq to polyadenylate mRNA’s containing 3’ Rho-19

independent transcription terminators (33). Since PAP I forms a complex with Hfq (33), we 20

hypothesized that an ∆hfq null allele might also suppress the toxicity phenotype. A strain 21

carrying pZS*11sprE60 and a ∆hfq allele had a doubling time of 38.6 +/- 2.8 minutes, which was 22

very similar to that of the parent containing pZS*11sprE+ and a result comparable to that 23


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observed with a ∆pcnB allele (Table 2). Furthermore, inactivation of Hfq did not affect SprE60 1

protein levels (Fig. 1B, compare lanes 2 and 5). Interestingly, inactivation of Hfq in an 2

otherwise wild-type genetic background led to a two-fold increase in SprE levels (Table 2). We 3

conclude that the toxicity phenotype caused by high levels of SprE or by SprE60 requires the 4

presence of both PAP I and Hfq. 5

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Interactions of SprE and various ribonucleases 7

Since polyadenylation has been shown to affect the decay of a significant number of E. 8

coli mRNAs (21, 28, 33, 36), we tested whether any of the ribonucleases involved in mRNA 9

decay also interacted with SprE. Accordingly, the plasmid pZS*11sprE60 was transformed into 10

strains carrying one of the following loss-of-function mutations: rne-1 (RNase E), rnc∆38 11

(RNase III), rnb-296 (RNase II), pnp∆683 (polynucleotide phosphorylase, PNPase) or rnr::kan 12

(RNase R). Transformants were easily obtained in the rne-1, rnc∆38, rnb-296 and rnr::kan 13

strains and the phenotypic properties of these transformants were comparable to those observed 14

for a wild-type control carrying pZS*11sprE60 (data not shown). 15

In contrast, very slow-growing transformants were obtained with the pnp∆683 strain. 16

These transformants were extremely unstable throwing off faster growing variants, many of 17

which had lost the plasmid. It is thus likely that a pnp∆683 pZS*11sprE60 strain is a synthetic 18

lethal combination. Indeed, we were unable to construct this double mutant by generalized 19

transduction. Since it is known that poly(A) levels increase significantly in a pnp null mutant 20

(33), we hypothesized that SprE60 might also cause an increase in poly(A) levels such that a 21

synergistic interaction occurred in the pnp∆683 pZS*11sprE60 strain, leading to cell death. The 22

idea that SprE60 (or high levels of wild-type SprE) increases poly(A) levels also explains the 23


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suppression of the toxicity phenotype by pcnB and hfq null alleles, since these alleles lead to 1

reduced polyadenylation (33). 2

It has also been observed that PAP I overproduction leads to increased levels of poly(A) 3

and cell death over time (27). This cell death is suppressed by co-overproduction of RNase II, an 4

exonuclease known to degrade poly(A) tails (32). If SprE60 toxicity were also the result of 5

increased poly(A) levels, increasing levels of RNase II might suppress the toxicity. Accordingly, 6

we transformed a compatible plasmid containing rnb under control of its native promoter into 7

strains carrying pZS*11sprE+ and pZS*11sprE60. Overproduction of RNase II in combination 8

with pZS*11sprE+ resulted in a doubling time of 47.9 +/- 4.6 minutes, and in combination with 9

pZS*11sprE60, the doubling time was nearly identical at 47.7 +/- 3.8 minutes (Table 2). We 10

conclude that the toxicity caused by SprE60 was at least partially suppressed by increasing the 11

levels of RNase II. 12

13

SprE affects intracellular levels of polyadenylation 14

Since inactivation of PAP I suppressed the growth defect associated with overproduction 15

of SprE60, we hypothesized that SprE played some role in the regulation of intracellular levels of 16

poly(A). To test this hypothesis directly, we measured poly(A) levels in exponentially growing 17

cells using RNA-DNA dot blots. As seen in Figure 2, the presence of pZS*11sprE60 led to a 2-18

fold increase in poly(A) levels compared to the wild type control. When the same experiment 19

was carried out in either a ∆pcnB/ pZS*11sprE60 or hfq-1/ pZS*11sprE60 strain, poly(A) levels 20

were significantly lower than those observed in the wild type control. These data correlated with 21

the shorter generation times observed in these strains, relative to the sprE+/ pZS*11sprE60 strain 22

(Table 2). 23


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Since overproduction of the SprE60 protein led to a significant increase in total poly(A) 1

levels (Fig. 2), we tested to determine if wild-type SprE was involved in the regulation of 2

intracellular levels of polyadenylation. In agreement with previously reported results in the 3

MG1655 genetic background (27, 33), poly(A) levels were reduced 70% and 60%, respectively, 4

in ∆pcnB and hfq-1 strains (Fig. 2), while deletion of the sprE locus caused a 40-50% reduction 5

in poly(A) levels in exponentially growing cells (Fig. 2). In contrast, poly(A) levels were 6

unchanged in a clpX null strain (Fig. 2), which suggests that the RpoS-degradation function of 7

SprE is separate and distinct from the polyadenylation phenotype. 8

9

Deletion of sprE results in increased mRNA half-lives in exponentially growing cells 10

The data presented above shows that deletion of the SprE protein leads to a significant 11

reduction in the intracellular poly(A) levels, suggesting a role of SprE in the regulation of the 12

polyadenylation pathway. Since it has been shown that the half-lives of specific mRNAs are 13

increased in the absence of polyadenylation (21, 33, 36), we predicted that for those mRNAs 14

whose decay is stimulated by polyadenylation we would see comparable half-lives in ∆pcnB and 15

∆sprE mutants. Accordingly, we used Northern blot analysis to determine the half-lives of six 16

specific mRNAs. As shown in Table 3, the half-lives of 5/6 of the tested mRNAs were increased 17

in both the ∆pcnB and ∆sprE strains. The single exception, rplY, had identical half-lives in all 18

genetic backgrounds. This result was consistent with earlier observations that the half-life of 19

rplY is not affected by the level of polyadenylation (29). Further support for SprE’s direct role in 20

the polyadenylation and degradation of mRNAs was derived from the decay patterns observed 21

for at least one of the six transcripts. As shown in Figure 3, specific breakdown products 22

(arrows) of the yfiA mRNA transcript accumulated in both the ∆pcnB and ∆sprE strains, but were 23


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barely detectable in the wild-type control. In fact, relative to the amount of full-length yfiA 1

mRNA in this analysis, the amount of the decay intermediate indicated by the upper arrow (Fig. 2

3) increased from <0.1 % at 0 min up to ∼10% at 32 min in the wild-type strain. In contrast, it 3

increased from 0.2% to 18% and 0.2% to 36% in the ∆pcnB and ∆sprE strains, respectively. 4

These results support a role for SprE in mRNA stability. 5

6

SprE does not affect PAP I protein levels 7

It has been shown that intracellular levels of polyadenylation can be affected by changing 8

the level of PAP I (27), altering the levels of either RNase II or PNPase (32), or by inactivating 9

Hfq, a component of the polyadenylation complex (33). Accordingly, we used Western blot 10

analysis to measure these protein levels in wild-type and ∆sprE strains. PAP I levels were 11

identical, within experimental error, in both strains (Fig. 4A). Similar results were obtained with 12

PNPase (data not shown). In contrast, there was a small but consistent 30-40% increase in the 13

level of RNase II (Fig. 4B). Clearly the decreased polyadenylation in strains lacking SprE is not 14

due to decreases in the levels of either PAP I or PNPase. Although it cannot be ruled out that the 15

decreased polyadenylation results from the small increase in RNase II observed in the absence of 16

SprE, we think this unlikely. 17

18

Phenotypic consequences of increased polyadenylation 19

During the course of this work we noticed that a strain carrying the rssA2 allele exhibited 20

an extended lag phase when exiting stationary phase. Specifically, after diluting an overnight 21

stationary phase culture of wild-type E. coli 100-fold into fresh medium, there was an 22

approximate 20 min delay before growth resumed. In contrast, rssA2 strains did not resume 23


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growth until 40 min after dilution (Fig. 5). In both the wild-type and rssA2 strains, CFU’s were 1

identical at time 0 indicating that a decreased number of viable cells was not the underlying 2

cause of the delay. Since deleting either rpoS or sprE had no effect on the lag time compared to 3

a wild-type control (data not shown), we hypothesized that the defect in the exit from stationary 4

phase might be related to altered polyadenylation in stationary phase cells resulting from 5

increased levels of the SprE protein. If this were the case, a ∆pcnB allele would be expected to 6

suppress this defect. As shown in Figure 5, this was in fact the case. 7

If changes in polyadenylation resulted in a defect in the exit from stationary phase, then 8

overproduction of PAP I from a plasmid should produce a similar phenotype. When pBMK28, a 9

plasmid carrying a pcnB-his6 allele under the control of an IPTG inducible promoter (33), was 10

introduced into a wild-type strain, even the uninduced transformants showed a 40 min delay in 11

the exit from stationary phase, identical to what was observed with the rssA2 mutant (Fig. 5). 12

Strikingly, in the absence of SprE, strains harboring pBMK28 did not show this defect (Fig. 5). 13

Note that the uninduced levels of PAP I-His6 produced under these conditions are not high 14

enough to cause the lethality associated with high-level overproduction of PAP I (27). We 15

further showed that in a strain that constitutively produces LacIQ (23), the exit defect associated 16

with pBMK28 was also suppressed (data not shown). Overproducing both PAP I and SprE 17

simultaneously resulted in additive effects, where growth had not resumed even after 60 minutes 18

post dilution (Fig. 5). Results presented in this section suggest that proper control of 19

polyadenylation is important for the exit from stationary phase. 20

21

SprE influences PAP I localization in a growth phase-dependent manner 22


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It has been previously reported that the first 24 amino acids of PAP I are sufficient for its 1

localization to the inner membrane (17). We hypothesized that SprE might affect intracellular 2

levels of polyadenylation by altering the cellular localization of PAP I. To test this directly, we 3

constructed a PAP I-GFP fusion protein. The fusion protein was able to complement the plasmid 4

copy-number defect associated with ∆pcnB strains (data not shown) and resulted in a 6-fold 5

increase in levels of polyadenylation (data not shown), demonstrating that the PAP I-GFP protein 6

is functional. 7

In a wild-type background during exponential growth, PAP I-GFP localized to the inner 8

membrane in 96.8% of the cells examined (Fig. 6A) as demonstrated by a ring of fluorescence at 9

the cell periphery. It should be noted that in many cases, in addition to rings, there appeared to 10

be localized foci of PAP I-GFP at the membrane. During stationary phase PAP I-GFP was no 11

longer membrane localized, but formed distinct foci in the cytoplasm in 94.4% of cells (Fig. 6B). 12

The spatial differences in PAP I localization may be important for regulating PAP I activity. In 13

sprE, rpoS and rssA2 strains, the PAP I-GFP localization in exponential phase was similar to that 14

observed in wild type; around 95% cells showed membrane localization (Fig. 6A). The rssA2 15

and rpoS strains appeared similar to wild-type in stationary phase as well. However, PAP I-GFP 16

was still localized to the membrane in 64.9% of cells in the sprE null strain in stationary phase 17

(Fig. 6B). This suggests that under these conditions, SprE is required for efficient release of 18

PAP I into the cytoplasm. Thus SprE affects not only the activity of PAP I in exponentially 19

growing cells, but also its cellular location in stationary phase cells. 20

21

Discussion 22


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It has been reported that the overproduction of wild-type SprE is toxic to cells (34, 43). 1

We have exploited the growth defect caused by SprE overproduction to uncover a novel function 2

of SprE. Our data indicate that in exponentially growing cells SprE stimulates polyadenylation, 3

thereby affecting the decay of specific mRNA species. Indeed, the SprE60 mutant protein 4

increases levels of polyadenylation at least two-fold and poly(A) levels are reduced two-fold in 5

the absence of SprE (Fig. 2). Thus SprE has two distinct functions, both of which affect the 6

transcriptome. SprE controls what is made by regulating the stability of RpoS and it also 7

controls which mRNAs are destroyed by stimulating polyadenylation. 8

While the exact changes in the polyadenylation profile of RNA substrates in the presence 9

of SprE60, or in the absence of SprE, are not yet known, we assume that these changes reflect 10

changes in the activity of PAP I. In support of this idea, removal of the SprE protein led to 11

almost identical changes in mRNA half-lives as that obtained upon inactivation of PAP I (Table 12

3). However, not all of the decay intermediates observed in the absence of PAP I were present in 13

the SprE mutant (Fig. 3). For example, out of the two yfiA decay intermediates that accumulated 14

in the ∆pcnB strain, only the larger species was stable in the ∆sprE mutant (Fig. 3). This could 15

indicate that the effect of SprE on mRNA decay is dependent on PAP I-mediated 16

polyadenylation as well as some other PAP I-independent mechanism. Furthermore, we cannot 17

rule out that the strong effect of both sprE and pcnB mutations on the ompA mRNA half-life 18

(Table 3) is due to a combination of effects related to other factors such as growth rate, Hfq 19

modulation, and the action of small RNA regulators that affect RNase E cleavage of this 20

transcript (35, 40, 47, 48). Nonetheless, our data strongly suggest that SprE and PAP I function 21

together during exponential phase to regulate polyadenylation and mRNA stability. 22


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Interestingly, connections between SprE and PAP I have been previously reported. 1

Sarkar et al. (43) identified SprE as a multi-copy suppressor of the pcnB plasmid copy-number 2

defect. PAP I controls the levels of an anti-sense RNA (RNA I) that inhibits an RNA replication 3

primer of ColE1-type plasmids (14). In the absence of pcnB, RNA I is stabilized, which prevents 4

plasmid replication. Overproduction of SprE suppresses this defect. Given our results, it seems 5

likely that the overproduction of SprE suppressed the pcnB defect either by stimulating PAP I-6

independent polyadenylation or by stimulating the degradation of the non-polyadenylated RNA I 7

species. One possibility is that SprE stimulates the other known polyadenylation enzyme 8

PNPase, which accounts for the residual polyadenylation in a pcnB null strain in exponential 9

phase cells (31). While it is true that we did not observe a significant increase in PNPase-10

dependent polyadenylation by SprE60 in the ∆pcnB background (Fig. 2), this result may simply 11

be due to the fact that the heteropolymeric tails generated by PNPase (32, 33) cannot be 12

efficiently detected with RNA-DNA dot blots using the oligo(dT)20 probes we employed. 13

Santos et al. (42) noted that in the absence of PAP I, RpoS levels were reduced 3-fold in 14

stationary phase cells. They also demonstrated that the effect seen was post-transcriptional and 15

specifically affected protein stability. They hypothesized that PAP I controls SprE’s activity, 16

and that is the cause for reduced RpoS levels. Taking into account our findings, it is plausible 17

that the reduction of RpoS levels the authors reported was due to the lack of SprE’s second 18

function. Perhaps when there is no PAP I, higher levels of SprE are available to promote RpoS 19

degradation. Indeed the rssA2::cam allele, which increases SprE levels four-fold, decreases 20

RpoS levels in all growth phases (39). What the authors perceived as PAP I regulation of SprE 21

may actually be a titration of SprE into the polyadenylation pathway. 22


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PAP I and SprE clearly function together during exponential phase, and we believe this is 1

true during stationary phase as well. Polyadenylation and mRNA stability in stationary phase 2

have not been extensively studied, and remain poorly understood. However, we believe that 3

SprE must play some role in regulating polyadenylation in stationary phase, since the 4

localization of PAP I changes in a SprE-dependent manner when growth ceases. Moreover, this 5

localization defect is associated with clear phenotypic changes. 6

In an otherwise wild-type background, PAP I-GFP was localized to the inner membrane 7

during exponential phase and was released from the membrane as cells entered stationary phase 8

(Fig. 6). This release of PAP I-GFP from the membrane was SprE-dependent; in cells lacking 9

SprE, PAP I-GFP remained membrane associated. These SprE-dependent changes in the 10

localization of PAP I-GFP provide evidence for a functional connection between PAP I and SprE 11

during stationary phase. It should be noted that the hybrid gene producing PAP I-GFP was 12

carried on a multi-copy plasmid, and thus the fusion protein is overproduced compared to the 13

wild-type endogenous PAP I. This overproduction facilitated visualization of the fluorescent 14

molecule. We assume, but have not proven, that this change in PAP I-GFP localization reflects 15

what happens with endogenous PAP I. If this is true then it seems likely that during exponential 16

phase, efficient polyadenylation takes place at the inner membrane. It is also possible that the 17

entire mRNA degradation process is compartmentalized. Indeed, it has been recently reported 18

that the mRNA degradosome is anchored to the inner membrane by the N-terminal portion of 19

RNase E (18). 20

It is important to note that the cells overproducing PAP I-GFP (Fig. 6) exhibit a defect in 21

the exit from stationary phase that is similar to that caused by overproducing PAP I-His6 (Fig. 5). 22

We believe that this exit defect is caused by the elevated activity of PAP I because removing 23


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SprE suppresses this exit defect. We have shown that removing SprE decreases PAP I activity in 1

exponential phase cells (Fig. 2), and we assume its removal has similar effects on PAP I-GFP 2

and PAP I-His6 in stationary phase as well. Strikingly, in reciprocal fashion, increased 3

production of SprE also causes a defect in the exit from stationary phase, and removal of PAP I 4

suppresses this defect. We assume that overproduction of SprE in stationary phase cells also 5

increases PAP I activity. Indeed, SprE60 increases polyadenylation in exponential phase cells 6

(Fig. 2). However, we do not know if the elevated activity of PAP I that occurs in stationary 7

phase cells under conditions of either PAP I or SprE overproduction is due to changes in PAP I 8

localization, activity, or both. Nonetheless, this reciprocal suppression provides additional 9

evidence for a functional connection between SprE and PAP I. 10

As noted above, little is known about polyadenylation and mRNA stability in stationary 11

phase cells, and we do not currently understand why increasing PAP I activity, either by 12

increasing production of PAP I or SprE, would cause a stationary phase exit defect. It seems 13

likely that this exit defect results from alterations in stationary phase polyadenylation. It is 14

possible that altered polyadenylation results in the depletion of an essential mRNA or non-15

coding RNA, or that the tRNA pool is damaged and therefore depleted. In wild-type cells, a 16

reduction of PAP I activity during stationary phase may help change the polyadenylation profile 17

of mRNA’s, and therefore greatly change which ones are degraded. The RNAs that are degraded 18

during exponential phase may be quite different from those that are degraded during stationary 19

phase. In fact, the stationary phase specific mRNAs generally contain polynucleotide tails (9), 20

which are most probably generated by PNPase (32), and the tails generated by PNPase are poor 21

substitutes for poly(A) tails (33). 22


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Upon entry into stationary phase, SprE levels are increased 2-3 fold (41). It is somewhat 1

paradoxical that SprE levels would increase when the protein is inactive with regard to RpoS 2

degradation. One proposed explanation is that increasing SprE levels in stationary phase is 3

necessary to promote the rapid destruction of RpoS when cells encounter nutrients and return to 4

exponential phase. This explanation seems unlikely because cells that only mildly overproduced 5

SprE exhibited a defect in the exit from stationary phase. Alternatively, we suggest that SprE 6

levels increase because the protein has an additional function in stationary phase, the regulation 7

of polyadenylation. While we currently have no evidence for a direct interaction between SprE 8

and PAP I, SprE clearly plays a role in the PAP I-pathway in exponential phase, and we have 9

documented numerous genetic interactions between sprE and pcnB that affect both exponential 10

and stationary phase phenotypes. Further studies are required to elucidate the exact mechanism 11

by which SprE contributes to the polyadenylation pathway. 12

13

Acknowledgements 14

This work was supported by grants from the National Institute of General Medical Sciences 15

(GM65216) to T.J.S. and (GM57220) to S.R.K. 16

17

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52. Zhou, Y., S. Gottesman, J. R. Hoskins, M. R. Maurizi, and S. Wickner. 2001. The 1

RssB response regulator directly targets sigma(S) for degradation by ClpXP. Genes Dev 2

15:627-37. 3

4

Figure Legends 5

Figure 1. The pcnB and hfq null alleles do not suppress the toxicity phenotype by lowering SprE 6

levels. Cells were grown to mid-exponential phase, and Western blot analysis was performed as 7

described in Materials and Methods. (A) Lane 1 (CNP108), Lane 2 (CNP153), Lane 3 (VC30), 8

Lane 4 (VC35), Lane 5 (VC56). Cultures were induced with 200 ng/mL ATC (lane 4) or 0.2% 9

arabinose (lane 5). (B) Lane 1 (VC35) Lane 2 (MUT60A), Lane 3 (VC179), Lane 4 (VC90), 10

Lane 5 (VC180), Lane 6 (CNP135). Abbreviations: Anhydrotetracycline (ATC). 11

Figure 2. SprE60 and deletion of sprE have opposite effects on intracellular levels of poly(A) 12

during exponential phase. DNA-RNA dot blots were performed as described in Materials and 13

Methods. All values are normalized to the wild-type level of poly(A) in exponential phase cells. 14

Figure 3. Decay of yfiA mRNA in the wild-type, ∆pcnB and ∆sprE strains. Total RNA (10 15

µg/lane) isolated at various time points after addition of rifampicin to stop the transcription were 16

separated on a 6% polyacrylamide/8M urea gel, transferred to a charged nylon membrane and 17

probed with 32P-labeled yfiA DNA as described in Materials and Methods. The figure is a 18

composite from two separate gels to accommodate all the lanes. The half-life of the full-length 19

transcript for is reported in Table 3. The arrows represent yfiA degradation intermediates. 20

Figure 4. Determination of (A) PAP I and (B) RNase II levels in the wild-type (MC4100) and 21

∆sprE (CNP58) strains. ∆pcnB (SK9176) and ∆rnb (CMA201) strains were included as 22


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controls. Cell extracts (50 µg for PAP I and 25 µg for RNase II per lane) were prepared and 1

separated on a SDS/10% polyacrylamide gel as described previously (30). The immunoblots 2

were probed with PAP I and RNase II specific antibodies using ECL Western blotting detection 3

reagents (GE Healthcare). The protein bands were detected and quantified with a 4

PhosphorImager (Storm 840 PC, GE Healthcare) using ImageQuant 5.2 software. RQ represents 5

average of 3-4 independent experiments. The protein level of the wild-type strain was set at one. 6

A star indicates non-specific hybridization. Both A and B are part of larger gels where non-7

relevant lanes were removed. 8

Figure 5. Overproduction of SprE or PAP I causes a defect in the exit from stationary phase. 9

Overnight cultures were back-diluted 1:100, and growth was monitored for 1 hour. MC4100 10

(closed squares), CNP108 (closed triangles), VC239 (open squares,), VC251 (closed circles), 11

VC221 (open triangles), VC253 (open circles). 12

Figure 6: SprE affects the localization of PAP I in a growth phase-dependent manner. Cells 13

were grown to exponential (A) or stationary phase (B), and were prepared for microscopy as 14

described in Materials and Methods. Since all 4 strains showed similar localization in log phase, 15

wild-type is depicted in panel A, as a representative localization pattern. The ∆sprE strain is 16

depicted as a representation in panel B, since both of the two types of stationary phase 17

localization patterns were observed. VC239 (wild type), VC244 (∆sprE), VC253 (rssA2), 18

VC252 (∆rpoS). 19

20

21

22


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Table 1. Strains and plasmids used in this study 1

Strain Genotype Source or

Reference

MC4100 F- araD139 ∆(argF-lac)U169 rpsL150 relA1

flbB5301 deoC1 ptsF25 rbsR

(10)

MG1693 F- thyA715 λ- rph-1 (2)

CNP58 MC4100 ∆sprE::tet This study

CNP108 MC4100 (λrpoS750’-‘lacZ) rssA2::cam This study

CNP135 MC4100 ∆hfq::kan This study

CNP153 MC4100 (λrpoS750’-‘lacZ) sprE::tet (37)

CMA201 MG1693 ∆rnb::tet (38)

ECK4175

MUT60A

BW25113 rnr::kan

VC30/pZS*11sprE60

(3)

This study

SK5665 MG1693 rne-1 (2)

SK7622 MG1693 rnc∆38::kan (4)

SK9176 MC4100 ∆pcnB::kan This study

SK10019 MG1693 pnp∆683::str/spc (28)

SK10486 MC4100 clpX::kan This study

SK10490 MC4100 rpoS::kan This study

SK10503 MC4100/pZS*11sprE60 This study

SK10505 MC4100/pZS*11sprE60 ∆pcnB::kan This study

SK10506 MC4100/pZS*11sprE60 hfq-1 This study


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TX2869 MC4100 hfq-1 (46)

VC30 MC4100 λφ(rpoS750’-‘lacZ) cysC::Tn10 This study

VC35 VC30/pZS*11sprE+ This study

VC56 MC4100 araR/pBADsprE+ This study

VC90 MC4100 ∆pcnB::kan This study

VC91 MC4100/pCMW1 This study

VC179 VC30/pZS*11sprE60

∆pcnB::kan This study

VC180 VC30/pZS*11sprE60 ∆hfq:kan This study

VC181 VC30/pZS*11 This study

VC218 VC30/pZS*11sprE60D58A This study

VC220 MC4100/pBMK28 This study

VC221 CNP58 sprE::tet pBMK28 This study

VC229 VC30/pDK24 (rnb+ Apr) This study

VC230 VC30 /pZS*11sprE+ (Kmr)/ pDK24 (rnb

+ Apr) This study

VC231 VC30/pZS*11sprE60 (Kmr), pDK24 (rnb+ Apr) This study

VC239 MC4100/ ppcnB-GFP This study

VC241 VC30 ∆pcnB::kan / pZS*11sprE+ This study

VC244 MC4100 ∆sprE::tet /ppcnB-GFP This study

VC247 MC4100 pcnB80::Tn10/pCMW1 This study

VC248 MC4100 pcnB80::Tn10/ppcnB-GFP This study

VC251 MC4100 rssA2::cam ∆pcnB::kan This study

VC252 MC4100 rpoS::Tn10/ppcnB-GFP This study

VC253 MC4100 rssA2::cam/ppcnB-GFP This study


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VC256

VC272

MC4100 ∆hfq::kan/ ppcnB-GFP

MC4100 rnr::kan

This study

This study

Plasmid Resistance Marker Source or reference

pZS*11 ApR (23)

pZS*11sprE+ ApR, KmR This study

pBADsprE+ ApR This study

pBMK28 CmR (33)

pDK24 ApR (11)

pCMW1 KmR (49)

ppcnB-GFP KmR This study

1

2

3

4

5

6

7

8

9

10

11

12

13


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Table 2. Growth rate and SprE levels in various strains 1

Genotype Doubling time (min)a SprE levelb

sprE+ 29.4 ± 2.6 1

rssA2 31.5 ± 2.9 4.1 ± 0.1

∆hfq 29.1 ± 3.7 2.2 ± 0.2

∆pcnB 28.8 ± 1.8 1.5 ± 0.3

sprE+/ pZS*11 33.8 ± 4.3 1

sprE+/ pZS*11sprE

+ 40.6 ± 4.5 5.2 ± 0.2

∆pcnB/ pZS*11sprE+ 32.6 ± 0.9 ND

sprE+/ pZS*11sprE60 61.8 ± 8.6 5.0 ± .1

∆pcnB/ pZS*11sprE60 35.4 ± 4.6 5.3 ± 0.1

∆hfq/ pZS*11sprE60 38.6 ± 2.8 5.0 ± 0.1

sprE+/ pZS*11sprE

60D58A 48.0 ± 5.0 ND

sprE+/pDK24 (rnb

+) 30.4 ± 1.8 ND

sprE+/ pZS*11sprE

+/ pDK24 (rnb

+) 47.7 ± 3.8 ND

sprE+/ pZS*11sprE60/ pDK24 (rnb

+) 47.9 ± 4.6 ND

a Three independent growth curves were measured, and doubling times were calculated as 2

described in Materials and Methods. 3


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bSprE protein levels were determined by Western blot analysis by normalizing levels to wild-1

type and averaging values from 4 independent experiments. A representative blot is shown in 2

Fig. 1. ND = not done 3

4

Table 3. Half-lives of specific mRNAs 5

Half-lives (min)

mRNA Wild-type ∆pcnB ∆sprE

rpsO

P1-RIII

P1-t1

1 ± 0.2

1.2 ± 0.2

1.9 ± 0.2

1.7 ± 0.1

2.0 ± 0.2

2.6 ± 0.0

yfiA 5.0 ± 1 6.3 ± 0.3 7.4 ± 0.5

gltA 2.0 ± 0.2 2.5 ± 0.2 3.0 ± 0.5

cspE 4.8 ± 0.2 5.3 ± 0.2 5.4 ± 0.2

ompA 7.0 ± 0.6 12 ± 2 11 ± 1

rplY 4.2 ± 0.1 4.3 ± 0.1 4.6 ± 0.2

Half-lives were determined as described in the Materials and Methods and represent the average 6

of at least two independent determinations. 7

8


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Figure 1

(A)

(B)

sprE

+ (pZS

*11s

prE60

)

sprE

+ (pZS

*11s

prE

+ )

∆

pcnB

(pZS

*11s

prE

60)

∆

pcnB

∆

hfq

(pZS*1

1spr

E60

)

∆

hfq

654321

SprE

ATC ++ + +

sprE

+(p

ZS

*11sp

rE+ )

Wild

typ

e

rssA

2

∆

sprE

4321sp

rE+ (p

BA

Dsp

rE+ )

5

Inducer - - - ++

SprE

--


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Figure 2

1

2

3

Wild

type

∆sprE

sprE

+ (pZS*1

1spr

E60)

∆pcnB

∆pcnB

(pZS*1

1spr

E60)

hfq-

1

hfq-

1 (p

ZS*11s

prE60

)

∆clpX

Re

lative

po

ly(A

) le

ve

ls


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Figure 3

0 1 2 4 8 16 0 1 2 4 8 16 0 1 2 4 8 16Time

(min)

Wild type ∆pcnB ∆sprE


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Wil

d t

yp

e

∆s

prE

∆p

cn

B

RQ 1 1+0.2 -

PAP I

**

(A)

RQ 1 1.3+0.2 -

Wil

d t

yp

e

∆s

prE

∆rn

b

RNase II

*

(B)

Figure 4


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0.05

0.10

0.15

0 20 40 60

Time (min)

OD

600

Wild type

rssA2

rssA2 ∆pcnB

ppcnB-his6

ppcnB-his6 ∆sprE

ppcnB-GFP rssA2

Figure 5


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Figure 6Not Membrane LocalizedMembrane Localized

Wild type

∆rpoS

∆sprE

rssA2

96.8%

95.1%

93.7%

95.0%

3.20%

4.90%

6.30%

5.00%

(A)

(B)

Not Membrane LocalizedMembrane Localized

Wild type

∆rpoS

∆sprE

rssA2

5.60%

2.60%

3.00%

64.9%

94.4%

97.4%

97.0%

35.1%

Exponential

Stationary


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