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The Pennsylvania State University
The Graduate School
Eberly College of Science
REGULATION AND PHYSIOLOGICAL ROLE
OF SSRA RNA
A Thesis in
Biochemistry, Microbiology, and Molecular Biology
by
Sue-Jean Hong
Submitted in Partial Fulfillment of the Requirements
for the Degree of
Doctor of Philosophy
August 2005
The thesis of Sue-Jean Hong was reviewed and approved* by the following:
Kenneth C. Keiler Assistant Professor of Biochemistry and Molecular Biology Thesis Adviser Chair of Committee
Philip C. Bevilacqua Associate Professor of Chemistry Craig E. Cameron Professor of Biochemistry and Molecular Biology Davis T. W. Ng Associate Professor of Biochemistry and Molecular Biology B. Tracy Nixon Associate Professor of Biochemistry and Molecular Biology Robert A. Schlegel Professor of Biochemistry and Molecular Biology Head of the Department of Biochemistry and Molecular Biology
*Signatures are on file in the Graduate School.
iii
ABSTRACT The last decade has witnessed a renaissance in the field of small regulatory RNAs.
All organisms ranging from bacteria to human contain a wealth of small regulatory RNAs
that function in a variety of cellular processes. The small regulatory RNAs are involved
in regulation of gene expression at both transcriptional and post-transcriptional levels, by
modifying chromatin structure, modulating transcription factor activity, and influencing
mRNA stability, processing, and translation.
One of the most interesting of these small regulatory RNAs is SsrA RNA. SsrA
with properties of both a tRNA and an mRNA carries out an extraordinary trans-
translation reaction that is ubiquitous in bacteria. In cases where a ribosome is arrested on
a selected mRNA, SsrA is recruited to the A site of the ribosome. By an unknown
mechanism, SsrA causes the ribosome to release the mRNA and resume translation using
a short open reading frame encoded within SsrA. This process serves to release the
stalled ribosome and adds a peptide tag to the end of the nascent polypeptide marking it
for degradation. This unique activity is required for such cellular processes as growth and
development, pathogenesis, symbiosis, and stress tolerance.
This study focuses on the regulation and physiological function of SsrA using
Caulobacter crescentus as a model organism. Asymmetric cell division and
differentiation of C. crescentus and the extensive knowledge of the molecular events
associated with the cell cycle provide an unique opportunity to study the influence of
SsrA activity on cell physiology. In C. crescentus, the initiation of chromosomal
replication is delayed during G1-S transition in the absence of SsrA activity. SsrA is also
iv
required for plasmid replication. The steady-state levels of SsrA is cell cycle regulated
such that its expression is high in both G1 and G2 phases but low in S phase.
Through detailed analysis of effects of two highly conserved proteins, RNase R
and SmpB, on the regulation of the abundance of SsrA, it has been demonstrated that
SsrA is specifically degraded by RNase R at a specific point in the cell cycle and this
timing may be regulated by SmpB. Proteomic studies of cellular substrates of SsrA reveal
that proteins in diverse functional categories are tagged by SsrA and at least one
consensus DNA motif exists that may activate the SsrA system. Genetic and proteomic
analyses suggest that SsrA may control plasmid replication by regulating the replication
initiation protein, Rep. Taken together, this study establishes a foundation for a more
comprehensive understanding of the regulation and physiological role of the small
regulatory RNA, SsrA.
v
TABLE OF CONTENTS
List of Figures vii List of Tables viii Acknowledgements ix Chapter 1. GENERAL INTRODUCTION: SSRA RNA IN EUBACTERIA 1
Introduction 2 SsrA RNA: a hybrid of tRNA and mRNA 2 Physiological roles of SsrA 9 SsrA associated protein factors 10
Model system: Caulobacter crescentus 12 SsrA in C. crescentus 13 References 16
Chapter 2. CELL-CYCLE REGULATED DEGRADATION OF SSRA IS 28 CONTROLLED BY RNASE R AND SMPB Abstract 29
Introduction 30 Results 34
C. crescentus RNase R specifically degrades SsrA RNA in vitro 34 RNase R is required for cell-cycle dependent degradation of SsrA RNA 36 Lack of RNase R alters cell-cycle expression of ssrA 37 Phenotype of the rnr deletion strain 38 SmpB protects SsrA RNA from RNase R degradation 39 Discussion 41 Materials and Methods 45 References 52 Chapter 3. PROTEOMIC STUDIES OF PHYSIOLOGICAL SUBSTRATES FOR 67 THE SSRA SYSTEM IN CAULOBACTER CRESCENTUS Abstract 68
Introduction 69 Results 72
Construction of functional SsrA-His6 72 Proteomes of SsrA tagging in C. crescentus 73 Identification of tagged proteins 73 Functional diversity of SsrA-tagged proteins 75 mRNA levels of SsrA-tagged genes 77 SsrA tagging sites and substrate selectivity 78 Discussion 80 Materials and Methods 83 References 86
vi
Chapter 4. CAULOBACTER CRESCENTUS REQUIRES SSRA ACTIVITY 109 FOR PLASMID REPLICATION
Abstract 110 Introduction 111 Results 113
Plasmid maintenance in ssrA deficient strains 113 Selection for mutants that can maintain plasmids in the absence of 114 SsrA activity SsrA tags Rep at the C terminus 116 Regulation of Rep by SsrA tagging 117
Discussion 120 Materials and Methods 123 References 129
vii
LIST OF FIGURES Figure 1.1. Schematic representation of SsrA from E. coli 24 Figure 1.2. Model for SsrA tagging mechanism 25 Figure 1.3. Cell cycle progression of C. crescentus 26 Figure 1.4. Schematic representation of transcription and maturation of SsrA in C. crescentus 27 Figure 2.1. RNase R activity in vitro 59 Figure 2.2. Degradation of SsrA RNA in wild-type and ∆rnr strains 60 Figure 2.3. Expression of SsrA RNA in the ∆rnr strain 61 Figure 2.4. Cell-cycle regulation of RNase R and SmpB protein levels 62 Figure 2.5. Binding kinetics of purified SmpB to SsrA RNA 63 Figure 2.6. Model for regulation of SsrA RNA by RNase R and SmpB 64 Figure 3.1. Schematic representation of wild-type SsrA and SsrA-His6 92 Figure 3.2. Detection of SsrA-His6-tagged proteins 93 Figure 4.1. Colony forming units of plasmids in C. crescentus strains 132 Figure 4.2. Schematic representation of a genetic selection for plasmids bypassing the requirement of SsrA activity 133 Figure 4.3. Properties of plasmid pPT1 and construction of variant pKJS2 plasmids 134 Figure 4.4. MALDI-TOF analysis of SsrA-tagged Rep 135 Figure 4.5. Overexpression of Rep does not harm cells 136 Figure 4.6. Expression of Rep in wild-type and ∆ssrA C. crescentus 137
viii
LIST OF TABLES Table 2.1. Growth parameters of wild type and ∆rnr strains 65 Table 2.2. SmpB specifically inhibits RNase R degradation of SsrA RNA 66 Table 3.1. Identified SsrA-tagged proteins and tagging determinants 94 Table 3.2. List of SsrA-tagged proteins 95 Table 3.3. Functional distribution of identified substrates 101 Table 3.4. Occurrence of a putative motif inducing SsrA tagging 102
ix
ACKNOWLEDGEMENTS
It has been my great honor and privilege to learn scientific thinking from my
advisor, Dr. Kenneth C. Keiler. During my short stay with him, I have learned the art of
performing science and presenting it in an exciting manner. His enthusiasm and strong
motivation to explore challenging questions has greatly shaped and broadened my
perspective of science. Thanks to him, I feel confident in accepting scientific challenges
and venturing into new fields of research. His words of encouragement always motivated,
inspired, and helped me to view matters from a different perspective. I am greatly
indebted to him for his guidance, support, and most of all patience.
During my graduate career, I have been fortunate to have a second advisor. With
all my heart, I would like to express my appreciation to my previous advisor, Dr. Susan
M. Abmayr for her guidance, support, and scientific advice. Her willingness to educate
me to understand the fundamental concepts of scientific research will always remain with
me. I am very grateful to all my committee members, Dr. Davis Ng, Dr. Craig Cameron,
Dr. Tracy Nixon, and Dr. Philip Bevilacqua for their helpful discussions and guidance. I
would like to especially thank Dr. Tracy Nixon for creating the proteomic database,
which has been an indispensable part for the Chapter 3 of this thesis.
I am very thankful to the past and present Keiler lab members, Dr. Faith Harrison,
Lin Cheng, Dennis Lee, Ling Li, Jay Russell, BryanVenters, Monica Guo, Jake Wesley,
Anastasiya Yakhnin, Iza Petrykowska, Quyen-Anh Tran, Tomoyo Takagi, and Priscilla
Tee for their support and helpful discussions. They certainly made the Keiler lab to be the
most exciting place to carry out science. I would like to express my gratitude to my
x
previous lab members, Dr. Lakshmi Balagopalan, Dr. Rhakee Banerjee, and the late
Malabika Chakravarti for their support and scientific advice at the beginning of my
graduate career.
I am very grateful to all my friends and colleagues in BMB department for their
friendship, help, and support.
My family has provided me a tremendous amount of encouragement and love
during my time in graduate school. I am indebted to my brother, Sung-Sul and his family,
Kum-Ok, Dennis, and Karen for their love and support both emotionally and financially.
I am also indebted to my sister, Eun-Hwa who has been with me in every step of my
graduate career for her endless love and encouragement. I am grateful to her for teaching
me the value of optimism.
xi
In loving memories of my mum and dad,
1
Chapter 1
General Introduction:
SsrA RNA in Eubacteria
2
A. Introduction
When cells face suboptimal chemical and physical conditions, global regulatory
networks must be coordinated to optimize the use of available nutrients and to increase
the chance of survival. In addition to protein regulators, small regulatory RNAs in many
cases add another layer of regulation to these systems. It is now evident that small
regulatory RNAs functioning in a variety of physiological conditions are present in both
prokaryotes and eukaryotes (Bartel, 2004; Gottesman, 2004). In prokaryotes, small
regulatory RNAs function by three general mechanisms. First, direct interaction with and
modification of a protein can directly affect gene expression (Romeo, 1998; Wassarman
and Storz, 2000). Second, antisense base-pairing to target mRNAs alters the structure
and/or stability of the messages, resulting in inhibition or stimulation of ribosome
binding, and ultimately changing translational efficiency (Gottesman, 2004; Storz et al.,
2004). SsrA RNA, a small RNA regulator in the third class acts by trans-translation
(Keiler et al., 1996). Although SsrA RNA is widely distributed in every bacterial species
(Gueneau de Novoa and Williams, 2004; Keiler et al., 2000), it is largely unknown how it
is regulated and what its biological role is. The function and regulation of this SsrA RNA
in Caulobacter crescentus is the focus of this study.
B. SsrA RNA: a hybrid of tRNA and mRNA
SsrA (small stable RNA A) was originally identified in Escherichia coli as part of
the 10S RNA (10Sa RNA) fraction (Lee et al., 1978; Ray and Apirion, 1979; Subbarao
and Apirion, 1989). ssrA has subsequently been found in every bacterial genome, in some
plastid genomes, and in some mitochondrial genomes (Gueneau de Novoa and Williams,
3
2004; Keiler et al., 2000). In E. coli, SsrA is encoded by a single gene (Chauhan and
Apirion, 1989; Komine et al., 1994). Transcription of ssrA appears to be controlled by a
σ70-type promoter and an ρ-independent terminator to yield a 457-nucleotide precursor
(pSsrA) (Komine et al., 1994). This precursor is then processed at both ends to create a
363-nucleotide mature form. In E. coli, RNase P removes seven nucleotides from the 5’
end of the pSsrA to generate the mature 5’ terminus (Komine et al., 1994). The mature 3’
end of E. coli SsrA is generated by an initial RNase III and/or RNase E cleavage
followed by RNase T and RNase PH-mediated trimming (Li et al., 1998; Lin-Chao et al.,
1999; Srivastava et al., 1992; Srivastava et al., 1990). SsrA is a unique tRNA-mRNA
hybrid (and hence, it is also named as tmRNA, Figure 1.1). It contains two distinct
functional domains, a tRNA-like domain that mimics part of alanyl-tRNA and an mRNA-
like domain that encodes a short polypeptide.
B1. tRNA-like domain
The potential tRNA-like activity of SsrA was first suggested by comparative
sequence analysis of SsrAs from E. coli, Alcaligenes eutrophus, Mycobacterium
tuberculosis, Bacillus subtilis, and Mycoplasma capicolum (Komine et al., 1994; Ushida
et al., 1994). The 5’ and 3’ ends of the RNA base-pair forming a tRNA-like domain with
an acceptor stem with a 3’-end CCA aminoacylation sequence and a TΨC arm (Figure
1.1; Komine et al., 1994). As observed in alanyl-tRNA, the acceptor stem of SsrA also
contains a G:U wobble at the third base pair position, which is recognized by analyl-
tRNA synthetase (Komine et al., 1994). Further evidence for tRNA-like functionality
4
was provided by in vitro assays demonstrating that SsrA can be charged with alanine by
alanyl-tRNA synthetase (Komine et al., 1994; Ushida et al., 1994).
B2. mRNA-like domain
Instead of an anticodon stem-loop observed in a canonical tRNA, the
corresponding stem in SsrA is extended and closed by a large loop of RNA (Figure 1.1;
Williams and Bartel, 1996). Within this loop, there is an mRNA-like domain that
contains a short open reading frame (ORF). A translated tag peptide from this ORF was
first discovered at the C termini of truncated versions of the murine interleukin-6 (IL-6)
overexpressed in E. coli (Tu et al., 1995). However, the tag peptide from SsrA ORF was
not observed when IL-6 was overexpressed in ∆ssrA E. coli, suggesting that SsrA
functions as an mRNA for the tag peptide (Tu et al., 1995).
B3. A trans-translational model for SsrA activity
The most important evidence for SsrA acting both as a tRNA and an mRNA was
provided by the trans-translational model for SsrA activity as part of the mechanism for
translational quality control (Figure 1.2; Keiler et al., 1996). This model was proposed
based on the similarity observed between the hydrophobic C-terminal amino acid
sequence of the ssrA-encoded tag peptide and that recognized by a periplasmic protease,
Tsp (Keiler et al., 1996). Normally, translation stops when the ribosome encounters a
stop codon in an mRNA, releasing the finished polypeptide and recycling the ribosome.
If, however, ribosomes translate to the 3’ end of truncated transcripts lacking stop
codons, they cannot dissociate from the transcripts. By an unknown mechanism, SsrA
5
charged with alanine is recruited to the A site of the trapped ribosomes and accepts the
nascent polypeptide by transpeptidation. Translation then shifts from the original mRNA
to the ORF within SsrA and normal translation resumes from this ORF. Termination at a
stop codon within the SsrA ORF results in release of trapped ribosomes and a
polypeptide with an SsrA-encoded peptide tag at its C terminus. The SsrA tag directs the
polypeptide for rapid degradation by periplasmic protease Tsp, cytoplasmic protease
complexes ClpXP and ClpAP, and membrane-anchored protease HflB/FtsH (Gottesman
et al., 1998; Herman et al., 1998; Keiler et al. 1996).
Many aspects of the model for SsrA activity have been supported by experimental
data. For example, the λ repressor N-terminal domain and cytochrome b562 translated
from truncated mRNAs lacking in-frame stop codons were tagged at the C termini and
are rapidly degraded (Keiler et al., 1996). Moreover, the SsrA-encoded tag was found in
in vitro translation products in the presence of wild-type SsrA or a variant SsrA (SsrA
encoding proteolysis-resistant tag) and polyuridine or mRNA encoding the N-terminal
domain of λ repressor without a termination codon (Himeno et al., 1997; Roche and
Sauer, 1999). SsrA is associated with 70S ribosomes but not 30S or 50S subunits
(Komine et al., 1996; Tadaki et al., 1996; Ushida et al., 1994) and this association
requires aminoacylation of SsrA with alanine (Tadaki et al., 1996). A recent cryo-
electron microscopy (cryo-EM) structure of SsrA in complex with the ribosome has
provided the first detailed insights into the trans-translational mechanism (Valle et al.,
2003). Using purified components from Thermus thermophilus, a ribosome complex
stalled at the end of a short mRNA was reacted with alanyl-SsrA, SmpB (a protein
required for SsrA activity, Section D), elongation factor Tu (EF-Tu, Section D) in the
6
presence of an antibiotic which prevents dissociation of EF-Tu and thereby halts the SsrA
complex in the ribosomal A site. The cryo-EM images of the complex reveal EF-Tu-
bound tRNA-like domain of SsrA engages the ribosomal A site in a similar manner to
that of normal tRNAs. SmpB makes bridging contacts between the tRNA-like domain of
SsrA and the ribosome. A large part of SsrA loops around the 30S subunit positioning the
mRNA-like domain of SsrA near the normal mRNA entrance to the A site (Valle et al.,
2003).
B4. Identified determinants of SsrA tagging
Understanding whether SsrA is also employed to mediate other types of
translational failures is fundamental to a complete understanding of its biological
function. Moreover, if SsrA can assist with a wider range of translational defects, it is
important to understand what features of such events lead to its recruitment. The trans-
translational model was initially proposed based on one specific condition where
ribosomes are stalled at the end of mRNAs lacking in-frame stop codons (Keiler et al.,
1996). However, recent studies have shown that SsrA can also be recruited by stalled
ribosomes either within an ORF or at termination codons while translating intact mRNAs
(Hayes et al., 2002a; Hayes et al., 2002b; Roche and Sauer, 1999; Roche and Sauer,
2001).
B4a. Incomplete mRNAs without stop codons
Truncated mRNAs without termination codons could result from mRNA damage,
premature transcription termination, or degradation or cleavage by ribonucleases. The E.
7
coli toxins RelE, MazF, and ChpB have been shown to cleave mRNAs in ribosomal A
site and induce SsrA tagging (Christensen and Gerdes, 2003; Christensen et al., 2003;
Pedersen et al., 2003). Ribosome stalling and SsrA-tagging occur during translation of
such mRNAs because release factors are not recruited (Keiler et al., 1996; Roche and
Sauer, 1999; Roche and Sauer, 2001). Tagging of E. coli Lac Repressor is mediated by
this mechanism (Abo et al., 2000). Binding of Lac Repressor to its operators within the
lacI gene blocks RNA polymerase from completing lacI transcription, resulting in
truncated transcripts lacking a stop codon which in turn activate SsrA. In the absence of
ssrA, however, Lac Repressors are produced from the truncated mRNAs and bind to the
lac operators. This increase in operator occupancy leads to a significant delay in
induction of the lac operon in ∆ssrA cells (Abo et al., 2000). This suggests that SsrA
plays an important regulatory role in gene expression by controlling protein levels.
B4b. Complete mRNAs with rare codons
It has been shown that the N-terminal domain of λ repressor translated from
hybrid transcripts containing rare AGA and CGA arginine codon repeats is tagged by
SsrA (Roche and Sauer, 1999). The level of the cognate arginyl-tRNA is the major
determinant for tagging induced by rare AGA codons. Under normal growth conditions,
SsrA tagging is observed only if a transcript contains two or more consecutive codons.
Overexpression of the rare cognate tRNA eliminates tagging at rare codons, whereas
depletion of the cognate tRNA results in tagging at single rare codons (Roche and Sauer,
1999). These results indicate that ribosome stalling at internal sites (clusters of rare
arginine codons) on complete mRNAs induces SsrA tagging at rare codons.
8
B4c. Complete mRNAs with inefficient stop codons and/or specific sense codons
Recent identification of endogenous proteins tagged by SsrA in E. coli reveals
that tagging is affected by the identity of the stop codon and the final sense codons of the
gene (Hayes et al., 2002a; Hayes et al., 2002b; Roche and Sauer, 2001). In E. coli, YbeL,
LacR, GalE, and RbsK are tagged at or near the C terminus (Roche and Sauer, 2001).
SsrA tagging of RbsK (an enzyme catalyzing the conversion of ribose to ribose 5-
phosphate) is determined by the combination of two rare arginine codons (AGG) and an
adjacent inefficient opal (UGA) stop codon (Hayes et al., 2002b). Tagging is observed at
both rare arginine codons and at the opal stop codon. Mutating the rare arginine codons
(AGG) (or inefficient opal stop codon) to the most common arginine codons (CGU) (to a
more efficient ochre (UAA) stop codon) leads to greatly reduced SsrA tagging.
Furthermore, overexpression of the rare cognate tRNA leads to a large decrease in
tagging (Hayes et al., 2002b).
Mutational analyses show that a C-terminal proline residue is the major
determinant for SsrA tagging of YbeL at a stop codon (Hayes et al., 2002a). This is
confirmed by SsrA-tagging of C-terminal proline-substituted thioredoxin, which is not
normally tagged (Hayes et al., 2002a). Tagging of YbeL is not affected by the identity of
the C-terminal proline codon, whereas the incorporation of proline analogs such as
azetidine-2-carboxylic acid, γ-thiaproline, and 3, 4-dehydroproline into YbeL affects the
extent of tagging at a stop codon (Hayes et al., 2002a). These data suggest that YbeL
tagging is largely determined by the chemical nature of the C-terminal proline residue of
the nascent polypeptide.
9
Taken together, these observations suggest that SsrA is capable of resolving a
variety of problems arising during protein synthesis, the common denominator being a
stalled ribosome on a transcript.
C. Physiological roles of SsrA
Genes encoding SsrA have been discovered in all known bacterial genomes and in
some mitochondria and chloroplasts (Gueneau de Novoa and Williams, 2004; Keiler et
al., 2000), suggesting an important SsrA role for cellular physiology. However, it is
largely unknown why it is conserved or what the important role is.
C1. Function in bacterial pathogenesis
In a pathogenic bacterium Neisseria gonorrhoeae, SsrA activity is essential for
viability (Huang et al., 2000). In two additional pathogenic strains, ssrA is dispensable for
normal growth but is required for full virulence (Baumler et al., 1994; Julio et al., 2000).
In Salmonella typhimurium, Tn10 transposon insertion in smpB (Section D) locus results
in a decreased ability to survive within murine macrophages that is correlated with a
reduced virulence in mice (Baumler et al., 1994). In Salmonella enterica serovar
Typhimurium, ssrA deletion results in a defect in virulence in mice and defects in
virulence gene expression (Julio et al., 2000).
C2. Function in α-proteobacterial linage
ssrA in Bradyrhizobium japonicum was identified in a Tn5-mediated genetic
screen for mutations that prevent growth in nodules at the roots of soybean plants
10
(Ebeling et al., 1991). In the absence of ssrA, B. japonicum grows slightly slower than the
wild type under free-living conditions. Furthermore, B. japonicum lacking SsrA activity
displays a severe defect in colonizing the root nodules (Ebeling et al., 1991), suggesting
an important SsrA function in symbiosis. In C. crescentus, SsrA is required for cell cycle
progression by controlling initiation of DNA replication (Section F; Keiler and Shapiro,
2003a).
C3. Other functions under stress conditions
Although ssrA is not essential for E. coli growth in culture, ∆ssrA strains exhibit
slower growth, impaired recovery from carbon starvation (Oh and Apirion, 1991),
reduced growth at 45oC and motility on semisolid agar (Komine et al., 1994), increased
expression or activity of Alp protease (Kirby et al., 1994), and inhibition of phage growth
(Karzai et al., 1999; Retallack et al., 1994). Similar phenotypes are observed for ∆ssrA B.
subtilis. SsrA is required for growth under conditions such as temperature over 40oC and
in elevated concentrations of ethanol or cadmium chloride (Muto et al., 2000).
D. SsrA associated protein factors
Several proteins are necessary for SsrA-mediated tagging. SmpB (Small Protein
B), an essential cofactor for SsrA function, binds specifically to SsrA with high affinity
(Jacob et al., 2005; Karzai et al., 1999; Metzinger et al., 2005; Sundermeier et al., 2005)
and mediates SsrA interaction with the ribosome (Karzai et al., 1999). Recent mutational
analyses of the C-terminal domain of SmpB suggest that SmpB may play a role in
productive positioning of SsrA in the ribosomal A site (Sundermeier et al., 2005). In C.
11
crescentus, SmpB is also required for SsrA stability (Keiler and Shapiro, 2003b). In both
E. coli and C. crescentus, ∆smpB strains display the same phenotypes as those observed
in ∆ssrA strains (Karzai et al., 1999; Keiler and Shapiro, 2003b).
Two general translation factors are also important for SsrA tagging. Alanine-
tRNA synthetase mediates aminoacylation (Komine et al., 1994; Ushida et al., 1994). EF-
Tu protects aminoacyl moiety of alanylated SsrA (Rudinger-Thirion et al., 1999) and
delivers SsrA to ribosome (Valle et al., 2003).
Several other proteins that copurify with SsrA-SmpB have also been identified in
E. coli (Karzai and Sauer, 2001). Ribosomal protein S1 binds to SsrA in vitro and in vivo
(Karzai and Sauer, 2001; McGinness and Sauer, 2004; Wower et al., 2000) and may
facilitate the association of SsrA with the ribosome. RNase R, encoded by rnr, is a highly
conserved exoribonuclease (Zuo and Deutscher, 2001) and degrades many natural RNAs
(Cheng and Deutscher, 2002; Cheng and Deutscher, 2005). In the absence of rnr, higher
levels of SsrA-tagging of natural substrates are observed compared to the isogenic wild
type strain (Karzai and Sauer, 2001). RNase R is required for the expression of the
virulence phenotype of Shigella flexneri and enteroinvasive E. coli (Cheng et al., 1998)
and its expression is induced by ssrA in S. enterica serovar Typhimurium (Julio et al.,
2000). Biochemical interactions are established between SsrA and the phosphoribosyl
pyrophosphate synthetase (PrsA), and SsrA-associated factor (SAF) encoded by the yfbG
gene (Karzai and Sauer, 2001). However, the biological roles of these physical
interactions remain to be elucidated.
12
E. Model system: Caulobacter crescentus
The gram-negative aquatic bacterium C. crescentus undergoes a cell cycle which
is tied to two distinct cell morphologies: swarmer cells and stalked cells (Figure 1.3;
Ryan and Shapiro, 2003). C. crescentus begins the cell cycle as a motile swarmer cell
containing a polar flagellum and pili. The swarmer cell cannot initiate DNA replication
and remains in G1 phase with a single chromosome. The swarmer cell differentiates into
a sessile stalked cell by shedding flagellum and retracting pili. A stalk, which is a narrow
elongation of the cell wall and membranes, is constructed at the same location as the
discarded flagellum. Coincident with this morphological transition, the cell enters S
phase and initiate DNA replication. DNA replication and segregation of daughter
chromosomes to opposite ends of the growing predivisional cell occur during S phase and
a brief G2 phase. Before dividing, the predivisional cell also builds a new flagellum and
pili at the pole opposite the stalk. Once the flagellum is complete, the predivisional cell
divide asymmetrically to yield two daughter cells. One daughter cell is a stalked cell that
immediately reinitiates another round of S phase and the other daughter cell is a swarmer
cell that cannot start DNA replication until after the obligate swarmer- to stalked cell
differentiation step. This entire developmental progression can be followed because it is
easy to isolate a pure population of swarmer cells which go through the cell cycle
synchronously. Furthermore, the complete genome of C. crescentus has been sequenced
(Nierman et al., 2001). These experimental advantages make C. crescentus the simplest
model system for studying the bacterial cell cycle as well as cell cycle regulated events.
13
F. SsrA in C. crescentus
F1. Characteristics of SsrA
There are two significant differences between SsrA from E. coli and C. crescentus
with respect to the composition and regulation of RNA. Most SsrAs, including E. coli
SsrA, consist of a single RNA chain. SsrA in C. crescentus, related α-proteobacteria, and
cyanobacteria is composed of two RNA molecules due to circular gene permutation. That
is, the segment normally at the 3’ end of ssrA is located upstream of the segment
normally at the 5’ end (Keiler et al., 2000).
The ssrA gene in C. crescentus is transcribed as a single RNA, pre-SsrA, which
contains a hairpin loop connecting the tRNA-like 5’ and 3’ ends. This pre-SsrA is then
processed, presumably by the same enzymes used for one-piece SsrA processing (Section
B), to produce mature SsrA composed of a coding RNA and an acceptor RNA (Figure
1.4). This two-piece C. crescentus SsrA can tag proteins translated from a model mRNA
without a stop codon, and these tagged peptides are rapidly degraded in vivo (Keiler et
al., 2000). These data suggest that the two-piece C. crescentus SsrA is functional and acts
in a similar manner to the one-piece SsrA.
The second difference between SsrA from E. coli and C. crescentus is that SsrA
in C. crescentus is cell cycle regulated (Keiler and Shapiro, 2003a). Northern blot
analyses on total RNA isolated from synchronized cultures of C. crescentus show that the
steady-state levels of SsrA increase during the swarmer- to stalked-cell transition and
decrease in stalked cells. Finally, the amount of SsrA increases slowly in predivisional
cells (Keiler and Shapiro, 2003a).
14
This cell-cycle dependent abundance of SsrA is controlled by a combination of
temporally regulated transcription and cell-type specific RNA degradation (Keiler and
Shapiro, 2003a). The ssrA promoter activity was determined by measuring the activity of
a lacZ reporter at different points in the cell cycle. ssrA transcription increases during the
swarmer- to stalked-cell transition, just before the increase in steady-state levels of SsrA.
This increase in transcription can account for the increase in SsrA abundance early in the
cell cycle. However, the rapid decrease in SsrA levels during the stalked-cell stage
suggests that SsrA is degraded. The half-life of SsrA in synchronized cultures is
measured by inhibiting transcription and monitoring the decay of RNA by Northern
blotting. SsrA has a half-life of longer than 50 min. in swarmer cells, but only about 5
min. in stalked cells.
This stalked-cell specific degradation of SsrA suggests that there must be protein
factors responsible for the regulation of SsrA. Investigating the roles of two such
proteins, RNase R and SmpB is the subject of Chapter 2.
F2. Phenotypes of ∆ssrA in C. crescentus
In C. crescentus deleted for ssrA or smpB, there is a specific delay in the
swarmer- to stalked-cell transition resulting in a longer cell-cycle duration compared to
wild type. The cell cycle is delayed because there is about 40 min delay in initiation of
DNA replication, suggesting that SsrA is required for proper timing of DNA replication
(Keiler and Shapiro, 2003b). However, the factors that are responsible for this cell cycle
defect are not known. Identification and characterization of such factors are the focus of
Chapter 3.
15
In addition to the delay in replication initiation, there is a defect in plasmid
replication in the absence of ssrA or smpB (K. C. Keiler, unpublished data). pBBR1-
based broad-host range plasmids are normally replicated in wild type C. crescentus.
However, these plasmids are not maintained in ∆ssrA or ∆smpB strains unless the
plasmid contains a copy of ssrA or smpB. The basis of this defect has not been
determined and characterization of the SsrA-dependent pathway responsible for plasmid
replication is the focus of Chapter 4.
16
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24
tRNA-like domain
mRNA-like domain
Figure 1.1. Schematic representation of SsrA from E. coli. tRNA-like domain is represented by a green line. Red hatched bar represents an open reading frame encoding a proteolysis-inducing tag within the mRNA-like domain, adapted from Keiler KC, Shapiro L, Williams KP (2000) Proc Natl Acad Sci U S A 97:7778-7783.
25
Fi
gure
1.2
. Mod
el fo
r Ssr
A ta
ggin
g m
echa
nism
. A
ribo
som
e sta
lled
on a
n ab
erra
nt m
RNA
recr
uits
alan
yl-S
srA
to th
e rib
osom
al A
site
. The
nas
cent
pol
ypep
tide
chai
n is
then
tra
nsfe
rred
to th
e A
la o
f ala
nyl-S
srA
and
pro
tein
synt
hesis
resu
mes
, but
now
usin
g Ss
rA a
s its
tem
plat
e. S
srA
-tem
plat
ed
elon
gatio
n an
d te
rmin
atio
n re
sult
in th
e rel
ease
of t
he ta
gged
pol
ypep
tide
whi
ch is
subs
eque
ntly
reco
gniz
ed a
nd d
egra
ded
by
prot
ease
s, ad
apte
d fro
m K
eile
r KC,
Wal
ler P
R, S
auer
RT
(199
6) S
cien
ce 2
71:9
90-9
93.
26
Fi
gure
1.3
. Cel
l cyc
le p
rogr
essio
n of
C. c
rece
ntus
. M
otile
swar
mer
cel
ls di
ffere
ntia
te in
to st
alke
d ce
lls a
t the
G1-
S tra
nsiti
on b
y sh
eddi
ng th
eir p
olar
flag
ellu
m, g
row
ing
a sta
lk a
t th
at si
te, a
nd in
itiat
ing
DN
A re
plic
atio
n. D
NA
repl
icat
ion
cont
inue
s as c
ells
elon
gate
and
incr
ease
in m
ass d
urin
g S
phas
e. A
ne
w fl
agel
lum
is a
ssem
bled
in th
e pre
divi
siona
l cel
ls at
the
pole
opp
osite
the
stalk
. On
com
plet
ion
of D
NA
repl
icat
ion,
an
asym
met
ric c
ell d
ivisi
on g
ener
ates
two
new
dau
ghte
r cel
ls (G
2 ph
ase)
. Ele
ctro
n m
icro
grap
hs a
re a
dapt
ed fr
om K
elly
, AJ
(199
8) G
enes
Dev
12:
880-
893.
27
Figure 1.4. Schematic representation of transcription and maturation of SsrA in C. crescentus. The ssrA gene is transcribed as a single RNA, pre-SsrA, in which the tRNA-like 5’ and 3’ ends are connected by a hairpin loop. Processing of the loop by nucleases produce mature SsrA composed of a coding RNA and an acceptor RNA, adapted from Keiler KC, Shapiro L, Williams KP (2000) Proc Natl Acad Sci U S A 97:7778-7783.
28
Chapter 2
Cell-cycle regulated degradation of SsrA is controlled by
RNase R and SmpB
Publication:
Hong, S. J., Tran, Q. A., and Keiler, K. C. (2005). Cell cycle-
regulated degradation of tmRNA is controlled by RNase R and
SmpB. Mol Microbiol 57, 565-575.
29
ABSTRACT
The production and removal of regulatory RNAs must be controlled to ensure
proper physiological responses. SsrA RNA (tmRNA), a regulatory RNA conserved in all
bacteria, is cell-cycle regulated and is important for control of cell cycle progression in
Caulobacter crescentus. We report that RNase R, a highly conserved 3’to 5’
exoribonuclease, is required for the selective degradation of SsrA RNA in stalked cells.
Purified RNase R degrades SsrA RNA in vitro, and is kinetically competent to account
for all SsrA RNA turnover. SmpB, a tmRNA-binding protein, protects SsrA RNA from
RNase R degradation in vitro, and the levels of SmpB protein during the cell cycle
correlate with SsrA RNA stability. These results suggest that SmpB binding controls the
timing of SsrA RNA degradation by RNase R. We propose a model for the regulated
degradation of SsrA RNA in which RNase R degrades SsrA RNA from a non-tRNA-like
3’ end, and SmpB specifically protects SsrA RNA from RNase R. This model explains
the regulation of SsrA RNA in other bacteria, and suggests that a highly conserved
regulatory mechanism controls SsrA activity.
30
INTRODUCTION
Regulatory RNAs play a prominent role in a number of physiological processes in
both prokaryotes and eukaryotes, including regulation of gene expression (Szymanski
and Barciszewski, 2003), remodeling and modification of chromatin structure (Akhtar,
2003), modulating the activity of proteins (Wassarman and Storz, 2000), and controlling
mRNA stability, processing, and translation (Bartel, 2004; Gottesman, 2004; Storz et al.,
2004). In addition, most regulatory RNAs are expressed in highly specific patterns in
relation to development and certain environmental conditions, suggesting that they play
important roles in the cell (Bartel, 2004). The expression of regulatory RNAs under
specific physiological conditions implies that these RNAs are produced and removed in a
coordinated fashion to ensure proper signaling and physiological responses. While there
are many examples of specific production of a regulatory RNA, there are far fewer cases
of specific removal of a regulatory RNA. One example of regulatory RNA degradation is
sRNAs in Escherichia coli. sRNAs base-pair with target mRNAs and attract RNase E for
the coupled degradation of the sRNA and its target. The RNA chaperone Hfq is required
to regulate this process (Gottesman, 2004). Here we report a mechanism for specifically
removing a highly conserved regulatory RNA, SsrA, at a single point in the cell cycle.
Analogous to proteolysis of regulatory proteins, SsrA RNA is specifically degraded by a
conserved ribonuclease, RNase R, but stabilized by an RNA-binding protein, SmpB.
SsrA (also known as tmRNA and 10Sa RNA) is a small, highly structured RNA
that is found in all bacteria and in chloroplasts and mitochondria of some eukaryotes
(Felden et al., 1999; Gueneau de Novoa and Williams, 2004; Jacob et al., 2004; Keiler et
al., 2000). SsrA interacts with selected ribosomes to add a peptide tag to nascent
31
polypeptides that are subsequently recognized and degraded by intracellular proteases
(Gottesman et al., 1998; Karzai et al., 2000; Keiler et al., 2000). SsrA activity is proposed
to regulate gene expression and provide a quality-control mechanism for translation (Abo
et al., 2000; Hayes et al., 2002a, 2002b; Karzai et al., 2000; Keiler et al., 1996; Ranquet
et al., 2001; Roche and Sauer, 1999, 2001; Sunohara et al., 2002). The phenotypes of
ssrA deletion stains in several bacterial species suggest that SsrA functions are important
in bacterial physiology (Ebeling et al., 1991; Julio et al., 2000; Keiler and Shapiro,
2003a; Withey and Friedman, 1999).
In Caulobacter crescentus, SsrA is required for coordinated cell cycle progression
(Keiler and Shapiro, 2003a), and both the synthesis and degradation of SsrA are tightly
controlled during the cell cycle (Keiler and Shapiro, 2003b). The cell cycle of C.
crescentus is linked to developmental progression such that initiation of DNA replication
is coincident with differentiation from the swarmer cell stage (G1 phase) to the stalked
cell stage (S phase) (Hung et al., 1999). SsrA RNA is transcribed just before this G1-S
transition, specifically degraded during early S phase, and re-synthesized late in S phase
(Keiler and Shapiro, 2003b). Deletion of the ssrA gene results in a disruption of the cell
cycle at the G1-S transition, consistent with an important role for the timing of SsrA
RNA synthesis and degradation (Keiler and Shapiro, 2003a). The striking temporal
expression pattern of SsrA RNA during the cell cycle raises the question of what
regulatory factors control SsrA RNA expression and activity.
In C. crescentus, as in all other � -proteobacteria, SsrA RNA is composed of two
RNA molecules due to a circular permutation in the ssrA gene (Keiler et al., 2000). The
ssrA gene is transcribed as a single RNA, pre-SsrA, which is predicted to form a similar
32
secondary structure to the mature SsrA except that the tRNA-like 5’ and 3’ ends are
connected by a hairpin loop. This loop is then excised to produce mature SsrA, composed
of the coding RNA and the acceptor RNA (Figure 2.1A). In other bacteria such as E. coli,
SsrA is made as a single transcript and the ends are processed in the same manner as for
tRNAs. In all cases, processing of pre-SsrA to the active mature form is mechanistically
and temporally distinct from the regulatory removal of SsrA, just as post-translational
proteolytic processing is distinct from regulatory protein degradation.
Several proteins are required for SsrA activity, including the general translation
factors EF-Tu and alanine-tRNA synthetase, and the SsrA-binding protein SmpB. SmpB
binds with high affinity to the tRNA-like domain of SsrA RNA (Barends et al., 2001;
Karzai et al., 1999), and is required for interaction of SsrA RNA with the ribosome
(Karzai et al., 1999). In C. crescentus, SmpB is required for normal steady-state levels of
SsrA RNA, and a deletion of smpB has the same phenotype as a deletion of ssrA (Keiler
and Shapiro, 2003a). SmpB is widely distributed in bacteria, and SmpB homologues have
been identified in almost all species in which SsrA has been found.
RNase R co-purifies at sub-stoichiometric levels with the SsrA-SmpB complex
from E. coli but is not required for SsrA activity, and the significance of this interaction is
unknown (Karzai and Sauer, 2001). RNase R, encoded by the rnr or vacB (virulence-
associated) gene, was originally identified as a residual 3’ to 5’ exoribonuclease activity
in an E. coli strain devoid of RNase II (Kasai et al., 1977), and is a member of the RNR
superfamily of exoribonucleases (Zuo and Deutscher, 2001). RNase R is highly
processive and can degrade RNAs with significant secondary structure, such as rRNA
and mRNAs containing stable stem-loops (Cheng and Deutscher, 2005), although tRNAs
33
are poorly degraded. Despite biochemical association with the E. coli SsrA-SmpB
complex, RNase R has not been implicated in the degradation of SsrA RNA or damaged
mRNAs, and its role in the SsrA pathway has been unclear. The data presented here
demonstrate that SsrA RNA in C. crescentus is specifically degraded by RNase R at a
specific point in the cell cycle, and that this timing may be regulated by SmpB.
34
RESULTS
C. crescentus RNase R specifically degrades SsrA RNA in vitro
RNase R was identified as a candidate for the nuclease responsible for complete
degradation of SsrA RNA in stalked cells because RNase R co-purifies with SsrA RNA
in E. coli (Karzai and Sauer, 2001), and it is a highly processive 3’ to 5’ exonuclease that
is capable of degrading RNAs with significant secondary structure (Cheng and
Deutscher, 2005; Cheng and Deutscher, 2002, 2003). The C. crescentus homologue of
rnr was identified by sequence similarity to the E. coli gene. The predicted amino acid
sequences of the C. crescentus and E. coli RNase R are over 32% identical and 48%
similar, and both contain a conserved RNase II family signature in the central region and
a C-terminal S1 RNA-binding domain.
To determine if RNase R can directly degrade SsrA RNA, a histidine-tagged
variant of C. crescentus RNase R was purified and assayed in vitro. When incubated with
mature SsrA RNA, RNase R rapidly degraded both the coding and acceptor RNAs. More
than 70% of the mature SsrA RNA was completely degraded within 5 min, corresponding
to a minimum initial rate of 14 min-1 (Figure 2.1B panel 1 and C). The remaining RNA is
significantly more stable, suggesting that there may be two RNA populations in the
mature SsrA RNA preparation. In contrast to the mature SsrA RNA, when either the SsrA
acceptor RNA alone or the SsrA coding RNA alone was incubated with RNase R, neither
was degraded (Figure 2.1B panels 2 and 3). These data indicate that mature SsrA RNA,
but not its individual components, is recognized by RNase R as a substrate. The residual
RNA observed after 5 min incubation of mature SsrA RNA with RNase R may represent
a sub-population of SsrA acceptor RNA and SsrA coding RNA that is not correctly
35
folded, and therefore is not recognized by RNase R. Like mature SsrA RNA, pre-SsrA
RNA was a substrate for RNase R, but was degraded at a slightly slower rate (Figure
2.1B panel 4 and C). For comparison, the degradation of a 17-mer oligo(A) RNA, a good
substrate for E. coli RNase R (Cheng and Deutscher, 2002), and tRNA, a poor substrate
for E. coli RNase R (Cheng and Deutscher, 2002), were assayed under the same
conditions as SsrA RNA (Figure 2.1B panels 5 and 6, and C). Oligo(A) RNA was
degraded 3-fold slower than mature SsrA RNA, whereas very little activity against tRNA
was observed. To obtain more detailed kinetic parameters, the degradation of pre-SsrA
RNA and poly(A) RNA was measured using a TCA precipitation assay and the apparent
kinetic constants were determined (Figure 2.1C). Both substrates were efficiently
degraded by RNase R, with very similar kcat and KM values. These data indicate that pre-
SsrA RNA is as good a substrate for RNase R as poly(A) RNA, and mature SsrA RNA is
the best substrate yet identified.
The degradation of SsrA RNA appears to be processive, because no degradation
intermediates were observed for any substrate in the electrophoresis-based assay (Figure
2.1B). Of particular interest, no processing of pre-SsrA to the mature form could be
detected (Figure 2.1B panel 4). To ensure that the observed degradation of pre-SsrA is in
fact complete degradation and not processing, aliquots of radiolabeled pre-SsrA were
removed during incubations with RNase R and resolved on a polyacrylamide gel. Again,
no degradation intermediates were observed at any point in the time course (Figure 2.1B
panel 4), indicating that pre-SsrA was degraded into fragments <32 nucleotides in length.
These data are consistent with degradation of SsrA RNA and other substrates to
individual nucleotides, as would be expected for a highly processive exonuclease.
36
RNase R is required for cell-cycle dependent degradation of SsrA RNA
To determine if RNase R degrades SsrA RNA in vivo as well as in vitro, a
deletion of rnr was constructed in C. crescentus and the stability of SsrA RNA was
measured in the Δrnr strain. In log-phase cultures of the Δrnr strain, both coding and
acceptor SsrA RNAs have half-lives longer than 30 min (Figure 2.2A). This rate
represents a significant stabilization over the 4-5 min half-life observed for the mature
SsrA RNAs in wild-type cells (Keiler and Shapiro, 2003b). The decay of pre-SsrA RNA
is also stabilized in the Δrnr strain, with a half-life of 4.7 ± 0.1 min compared to 2.5 ± 0.2
min for wild type. The half-lives of two mRNAs, pilA and a variant of λ repressor, were
1-2 min in the Δrnr strain (data not shown), the same as in wild type. These data indicate
that deletion of rnr does not result in a general stabilization of all RNAs, but SsrA RNA
is specifically stabilized.
In principle, the extended half-life of SsrA RNA in log-phase populations of the
Δrnr strain could be due to loss of specific degradation of SsrA RNA in stalked cells, or
to partial stabilization of SsrA RNA throughout the cell cycle. To distinguish between
these possibilities, the RNA degradation assays were repeated in pure populations of
swarmer cells and stalked cells. In swarmer cells isolated from the Δrnr strain, the mature
SsrA RNAs have half-lives >30 min, just as in the wild-type strain (Figure 2.2B).
However, in stalked cells the half-lives are also >30 min, compared to 4-5 min for wild
type (Figure 2.2C). Thus, in the Δrnr strain the cell-type specific degradation of SsrA is
eliminated and SsrA RNA is stable in all cell types.
If RNase R is the sole nuclease responsible for the turnover of mature SsrA RNA
in vivo, then it must degrade SsrA RNA with a rate sufficient to account for the observed
37
turnover. During the C. crescentus stalked cell stage, SsrA RNA is almost completely
degraded within 30 min, corresponding to a 4-5 min half-life. Estimates for the
concentration of SsrA RNA in C. crescentus indicate that there are approximately 2000
molecules per cell (K. C. Keiler, unpublished observations). In this case, the degradation
of mature SsrA RNA occurs with a rate of approximately 67 molecules/min (= 2000
molecules/30 min.). Since the observed rate of degradation of SsrA RNA by RNase R in
vitro is 14 mol SsrA RNA/ min / mol RNase R, there must be at least 5 molecules of
RNase R per cell to account for the observed degradation of SsrA RNA. Western blots
calibrated with purified RNase R protein indicate that there are at least 1000 RNase R
molecules per cell (data not shown). Therefore, assuming the conditions in vitro
approximate the conditions in vivo, RNase R is kinetically competent to be the sole
ribonuclease degrading SsrA RNA. Taken together, these in vivo and in vitro results
suggest that RNase R is directly responsible for degrading SsrA RNA in a cell-type
specific manner.
Lack of RNase R alters cell-cycle expression of ssrA
To determine if loss of cell-type specific degradation leads to over-accumulation
of mature SsrA RNA, total RNA was isolated from log-phase cultures of wild type and
the Δrnr strain, and the amount of SsrA RNA was measured by Northern blotting. The
amount of mature SsrA RNA was the same in the Δrnr strain as wild type (Figure 2.3A).
Therefore, loss of SsrA RNA degradation does not alter the steady-state level of SsrA
RNA. Even if the total amount of SsrA RNA is the same in a mixed culture, it is possible
that the cell-cycle regulation of expression is altered. To investigate whether RNase R-
38
mediated degradation is necessary for control of the pattern of ssrA expression through
the cell cycle, the amount of SsrA RNA in synchronous cultures of the Δrnr strain was
assayed by Northern blotting. In the Δrnr strain the cell-cycle expression pattern of SsrA
RNA was disrupted (Figure 2.3B & C). There was <2-fold change in steady-state level
over the course of the cell cycle, compared to a 5-fold change for wild type (Figure 2.3C;
Keiler and Shapiro, 2003b), and there was no decline during the stalked cell stage (30
min to 60 min after synchronization). Therefore, RNase R is necessary for degradation of
SsrA RNA and this degradation is required for cell-cycle regulation of SsrA RNA levels.
Phenotype of the rnr deletion strain
To investigate whether lack of RNase R and the resulting constitutive expression
of SsrA are detrimental to the cell, the morphology, growth, and cell cycle progression of
the Δrnr strain were assayed. The Δrnr strain showed no changes in cellular morphology
when examined by light microscopy. In addition, the growth rate of the Δrnr strain during
log-phase in complex and defined media and the timing of cell cycle events such as
initiation of DNA replication, expression of the cell-cycle regulated proteins CtrA and
McpA, loss of motility, and cell division, were not significantly different from those of
wild type C. crescentus (Table 2.1). Therefore, there is no significant defect in the cell
cycle in the absence of RNase R. These data indicate that RNase R activity is not
essential under culture conditions, and constitutive expression of SsrA RNA does not
disrupt the cell cycle under the growth conditions assayed. Because a deletion of ssrA
causes a significant defect in the cell cycle (Keiler and Shapiro, 2003a) and SsrA RNA is
normally cell-cycle regulated (Keiler and Shapiro, 2003b), the lack of a phenotype when
39
SsrA RNA is constitutively expressed suggests that there may be redundant mechanisms
to control SsrA activity.
SmpB protects SsrA RNA from RNase R degradation
SsrA RNA is specifically degraded in the stalked cell. One possible mechanism
for stalked cell-specific degradation would be to limit expression of RNase R to this cell
type. However, Western blots showed that the level of RNase R does not fluctuate
through the cell cycle in a pattern that would explain cell-type specific degradation of
SsrA RNA (Figure 2.4). Instead, RNase R protein accumulates through the cell cycle.
These results exclude a mechanism of temporal separation of SsrA RNA and RNase R,
and indicate that RNase R degradation of SsrA RNA is regulated by additional factors in
vivo.
One candidate for a regulator of SsrA RNA degradation is the SsrA-binding
protein SmpB. The steady-state level of SsrA RNA is decreased by 90% in a ∆smpB
strain, consistent with an increase in SsrA RNA degradation in the absence of SmpB
(Figure 2.3A; Keiler and Shapiro, 2003a). To determine if SmpB can directly protect
SsrA RNA from degradation by RNase R, the interactions among these molecules were
assayed in vitro. First, direct association of SmpB with SsrA RNA was examined by
equilibrium filter-binding assays. Purified SmpB bound to SsrA RNA with a Kd = 1.8 ±
0.1 nM (Figure 2.5), an affinity similar to that reported for E. coli SmpB and SsrA RNA
(Jacob et al. 2005; Sundermeier, et al., 2005). Second, the effect of SmpB on degradation
of pre-SsrA RNA by RNase R was assayed in vitro. As shown in Table 2.2, incubation of
40
SsrA RNA with SmpB prior to the addition of RNase R protected SsrA RNA from
degradation. When SsrA RNA was pre-incubated with 20 µM SmpB, no SsrA RNA
degradation could be detected after 2 h, indicating at least a 14-fold decrease in the
degradation rate. The degradation of oligo(A) RNA was not affected by pre-incubation
with SmpB (Table 2.2), indicating that SmpB is not a general inhibitor of RNase R, but
specifically inhibits the degradation of SsrA RNA.
Western blotting of lysates from synchronous cultures showed that the amount of
SmpB protein fluctuates during the cell cycle (Figure 2.4), with a similar pattern to SsrA
RNA (Figure 2.3C). SmpB levels are low in early swarmer cells, but increase
dramatically during the G1-S transition, between 15 and 30 min after synchronization.
During this interval, SsrA RNA accumulates in the cell. SmpB is rapidly removed from
the cell after the initiation of DNA replication, between 30 and 45 min after
synchronization, when SsrA RNA is being degraded. SmpB protein levels increase again
after 60 min, when SsrA RNA levels also increase. Thus the pattern of SmpB protein
level throughout the cell cycle corresponds precisely to the stability of SsrA RNA. The
pattern of SmpB protein level is unchanged in the ∆rnr strain, indicating that regulation
of SmpB does not require RNase R activity (data not shown). Moreover, because SsrA
RNA levels are constitutive in the ∆rnr strain, regulation of SmpB abundance does not
depend on the expression pattern of SsrA RNA. Because SmpB binds with high affinity
to SsrA RNA and protects SsrA RNA from degradation by RNase R in vitro, and SmpB
has the same cell-cycle regulated expression pattern as SsrA RNA in vivo, it is likely that
SmpB is required for SsrA RNA stability as well as activity.
41
DISCUSSION
The data presented here demonstrate that RNase R is the nuclease responsible for
cell-type specific degradation of SsrA RNA. In the absence of RNase R there is no
degradation of SsrA RNA, and RNase R is kinetically competent to account for the
observed degradation of SsrA RNA. The cell-cycle regulation of this degradation in vivo
is likely to be controlled by SmpB binding to SsrA RNA. The model presented in Figure
2.6 is sufficient to explain the data reported here, as well as known phenotypes of
mutations in genes encoding SsrA, SmpB, and RNase R from other bacteria. In this
model, SmpB protects SsrA RNA from degradation by RNase R at a non-tRNA-like 3’-
end.
The substrate specificity of RNase R indicates that degradation of SsrA RNA
requires an element of the folded structure, and is likely to initiate at the 3’ end of the
SsrA coding RNA in vivo. RNase R rapidly degrades both mature SsrA RNA and pre-
SsrA RNA, but will not degrade the SsrA coding RNA or the SsrA acceptor RNA alone.
This selectivity indicates that RNase R requires some feature of the folded SsrA RNA
structure that is not present in the isolated coding and acceptor RNAs. One possibility is
that RNase R specifically binds to part of the SsrA RNA folded structure. Alternatively,
an important sequence determinant, such as the 3’ end, may be obscured when the
acceptor RNA and the coding RNA are not properly folded. In either case, degradation of
folded SsrA RNA is likely to initiate at the 3’ end of the SsrA coding RNA, because
RNase R degrades both mature SsrA RNA and pre-SsrA RNA substantially faster than
tRNA. The 3’ end of tRNA and the 3’ end of the acceptor RNA in mature SsrA have
similar sequence and structure, and are largely constrained by base-pairing interactions.
42
In contrast, pre-SsrA RNA and the coding RNA in mature SsrA have a non-tRNA-like 3’
end that may be accessible to RNase R. Therefore, the stability of mature SsrA RNA in
vivo could be regulated by protecting or exposing the 3’ end of the SsrA coding RNA.
SmpB could limit access of RNase R to the 3’ end of the SsrA coding RNA either
by directly binding this region or by promoting a three-dimensional RNA structure that
buries this 3’ end. Biochemical and structural studies on SsrA-SmpB complexes from
other species demonstrate that SmpB binds to the tRNA-like domain of SsrA RNA, but
do not rule out a second contact with another portion of SsrA RNA (Barends et al., 2001;
Gutmann et al., 2003; Valle et al., 2003). The cell-cycle regulation of SmpB protein
levels could then control the susceptibility of SsrA RNA to RNase R degradation. When
SmpB is present in swarmer cells and pre-divisional cells, SsrA RNA is protected from
RNase R and is stable, but after the G1-S transition SmpB levels decrease and SsrA RNA
is degraded by RNase R. If this model is correct, the key event to degrading SsrA RNA in
stalked cells is proteolysis of SmpB, and persistence of SmpB protein through the cell
cycle should produce constitutive expression of SsrA RNA. Future studies using
mutations that alter SmpB proteolysis will be required to confirm the role of SmpB in
SsrA RNA stability.
The cell-cycle regulation of SmpB abundance also explains why a deletion of rnr
that results in constitutive ssrA expression has no phenotype. SmpB is required for
association of SsrA RNA with the ribosome (Karzai et al., 1999), so SsrA activity will be
properly controlled, even if SsrA RNA is ectopically produced, as long as SmpB levels
are cell-cycle regulated. The dual regulation of SsrA RNA activity by SmpB binding and
degradation of SsrA RNA by RNase R provides a redundant regulatory network to
43
control SsrA activity through the cell cycle, analogous to the control of key protein cell-
cycle regulators such as CtrA (Ryan and Shapiro, 2003).
Why is SsrA RNA stable in E. coli even though RNase R co-purifies with SsrA
(Karzai and Sauer, 2001)? In the context of the model presented in Figure 2.6, the lack of
degradation of E. coli SsrA RNA by RNase R can be explained by the absence of a non-
tRNA-like 3’ end. If RNase R can not recognize the tRNA-like 3’ end of SsrA, then E.
coli SsrA RNA would have to unfold or be cleaved by another nuclease before RNase R
could degrade it. Significantly, this model is consistent with the observed specificity of
RNase R for its other known substrates in vivo, rRNAs and mRNAs with significant
secondary structure. RNase R degrades intact rRNAs with a rate 5-fold lower than
poly(A) RNA (Cheng and Deutscher, 2002). Likewise, mRNAs containing stable stem-
loop structures are degraded rapidly by RNase R after polyadenylation by poly(A)
polymerase (Cheng and Deutscher, 2005). Thus, E. coli SsrA RNA would only be a good
substrate for RNase R if it is damaged, but the two-piece construction of C. crescentus
SsrA RNA would provide access to a 3’ end without the involvement of another
nuclease. Interestingly, E. coli SsrA RNA is cleaved internally in a reaction that is
stimulated by toxins such as RelE (Christensen and Gerdes, 2003) and MazF
(Christensen et al., 2003). The proposed model predicts that cleaved SsrA RNA
generated under stress conditions would be rapidly degraded by RNase R. Furthermore,
the opportunity to regulate SsrA RNA by RNase R-mediated degradation may have
contributed to the selective advantage that has produced at least three lineages of
circularly permuted ssrA genes (Sharkady and Williams, 2004).
44
It has been suggested that RNase R is required for the processing of pre-SsrA in
E. coli under cold shock conditions (Cairrao et al., 2003). RNase R is not induced under
cold shock conditions in C. crescentus, and is not required for processing pre-SsrA RNA
under cold shock or heat shock conditions, during log-phase growth, or during stationary
phase (Figure 2.3A and data not shown). RNase R degrades pre-SsrA RNA in vitro, but
no intermediate products that would be consistent with processing to the mature form
were observed in vitro or in vivo. Instead, pre-SsrA RNA is completely degraded by
RNase R in a processive manner.
SsrA and RNase R are each required for virulence in pathogenic bacteria, and the
results presented here raise the possibility that these two molecules function together
during pathogenesis. SsrA is required for virulence in Salmonella typhimurium (Julio et
al., 2000), whereas RNase R is essential for virulence in Shigella flexneri and
enteroinvasive E. coli (Cheng et al., 1998). If RNase R is required because it must
degrade SsrA RNA in these species during pathogenesis, then both the presence and the
removal of SsrA would be critical for pathogenic processes. This dependence would put
SsrA at the center of an uncharacterized pathway for regulation of processes important
for pathogenesis, and suggests that the SsrA pathway might be a useful target for
antibacterial drugs. It will be crucial to examine the role and regulation of SsrA, SmpB,
and RNase R during pathogenesis to test these possibilities.
45
MATERIALS AND METHODS
Bacterial strains and plasmids
The wild-type C. crescentus strain used in this study is CB15N (Evinger and
Agabian, 1977). C. crescentus strains were grown at 30oC in M2G, M2X, or PYE
medium (Ely, 1991) supplemented with 1-2 µg/ml chloramphenicol, 5-20 µg/ml
kanamycin, or 25-50 µg/ml spectinomycin as necessary, and monitored by optical density
at 660 nm. The Δrnr strain was constructed by engineering an in-frame deletion of all but
six codons of the rnr open reading frame using the two-step recombination method as
previously described (Gay et al., 1985), and verified by PCR analysis and Southern
blotting. E. coli strains were grown at 37°C in Luria-Bertani broth (Sambrook et al.,
1989) supplemented with 50-100 µg/ml ampicillin, 20-30 µg/ml chloramphenicol, or 30-
50 µg/ml kanamycin as necessary, and monitored by optical density at 600 nm. pSmpBa
was constructed by amplifying the coding sequence of the smpB gene from C. crescentus
genomic DNA by PCR and cloning the product into the plasmid pET28a (Novagen) to
produce a gene encoding N-terminal His6-tagged SmpB. To generate pSmpBb, the smpB
gene was amplified by PCR using primers to add a His6 tag at the 5’ end of the smpB
coding sequence, and the product was cloned into plasmid pML81 under control of the
xylose-inducible promoter (Meisenzahl et al., 1997). To produce RNase R with a His6 tag
at the C terminus, plasmid pRNR was constructed by amplifying the coding region of rnr
from C. crescentus genomic DNA by PCR and cloning the product into pET-21a
(Novagen).
46
Cell-cycle experiments
Synchronized cultures of C. crescentus were obtained by isolation of swarmer
cells from Ludox density gradients (Evinger and Agabian, 1977). Aliquots of
synchronized cultures were removed every 15 min for analysis by flow cytometry,
Western blotting, or Northern blotting. The timing of loss of motility and cell division in
these cultures was estimated by visual inspection using light microscopy. The levels of
SmpB, RNase R, CtrA, and McpA were analyzed by Western blotting followed by
quantification using ImageQuant software (Molecular Dynamics). Flow cytometry assays
for DNA content and initiation of replication were performed as previously described
(Winzeler and Shapiro, 1995).
Northern blotting and RNA turnover
Total RNA was isolated using the hot phenol method (Sambrook et al., 1989).
Northern blotting was performed after separating equal quantities of total RNA on
polyacrylamide-urea gels. Specific RNAs were visualized by hybridization with [32P]-
labeled DNA probes generated from PCR products using the QuickPrime protocol
(Amersham Biosciences), and quantified using a PhosphorImager with ImageQuant
software. As a control for loading and transfer of RNA, the blots were re-probed with 5S
rRNA, which does not fluctuate through the cell cycle (Keiler and Shapiro, 2003b). RNA
decay experiments were performed by inhibiting transcription with rifampicin (15 µg/ml
final concentration) and assaying the fate of existing RNAs by Northern blotting as
previously described (Keiler and Shapiro, 2003b).
47
Protein purification and antisera production
His6-SmpB for binding assays was produced from log-phase cultures of E. coli
strain BL21(DE3)/pLys (Novagen) bearing plasmid pSmpBa by growth in the presence
of 1 mM isopropyl-β-D-thiogalactopyranoside for 3 h. The culture was quickly cooled on
ice, cells were harvested by centrifugation, and the cell pellet was suspended in 5 ml
buffer A (50 mM NaH2PO4 (pH 8.0), 300 mM NaCl, 5 mM β-mercaptoethanol, 10 mM
imidazole) per gram wet weight and lysed by sonication. The lysate was cleared by
centrifugation at 10,000 x g for 30 min and the supernatant was added to 0.5 ml Ni2+-
nitrilotriacetic acid (NTA) resin (Qiagen) equilibrated in buffer A. After mixing for 1 hr
at 4°C, the resin was packed into a column, washed with 100 ml buffer A, 100 ml buffer
A20 (A buffer with 20 mM imidazole), and eluted with 10 ml buffer B (50 mM MES (pH
6.5), 300 mM NaCl, 5 mM β-mercaptoethanol, 250 mM imidazole). The salt
concentration of eluted protein was diluted by the addition of 5 volumes of 50 mM MES
(pH 6.5) and applied to a MonoS HR 5/5 column (Amersham Biosciences) equilibrated in
buffer S100 (50 mM MES (pH 6.5), 0.1 M KCl), and developed with a 50 ml gradient
from 0.1–1 M KCl. Fractions containing the purified protein were identified by SDS-
polyacrylamide gel electrophoresis (PAGE), quickly frozen in liquid nitrogen, and stored
at –80°C.
His6-SmpB for antibody production was produced as described above with the
following exceptions. The cell pellet was suspended in buffer C (10 mM Tris-HCl (pH
8.0), 100 mM NaH2PO4, 8 M urea, 5 mM imidazole). The cleared supernatant was added
to 1 ml Ni2+-NTA resin equilibrated in buffer C. After mixing for 1 h at room
temperature, the resin was packed into a column, washed with 100 ml buffer C, 100 ml
48
buffer C20 (C buffer with 20 mM imidazole), and eluted with 10 ml buffer C containing
250 mM imidazole. Fractions were separated on a preparative SDS-polyacrylamide gel
and the band corresponding to the His6-SmpB protein was excised and used to immunize
rabbits. Immunization and sampling of the sera were performed by Josman LLC. (Napa,
CA). The antisera were affinity-purified using the original antigen coupled to AminoLink
Plus Coupling Gel (Pierce) and used for Western blotting.
RNase R-His6 for enzyme assays was produced and purified by Ni2+-NTA
chromatography as described for His6-SmpB from E. coli strain BL21(DE3)/pLys bearing
plasmid pRNR with the following exceptions. Buffer A for RNase R-His6 purification
was prepared without β-mercaptoethanol. The salt concentration of eluted protein from
Ni2+-NTA was diluted by the addition of 5 volumes of 10 mM Tris-HCl (pH 7.6) and
applied to a MonoQ HR 5/5 column equilibrated in buffer Q100 (10 mM Tris-HCl (pH
7.6), 0.1 M KCl, 1 mM DTT, 0.5 mM EDTA), and developed with a 50 ml gradient from
0.1–1 M KCl. Fractions containing the purified protein were identified by SDS-PAGE,
pooled, and dialyzed against buffer Q500 (10 mM Tris-HCl (pH 7.6), 0.5 M KCl, 1 mM
DTT, 0.5 mM EDTA, 10% glycerol). The dialyzed sample was quickly frozen in liquid
nitrogen and stored at -80oC. Antisera against RNase R-His6 were generated as described
above.
RNA purification and labeling
Mature SsrA RNA was purified by affinity to SmpB. Log-phase cultures of C.
crescentus bearing pSmpBb were induced by the addition of xylose to 0.03%, and grown
for 3 h at 30°C in PYE broth. Cells were harvested by centrifugation, resuspended in 5 ml
buffer D (50 mM NaH2PO4 (pH 8.0), 150 mM NaCl, 5 mM imidazole) per gram wet
49
weight and lysed in a French press. The lysate was cleared by centrifugation at 10,000 x g
for 30 min and added to 2 ml Ni2+-NTA resin equilibrated in buffer D. After mixing for 1
h at 4°C, the resin was packed into a column, washed with 200 ml buffer D, and bound
protein was eluted with 10 ml buffer D containing 250 mM imidazole. The mature SsrA
RNA was extracted from fractions containing SmpB with acid phenol (pH 4.5)-
chloroform, precipitated with ethanol (Sambrook et al., 1989), and desalted with mini
Quick Spin RNA columns (Roche Applied Science). To remove contaminating RNA, the
mature SsrA RNA was separated by polyacrylamide-urea gel electrophoresis and the
bands corresponding to mature SsrA RNA were excised and soaked in Tris-EDTA buffer
(pH 7.0) containing 250 mM NaCl. The supernatant was extracted with acid phenol (pH
4.5)-chloroform, and the RNA was precipitated with ethanol and resuspended in water.
Mature SsrA RNA was dephosphorylated with alkaline phosphatase, labeled at the 5'-
ends with [γ-32P]ATP and T4 polynucleotide kinase, and purified using mini Quick Spin
RNA columns.
C. crescentus pre-SsrA was transcribed from a PCR-generated DNA fragment
using the T7 RiboMax Large Scale RNA Production System (Promega) as described in
the manufacturer’s instructions, and the transcribed RNA was separated from
unincorporated nucleotides and DNA fragments using the RNeasy MinElute Cleanup Kit
(Qiagen). The presence of predicted secondary structures was confirmed by chemical
probing, and the presence of unprocessed, pre-SsrA RNA was confirmed by Northern
blotting. Pre-SsrA, poly(A) RNA (Sigma), and E. coli tRNA (Sigma) were labeled at the
5’ end as described for mature SsrA RNA. 3’-end labeled pre-SsrA was prepared by
ligation of [α-32P]pCp using T4 RNA ligase, and purified using RNeasy MinElute
50
Cleanup Kit. 17-mer oligo(A) RNA was synthesized (IDT) and labeled at the 5’ end with
[γ-32P]ATP and T4 polynucleotide kinase, and purified using mini Quick Spin Oligo
columns (Roche Applied Science).
RNA binding assays
100 pM 5’-end labeled pre-SsrA was incubated with varying amounts of purified
His6-SmpB protein in 20 µl reactions containing buffer E (50 mM MES (pH 6.5), 200
mM KCl, 5% glycerol, 5 mM β-mercaptoethanol, 0.01% NP-40, and 0.1 mg/ml BSA, 10
U of RNasin (Promega)) for 30 min at room temperature. The reaction products were
passed under vacuum through a 0.45 µm nitrocellulose membrane filter (Millipore) that
had been presoaked with buffer E. The filter was washed with 500 µl of buffer E under
vacuum and dried, and radioactivity was determined by scintillation counting.
Ribonuclease assays
RNase R degradation assays were performed at 37°C in 10 µl total volume
containing labeled substrate, buffer R (20 mM Tris-HCl (pH 8.2), 100 mM KCl, 0.5 mM
MgCl2), and 11-223 nM purified RNase R. At each time point a 1 µl aliquot of the
reaction mixture was added to 250 µl ice-cold 5% TCA with 25 µg salmon testes DNA,
incubated on ice for 15 min, and passed under vacuum through a glass microfiber filter
(VWR) that had been presoaked with ice-cold 5% TCA. The filter was washed twice with
10 ml of ice-cold 5% TCA under vacuum and dried, and acid-insoluble radioactivity was
determined by scintillation counting. The fraction RNA remaining was plotted versus
time, and the initial rate of degradation was obtained from a linear fit to the data.
51
Apparent steady-state kinetic parameters were obtained from non-linear curve fitting of
plots of initial rate versus RNA concentration using Prism software (GraphPad).
RNase R degradation assays using polyacrylamide-urea gel analysis were
performed at 37°C in 80 µl total volume containing 5-10 µM 5’-end labeled substrates,
buffer R, and 50-100 nM purified RNase R. At each time point a 9 µl aliquot of the
reaction mixture was added to 2 volumes RNA loading buffer (Sambrook et al., 1989) to
stop the reaction, analyzed on polyacrylamide-urea gels and quantified using a
PhosphorImager with ImageQuant software.
For assays in the presence of SmpB, 5 µM pre-SsrA labeled at the 5’ end or 10
µM oligo(A) RNA labeled at the 5’ end was incubated in the presence of 0-20 µM SmpB
for 30 min at room temperature in buffer R, and RNase R was added to 50-100 nM final
concentration. The reaction was incubated at room temperature for 2 hours, and
degradation was monitored as above, except that acid-soluble radioactivity was
determined by scintillation counting. Relative activity was calculated by subtracting
background counts observed in the absence of RNase R, and normalizing the rate to that
observed in the presence of RNase R with no SmpB.
52
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Figure 2.1. RNase R activity in vitro. (A) Cartoon depicting forms of SsrA RNA in C. crescentus. The ssrA gene is transcribed as a single RNA, pre-SsrA, in which the tRNA-like 5’ and 3’ ends are connected by a hairpin loop. Excision of the internal loop produces mature SsrA composed of a coding RNA and an acceptor RNA. (B) RNase R degradation of 5’ [32P]-labeled RNA substrates was analyzed on polyacrylamide-urea gels, and representative gels are shown. The mobilities of intact RNA molecules and the dye front are indicated, and the approximate RNA size expected to comigrate with the dye is noted in parentheses. (C) Rates and kinetic parameters for RNase R degradation of RNAs in vitro. Rate/[RNase R] was calculated based on the fraction of RNA that remained after 5 min incubation with RNase R in assays as shown in panel B. KM and kcat were determined from TCA precipitation assay data by non-linear curve fitting of plots of initial rate versus substrate concentration.
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Figure 2.2. Degradation of SsrA RNA in wild-type and ∆rnr strains. (A) The decay of SsrA coding RNA (squares) and SsrA acceptor RNA (circles) in log-phase cultures of Δrnr (filled blue symbols) was assayed by inhibiting transcription and measuring the loss of SsrA RNAs by Northern blotting. Half-lives of SsrA RNAs were obtained by fitting the data to single exponential functions. Assays were repeated in pure populations of swarmer cells (B) and stalked cells (C). Published data (Keiler and Shapiro, 2003b) from wild type (open symbols) are shown for comparison.
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Figure 2.3. Expression of SsrA RNA in the Δ rnr strain. (A) Equal amounts of total RNA isolated from the wild-type (wt), ΔssrA, ΔsmpB, and Δrnr strains were analyzed by Northern blots probed for SsrA RNA. A representative Northern blot with bands corresponding to pre-SsrA RNA, SsrA coding RNA, and SsrA acceptor RNA is shown. Results for wt, ΔssrA, and ΔsmpB are from published data (Keiler and Shapiro, 2003a). (B) A representative Northern blot of total RNA isolated from synchronized cultures of the Δrnr strain probed for SsrA RNA. The stages of the cell cycle are indicated schematically (top), and arrows indicate the bands corresponding to the SsrA coding RNA and the SsrA acceptor RNA. (C) The amounts of SsrA coding RNA (squares) and SsrA acceptor RNA (circles) from 3 independent experiments as in panel B were quantified and normalized to the peak level of SsrA RNA. Published data (Keiler and Shapiro, 2003b) from wild type (open symbols) are shown for comparison.
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Figure 2.4. Cell-cycle regulation of RNase R and SmpB protein levels. Steady-state protein levels were determined in synchronized cultures by Western blots probed with antibodies specific to RNase R or SmpB. A representative Western blot for each protein is shown. The stages of the cell cycle are indicated schematically (top). Western blots were quantified, normalized to the 105 min time point, and the average of at least 3 experiments was plotted versus time after synchronization.
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Figure 2.5. Binding kinetics of purified SmpB to SsrA RNA. A curve for pre-SsrA binding by purified His6-SmpB. A Kd of 1.8 ± 0.1 nM was obtained by fitting data from three independent experiments to the following equation: 1/1+( Kd/[SmpB]). Error bars indicate the standard deviation.
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Figure 2.6. Model for regulation of SsrA RNA by RNase R and SmpB. (A) C. crescentus SsrA RNA is abundant in swarmer cells because RNase R cannot recognize either the tRNA-like 3’- end of the SsrA acceptor RNA or the 3’-end of the SsrA coding RNA. The 3’-end of the coding RNA may be protected by SmpB. In stalked cells, SmpB is absent and the 3’-end of SsrA coding RNA is exposed, resulting in recognition and degradation by RNase R. (B) E. coli SsrA RNA contains only a tRNA-like 3’-end that is not accessible to RNase R. If SsrA RNA is damaged or cleaved under stress conditions, the new 3’-end may be recognized by RNase R. Unfolding of the tRNA-like acceptor arm may lead to degradation by RNase R in either species.
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Table 2.1. Growth parameters of wild type and ∆rnr strains.
Wild type Δrnr Doubling time in PYE medium1 102 ± 4 110 ± 5 Doubling time in M2G medium1 153 ± 11 173 ± 11
Initiation of DNA replication2 22 ± 3 28 ± 2 Length of S-phase3 90 90
Loss of motility4 15-30 15-30 Cell division4 135-150 135-150
CtrA degradation/synthesis5 16 / 76 22 / 83 McpA degradation/synthesis5 40 / 91 42 / 98
1. The doubling time (min) during exponential growth with the standard deviation. 2. Time (min) at which 50% of cells have initiated DNA replication as assayed by flow cytometry after treatment of cultures with rifampicin. 3. Time (min) required after initiation for the average DNA content to reach the level corresponding to two chromosomes. 4. Time (min) of cell division and loss of motility are estimated from light microscopy study of synchronized cultures as interval during which more than 80% of the cells lost motility or divided. 5.Time (min) of protein degradation/resynthesis monitored by Western blotting.
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Table 2.2. SmpB specifically inhibits RNase R degradation of SsrA RNA.
Substrate pre-SsrA RNA oligo(A) RNA RNase R – + – + + + – + +
SmpB (µM) 0 0 2.5 2.5 10.0 20.0 0 0 10 % activity1
0 100 0 31 10 0
0 100 100 1. The counts released from 5’ [32P]-labeled substrates after 2 h incubation with RNase R expressed as a percentage of the value in the absence of SmpB for each substrate.
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Chapter 3
Proteomic studies of physiological substrates for the SsrA
system in Caulobacter crescentus
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ABSTRACT
SsrA is a small RNA that regulates translation in all bacteria by mediating the
addition of a peptide tag to proteins before they are released from the ribosome. To date,
however, only 12 cellular substrates of SsrA have been identified. Using a variant of
SsrA encoding a protease-resistant His6 tag, substrates of Caulobacter crescentus SsrA
were purified and identified by mass spectrometry. The more than 200 SsrA substrates
belong to multiple functional categories: 11 substrates are enzymes involved in DNA
replication, recombination, and repair, consistent with the known role for SsrA in
initiation of DNA replication; 18 substrates are enzyme components of amino acid
biosynthesis and degradation, suggesting a role for SsrA as a regulator of amino acid
metabolism; 36 substrates are involved in protein synthesis or degradation, indicating that
SsrA affects total protein production and stability in the cell; 29 substrates are membrane
proteins involved in transport, including 10 TonB-dependent receptors, suggesting a role
for SsrA in regulating membrane protein production or membrane transport; 23 substrates
are involved in energy metabolism, indicating that SsrA may play a role in tuning the
metabolic potential of the cell. Analysis of the protein sequence, mRNA structure, and
DNA sequence adjacent to the tagging site revealed a 15-base DNA motif downstream of
the substrate tagging sites. These results indicate that SsrA plays a central regulatory role
in many different aspects of the cell physiology and that there is a specific rule governing
substrate selection by SsrA.
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INTRODUCTION
A number of small regulatory RNAs have recently been identified in bacteria but
the functions of many of the RNAs are not known (Gottesman, 2004). Among these small
regulatory RNAs, SsrA is a specialized RNA with both canonical tRNA-like and mRNA-
like properties (Keiler et al., 1996; Komine et al., 1994; Tu et al., 1995; Ushida et al.,
1994). Using these dual activities, SsrA, together with the protein SmpB and other
translational cofactors, functions both to release stalled ribosomes and to target complete
or truncated proteins translated by the stalled ribosomes for rapid proteolytic degradation
(Karzai et al., 2000; Keiler et al., 1996). In a process referred to as trans-translation,
SsrA charged with alanine enters the A site of a stalled ribosome and accepts the growing
polypeptide by acting as an alanyl-tRNA. SsrA then serves as an mRNA providing a
complete short open reading frame to add a proteolytic degradation tag to the nascent
polypeptide. Upon translation termination, the tagged protein is released and degraded by
cellular proteases (Keiler et al., 1996).
What does SsrA do for cellular physiology? It is well conserved throughout
bacteria, suggesting that SsrA provides intrinsic advantages for survival and that its
function is conserved. In fact, ssrA is essential for growth in Neisseria gonorrhoeae
(Huang et al., 2000). In other bacteria such as Salmonella typhimurium, ssrA is required
for virulence (Baumler et al., 1994; Julio et al., 2000); in Caulobacter crescentus, ssrA is
required for cell cycle progression and plasmid replication (Keiler and Shapiro, 2003a;
Chapter 4). ssrA-deficient Escherichia coli exhibit various subtle phenotypes including
slower growth at high temperature (Komine et al., 1994; Oh and Apirion, 1991); reduced
motility (Komine et al., 1994); induction of Alp protease activity (Kirby et al., 1994), and
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inhibition of variant forms of phage λ and µ development (Karzai et al., 1999; Retallack
et al., 1994; Withey and Friedman, 1999). In Bacillus subtilis, ssrA is required for
optimal growth under stress conditions such as high temperature and high concentrations
of ethanol or cadmium (Muto et al., 2000). These phenotypes suggest SsrA plays a
critical regulatory role in bacterial physiology beyond its function for translational quality
control, but the precise details of its mechanism of action are not yet fully understood.
Despite the diverse physiological roles of SsrA, only twelve endogenous
substrates have been identified from B. subtilis and E. coli (Table 3.1; Abo et al., 2000;
Fujihara et al., 2002; Roche and Sauer, 2001). Analysis of four E. coli substrates revealed
three rules where endogenous mRNAs become targets for SsrA and protein tagging
(Table 3.1; Abo et al., 2000; Hayes et al., 2002a; Hayes et al., 2002b; Roche and Sauer,
2001). SsrA tagging occurs when ribosomes stall at the 3’ end of nonstop mRNAs
generated by premature termination of transcription (Abo et al., 2000), at clusters of rare
codons in an mRNA when the cognate tRNA is scarce (Hayes et al., 2002b; Roche and
Sauer, 1999), or at less efficient stop codons (Hayes et al., 2002b; Roche and Sauer,
2001). Also, the C-terminal amino acid sequence of some nascent peptides has been
found to be a major determinant for SsrA tagging (Hayes et al., 2002a; Roche and Sauer,
2001; Sunohara et al., 2002).
Although analyses of these substrates have definitely shed some light on how the
SsrA tagging system is activated, the activities of known tagged proteins do not explain
various cellular phenotypes mentioned above. Therefore, identification of more
endogenous substrates for SsrA and rules for substrate selectivity are necessary to gain
further insight into the molecular mechanisms underlying various phenotypes observed in
71
the absence of ssrA. Here, we report the identification of more than 200 C. crescentus
proteins that are tagged by a variant SsrA, SsrA-His6. Characterization of these tagged
proteins provides a more comprehensive understanding of the cellular roles of SsrA.
Analysis of DNA sequences around the tagged sites reveals at least one consensus motif
that may activate the SsrA system.
72
RESULTS
Construction of functional SsrA-His6
Endogenous proteins tagged by wild type C. crescentus SsrA with the peptide
AANDNFAEEFAVAA are difficult to detect because the tagged proteins are rapidly
degraded (Keiler et al., 2000). However, it has been demonstrated that proteins tagged by
a variant SsrA containing a protease-resistant sequence at their C terminus
(AANDNFAEEFAVDD) are stable (Keiler et al., 2000; Keiler and Shapiro, 2003). To
identify substrates for SsrA, we constructed another variant of SsrA, SsrA-His6. In this
SsrA-His6, the reading frame of the SsrA degradation tag is modified by PCR-directed
mutagenesis to encode six histidines as its C-terminal residues (Figure 3.1). This SsrA-
His6 adds the tag peptide AANDNFAEHHHHHH to in vivo targets and the resulting
tagged proteins are resistant to proteolytic degradation. The His6 epitope on proteins
tagged by SsrA-His6 also allows affinity purification of these substrates. A similar E. coli
SsrA construct has been used to identify SsrA-tagged proteins (Roche and Sauer, 2001).
To confirm that SsrA-His6 is active, we examined if phenotypes observed in the
absence of ssrA could be complemented by ssrA-His6. SsrA is required for normal cell
cycle progression (Keiler and Shapiro, 2003) and plasmid replication in C. crescentus
(Chapter 4). A multicopy plasmid expressing SsrA-His6 with the wild-type SsrA
promoter can be replicated and maintained in an ssrA-deletion strain as efficiently as a
comparable plasmid expressing wild-type SsrA. However, expression of SsrA-His6 does
not complement the cell cycle delay of ssrA-deficient cells, indicating that degradation of
one or more SsrA-tagged substrates is important for control of the cell cycle by SsrA.
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These data suggests that SsrA-His6 is functional in regard to releasing stalled ribosomes
and tagging the nascent proteins.
Proteomes of SsrA tagging in C. crescentus
To examine the complexity of SsrA-tagged proteins, C. crescentus with SsrA-His6
was cultured in rich medium and harvested during logarithmic growth. The cell lysates
were applied to a Ni2+-NTA column. The eluted proteins were separated by SDS-
polyacrylamide gel electrophoresis (PAGE) and visualized by Coomassie blue staining
(Figure 3.2). A control gel with samples from a mock purification using cells with no
plasmid showed a few bands including CobW, a histidine-rich protein that binds to Ni2+-
NTA resin nonspecifically (Nierman et al., 2001). In contrast, far more bands distributed
over a wide range of molecular weights were observed in the SDS-PAGE from the strain
containing SsrA-His6. Furthermore, the presence of stained bands with different
intensities suggests that some proteins are tagged by SsrA at a high rate. These results
indicate that a complex mixture of endogenous proteins is tagged by SsrA, even under
normal culture conditions.
Identification of tagged proteins
Sixteen of the most prominent SsrA-His6-tagged protein bands from the SDS-
PAGE were excised and digested with trypsin. The masses of the peptides after the
proteolytic digest were determined by matrix-assisted laser desorption ionization time-of-
flight (MALDI-TOF) mass spectrometry. The resulting sets of peptide masses from each
band were compared to the predicted tryptic fragments from the C. crescentus proteome
74
using the search program MS-Fit (Clauser et al., 1999). The criteria for positive
identification was recovery of at least four tryptic peptides including a peptide that
contains a part of the substrate fused to the SsrA-His6 peptide. These data analyses are
different from a standard proteomic analysis in several aspects. First, each excised band
from the one-dimensional SDS-PAGE may contain more than one protein. This results in
assigning multiple protein identities to each band. Second, SsrA tagging of proteins at
internal positions will lead to truncated proteins, and thus fewer peptides will be available
to match the full-length protein sequence. Third, SsrA tagging could occur at different
sites within a protein resulting in a heterogeneous set of junction peptides containing both
the protein and the SsrA-His6 tag sequences.
A total of 219 possible SsrA-tagged proteins (Table 3.2) were identified,
corresponding to about 6% of the estimated total number of C. crescentus proteins
(Nierman et al., 2001). A two-dimensional gel analysis revealed that 979 proteins of C.
crescentus proteome are expressed during exponential growth on minimal salts medium
with glucose as the carbon source (Grunenfelder et al., 2001). These proteins, however,
represent only highly abundant soluble cytoplasmic proteins in C. crescentus proteome
(Grunenfelder et al., 2001). Taking into account that 29 of identified SsrA substrates are
membrane-associated proteins, the remaining 190 substrates represent no more than 19%
of expressed cytoplasmic proteins encoded by the C. crescentus genome. These
substrates belonged to a variety of functional groups (Table 3.3): DNA metabolism;
protein synthesis, folding, and degradation; amino acid biosynthesis and degradation; cell
envelope and transport; metabolism; cellular processes such as chemotaxis and cell
division; transcription and regulation. No function could be assigned to 24.2% of the
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identified proteins. This is not surprising, as the C. crescentus genome was found to
contain 721 proteins with unknown function (19.2%) and 1012 hypothetical proteins
(26.9%), accounting for nearly 45% of the C. crescentus genome (Nierman et al., 2001).
This large subgroup of SsrA-tagged proteins with unknown function may represent novel
cellular pathways regulated by SsrA.
Functional diversity of SsrA-tagged proteins
Twelve proteins that function in DNA replication were identified as substrates for
SsrA, including a subunit of DNA polymerase III, a DNA primase, and a DNA gyrase. A
plasmid replication factor, the Rep protein of plasmid pJS14 (a pBBR1-based plasmid
used to deliver the ssrA-His6) was also identified as a substrate. These proteins account
for almost 20% of total proteins involved in DNA replication, recombination, and repair
in C. crescentus. SsrA has been implicated in the regulation of DNA replication due to
the fact that the strongest phenotype of the ssrA deletion strain is a delay in initiation of
DNA replication (Keiler and Shapiro, 2003). The identification of SsrA-tagged
replication proteins confirms that SsrA plays an active regulatory role in DNA
replication.
Thirty-six substrates are involved in protein synthesis or degradation. These
include components of several amino acid biosynthesis pathways, ribosomal proteins, and
general translation factors such as aminoacyl-tRNA synthetases and elongation factor Tu
(EF-Tu). A large group of proteolytic enzymes are also part of this functional category.
These substrates indicate that SsrA may control components of the translation machinery
and proteases, thereby affecting total protein production and stability in the cell. In
76
addition, the presence of protein cofactors associated with the SsrA system, alanyl-tRNA
synthetase, EF-Tu, ribosomal protein S1, and ClpP, suggests that SsrA may also
autoregulate its own activity.
A surprisingly large number of substrates are associated with cell membranes.
Ten TonB-dependent receptors, six ABC transporter components, and other transporters
are tagged by SsrA. C. crescentus is remarkably rich in TonB-dependent receptors, with
at least sixty-five gene products containing the TonB box (Nierman et al., 2001). These
receptors facilitate the transport of nutrients and macromolecules such as iron and
vitamin B12 into the periplasm. These macromolecular complexes are then transported
into the cell by ABC transporters (Ferguson and Deisenhofer, 2002). The identification of
TonB-dependent receptors and ABC transporter components may suggest that SsrA is
involved in some aspect of regulating membrane protein production or metabolite
transport. Of note, the six identified ABC transporter components are all ATP binding
cassettes, suggesting that SsrA may regulate membrane protein complexes by tagging a
specific component of the complexes.
Thirty-nine substrates are involved in metabolic functions, expanding a potential
role of SsrA in controlling cell growth. C. crescentus grows slow in the absence of ssrA
(Keiler and Shapiro, 2003), suggesting that SsrA may in fact regulate one or more
substrates of this group. Although a majority of these proteins are involved in energy
metabolism, additional proteins include enzymes that participate in fatty acid and lipid
metabolism, and cofactor biosynthesis. Eighteen known and putative transcription factors
or proteins involved in signal transduction pathways were tagged by SsrA. Such
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regulators included subunits of DNA-dependent RNA polymerase, σ70, hybrid histidine
kinase/response regulators, and a few potential transcriptional regulators.
Temporally regulated genes such as ctrA were maximally expressed at specific
times during C. crescentus cell cycle (Laub et al., 2000). A subset of proteins encoded by
cell cycle regulated genes are tagged by SsrA. However, these genes were distributed
throughout the entire cell cycle, with no evident correlation between time of expression
and SsrA-tagging.
mRNA levels of SsrA-tagged genes
To determine if SsrA regulates transcription of genes encoding the identified
substrates, changes in global transcription patterns were assayed in an ssrA deletion
strain. Using oligo microarrays with probes for 3767 predicted C. crescentus genes
(Nierman et al., 2001), we found that deletion of ssrA altered expression levels of about
200 genes (L. Ling and K. C. Keiler, unpublished data). The level of transcripts for
fifteen identified substrates increased in ssrA-deleted cells, indicating that they are
negatively controlled by SsrA (Table 3.2). In contrast, the mRNA levels of three
substrates decreased in ∆ssrA cells, suggesting that their expression is activated by SsrA
(Table 3.2). These results indicate that the loss of SsrA tagging due to the absence of ssrA
does not generally alter the mRNA levels of substrates, suggesting that most substrates
are regulated by SsrA post-transcriptionally.
78
SsrA tagging sites and substrate selectivity
The junction peptides containing both the substrate and the SsrA-His6 tag
sequences were identified for 201 proteins. Among these 201 proteins, 53 proteins had
two or more different junction peptides, indicating that they are tagged by SsrA at two or
more positions within the proteins. The SsrA-tagging sites within 18 identified substrates
could not be assigned due to the lack of a junction peptide. Some substrates cannot be
assigned a junction peptide because tagging occurs after a lysine or arginine residue. In
addition, the junction peptide could not be detected if the size of the tryptic peptide is
beyond the limit of detection by MALDI-TOF. In the future, these problems can be
circumvented by using a protease with different specificity from trypsin.
Identification of rules for SsrA substrate selectivity is important to understand the
molecular mechanism of SsrA in physiological processes. None of the previously defined
rules derived from studies on E. coli substrates explains more than 99.5% of C.
crescentus substrates. This result suggests that there are new sequence features that may
cause ribosome stalling leading to SsrA activation.
In principle, the determinants of SsrA tagging could be encoded in the DNA
sequence, RNA sequence or structure, or protein sequence. Therefore, regions upstream
and downstream of tagging sites of identified substrates were scanned with MEME
(Bailley and Elkan, 1994) to identify conserved sequences potentially representing
substrate selectivity rules. No common features were found in the protein sequence, but a
conserved 15 base pair (bp) motif (CGTCGCCCTGATCGA) was identified downstream
of the substrate tagging sites (Table 3.4). This motif could be found on either strand of
DNA adjacent to the tagging sites, suggesting that it may be an important DNA rather
79
than RNA determinant for SsrA tagging. The motif appeared 272 times out of 275
tagging sites.
80
DISCUSSION
In this study, we have used a variant SsrA, SsrA-His6 to identify SsrA substrates
in C. crescentus. The results demonstrate that the SsrA-tagging event occurs on a variety
of cellular mRNAs, resulting in a large number of tagged proteins in normally growing C.
crescentus.
In E. coli, a few determinants of SsrA tagging are documented based on
mutagenesis studies of four individual in vivo substrates (Table 3.1). Ribosome stalling
and SsrA-tagging occur during translation of truncated mRNAs lacking a stop codon
because translation release factors are not recruited (Abo et al., 2000). Such truncated
messages could result from premature transcription termination (Abo et al., 2000). A
combination of an inefficient opal stop codon (UGA) and cognate tRNA scarcity for rare
codons (Hayes et al., 2002b; Roche and Sauer, 2001) or the presence of a specific amino
acid (proline) just before the stop codon (Hayes et al., 2002a) generates ribosome
pausing, which can also lead to SsrA tagging.
Analysis of C. crescentus substrates shows that only one protein is tagged at a
rare codon, and that the rest of substrates do not follow the aforementioned rules.
Peptidyl-tRNA hydrolase (CC0484) is tagged immediately before a rare ATA codon,
which is present 782 times out of 1.2 × 106 codons in C. crescentus genome. Although
there is no information on tRNA abundance in C. crescentus, ribosome stalling at the rare
codon due to insufficient amount of cognate tRNA may lead to recruitment of SsrA and
subsequent tagging, as observed in E. coli. Twelve substrates are tagged at or within five
amino acid from their C terminus and do not appear to be associated with rare codons.
Furthermore, there are no examples of substrates ending in proline. Ribosomes arrested at
81
or near the stop codon because of an inefficient terminator may result in SsrA tagging of
these substrates. The remaining 188 substrates are tagged at internal positions.
These data indicate that rules for substrate selectivity in C. crescentus may be
different from those in E. coli. In fact, all four substrates characterized in E. coli are
associated with a different mechanism of ribosome stalling, suggesting that SsrA can be
recruited by a variety of features present in DNA, RNA, or protein levels. It is important
to note that only four substrates have been identified in E. coli (Abo et al., 2000; Roche
and Sauer, 2001), so it may be premature to conclude that there are different rules
governing substrate choice in C. crescentus and E. coli.
The presence of the 15-bp DNA motif downstream of 99% of SsrA tagging sites
suggests a simple model for mechanism for SsrA activity. The conserved motif may
function as a binding site for an unknown transcription factor that obstructs transcribing
RNA polymerase. The resulting truncated mRNAs without an in-frame stop codon will
cause ribosome stalling and recruit SsrA. A similar mode of SsrA activation was
observed in E. coli. Lac Repressor is tagged by SsrA because Lac Repressor binding to
lac operators within its gene could block completion of lacI transcription, resulting in
truncated lacI transcripts (Abo et al., 2000). Alternatively, the DNA motif may encode an
RNA element which is cut by a ribonuclease. Ribonuclease cleavage would produce
truncated mRNAs lacking an in-frame stop codon to terminate translation. Therefore,
ribosome stalling and SsrA tagging will occur during translation of such truncated
mRNAs. Though intriguing, the significance of the 15-bp DNA motif with respect to
SsrA substrate selectivity rules remains to be investigated. However, once this motif is
shown to be functionally important by mutational analysis, the SsrA-tagging signal will
82
provide a useful foundation for the bioinformatic identification of other likely SsrA
substrates.
The fact that SsrA tags a number of proteins involved in the regulation of multiple
biological processes makes SsrA a central regulator of cell physiology. Mutational
analysis of substrates identified in this study may lead to better insight into the
mechanism of substrate selection for SsrA.
83
MATERIALS AND METHODS
Bacterial strains and plasmids
The wild-type C. crescentus strain used in this study is CB15N (Evinger and
Agabian, 1977). C. crescentus strains were grown at 30oC in PYE medium (Ely, 1991)
supplemented with 1-2 µg/ml chloramphenicol as necessary, and monitored by optical
density at 660 nm. E. coli strains were grown at 37°C in Luria-Bertani broth (Sambrook
et al., 1989) supplemented with 20-30 µg/ml chloramphenicol as necessary, and
monitored by optical density at 600 nm. pSsrA-His6 was derived from pSsrA (Keiler and
Shapiro, 2003) by PCR-based mutagenesis to alter the last six codons of the open reading
frame to those encoding six histidines (Figure 3.1).
Purification of SsrA-tagged C. crescentus proteins
Strains CB15N and CB15N bearing pSsrA-His6 were grown in 6 L of PYE broth
supplemented with chloramphenicol at 30°C to optical density at 660 nm of 0.3-0.4. Cells
were harvested by centrifugation and resuspended in 5 ml L buffer (8 M urea, 100 mM
NaH2PO4, 10 mM Tris, 150 mM NaCl, 1 mM PMSF, 20 mM imidazole, pH 8.0) per
gram wet weight and lysed by stirring for 1 h followed by sonication. The lysate was
centrifuged for 30 min at 10,000 × g, and the supernatant was added to 5 ml Ni2+-
nitrilotriacetic acid (NTA) resin (Qiagen) equilibrated in L buffer. After mixing for 1 h,
the resin was packed into a column, washed sequentially with 200 ml LX buffer (L buffer
with X = 8, 6, 4, 2, and 0 M urea), and eluted with 10 ml L0 buffer with 500 mM
84
imidazole. Fractions containing tagged proteins were identified by SDS-polyacrylamide
gel electrophoresis, quickly frozen in liquid nitrogen, and stored at –80°C.
Identification of SsrA-tagged proteins by mass spectrometry
Purified tagged proteins were separated on a 4-15% gradient polyacrylamide gel
(Bio-Rad) and identified by Coomassie Brilliant Blue staining. Gel slices (approximately
0.5–1.0 cm) were excised and processed for peptide mass fingerprinting as described
(Shevchenko et al., 1996) with approximately 10 ng modified trypsin (Promega)/µl gel
slices. Tryptic peptides were eluted from the gel by multiple extractions with 5% formic
acid and 50% acetonitrile, and dried under vacuum to reduce the sample volume to 10-20
µl. Samples were analyzed by matrix-assisted laser desorption ionization time-of-flight
(MALDI-TOF) mass spectrometry using a Perspective Voyager-DE RP Biospectrometry
instrument (Stanford Protein and Nucleic Acid Facility). Initial peptide fingerprint
searches were performed using ProFound (prowl.rockefeller.edu/PROWL/prowl.html),
searching the NCBInr database of “other proteobacteria” sequences of all molecular
weights. To identify junction peptides containing the SsrA-His6 tag, a database containing
all translated putative open reading frames of the C. crescentus genome with the SsrA-
His6 tag (AANDNFAEHHHHHH) after each amino acid was created and searched using
MS-Fit (Clauser et al., 1999). An expected mass accuracy of 100 ppm was used. A
maximum of two missed enzymatic cleavage, and modification of cysteines by
carboxyamidomethylation were considered during the searches. For protein
identification, the requirement of successful matches entailed the junction peptide and a
minimum of four peptides.
85
Gene annotations and categorization
Most C. crescentus open reading frame (ORF) annotations were taken from
GenBank accession number (Nierman et al., 2001). For genes annotated in GenBank as
hypothetical or conserved hypothetical proteins, clusters of orthologous gene (COG)
descriptions were used if they were more detailed. Gene names shown in the tables came
from TIGR and generally refer to the homologous Escherichia coli gene (www.tigr.org).
Motif identification
The Multiple Em for Motif Elicitation (MEME) software (Bailley and Elkan,
1994) was used to identify matrix models describing amino acid or DNA sequence motifs
present upstream or downstream of gene products tagged by SsrA. The 11 (or 5) amino
acids upstream (downstream) or the 200 bases upstream and down stream of the tagged
amino acid of each protein were searched. Both strands of each sequence were searched
for DNA sequence motifs.
86
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Karzai, A. W., Roche, E. D., and Sauer, R. T. (2000). The SsrA-SmpB system for protein
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Roche, E. D., and Sauer, R. T. (1999). SsrA-mediated peptide tagging caused by rare
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Tu, G. F., Reid, G. E., Zhang, J. G., Moritz, R. L., and Simpson, R. J. (1995). C-terminal
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Figure 3.1. Schematic representation of wild type SsrA and SsrA-His6. Nucleotide and amino acid sequences of wild type (wt) SsrA and SsrA-His6 in the region encoding the peptide tag are shown. The nucleotides mutated in SsrA-His6 are underlined.
93
Figure 3.2. Detection of SsrA-His6-tagged proteins. SsrA-His6-tagged proteins were purified from a strain without SsrA-His6 (lane 2) or from a strain harboring SsrA-His6 (lane 3) through Ni2+-NTA chromatography and analyzed by SDS-PAGE. The gel was stained with Coomassie blue to visualize protein bands. The histidine-rich protein (CobW) interacting nonspecifically with Ni2+-NTA resin is indicated by an arrow.
94
Table 3.1. Identified SsrA-tagged proteins and tagging determinants.
Species or proteins Tagging determinant Reference E. coli Roche and Sauer, 2001 Lac Repressor mRNA without a stop codon Abo et al, 2000 RbsK mRNA with an inefficient opal stop
codon and rare Arg codons Hayes et al., 2002b
YbeL mRNA with a C-terminal Pro Hayes et al., 2002a GalE ND1 Roche and Sauer, 2001 B. subtilis Fujihara et al., 2002 YqaP ND1 YtoQ ND1 TreP ND1 YloN ND1 PerR ND1 TufA ND1 FolA ND1 GsiB ND1 1. Not determined.
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Table 3.2. List of SsrA-tagged proteins. Gene # tagging # Peptides % Coverage Gene product sites
DNA replication, recombination, and repair CC0346 priA 3 4/4/4 16/15/19 primosomal protein N CC1522 ligA 1 4 32 DNA ligase, NAD-dependent CC18441 mfd 2 5/5 10/11 transcription-repair coupling factor CC1926 dnaE 1 5 16 DNA polymerase III, alpha subunit CC2246 xseA 1 4 37 exodeoxyribonuclease large subunit CC2451 topA 1 5 24 DNA topoisomerase I CC2881 uvrC 1 5 35 excinuclease ABC, subunit C CC3049 dnaG 1 4 20 DNA primase CC3211 3 6/4/4 8/7/7 DNA polymerase III, alpha subunit, putative CC3538 1 4 32 helicase, UvrD/Rep family CC0378 ccrM 0 4 21 modification methylase CcrMI CC1580 gyrA 0 5 5 DNA gyrase subunit A rep 1 9 56 plasmid replication initiator Purines, pyrimidines, nucleosides, and nucleotides CC3492 nrdA 1 5 11 ribonucleoside-diphosphate reductase, alpha subunit CC3588 cmk 1 4 26 cytidylate kinase CC36301 purU 2 4/4 24/27 formyltetrahydrofolate deformylase Protein synthesis and protein fate CC00101 dnaK 4 8/8/8/8 21/21/25/23 dnaK protein CC01741 gspE 2 7/4 17/16 general secretion pathway protein E CC0197 rplS 1 4 34 ribosomal protein L19 CC0484 pth 1 4 35 peptidyl-tRNA hydrolase CC0701 ileS 1 8 12 isoleucyl-tRNA synthetase CC0721 efp 1 4 27 translation elongation factor P
CC0897 trmU 1 4 18 tRNA (5-methylaminomethyl-2-thiouridylate)-methyltransferase
CC0917 1 4 26 dnaJ family protein CC1009 rsaE 2 4/4 14/15 RsaA secretion system, membrane protein RsaE CC1048 1 4 11 prolyl oligopeptidase family protein CC1240 tufB 1 4 17 translation elongation factor EF-Tu CC1248 rplC 2 4/5 32/35 ribosomal protein L3 CC1260 rplE 1 5 51 ribosomal protein L5 CC1320 valS 2 4/4 5/33 valyl-tRNA synthetase CC1376 rplM 1 4 33 ribosomal protein L13 CC1892 aspS 1 5 11 aspartyl-tRNA synthetase CC1963 clpP 2 4/4 27/29 ATP-dependent Clp protease, proteolytic subunit CC2481 pepN 4 5/5/5/4 19/21/18/25 aminopeptidase N CC2511 rpsD 1 4 26 ribosomal protein S4 CC2529 alaS 2 5/5 12/11 alanyl-tRNA synthetase CC2544 2 5/4 14/43 aminopeptidase, putative CC2672 1 7 27 Xaa-Pro dipeptidase, putative CC2809 1 4 19 peptidase, M20/M25/M40 family CC31991 tufA 1 4 17 translation elongation factor EF-Tu CC3246 1 4 16 prolyl oligopeptidase family protein
96
Table 3.2. Continued. Gene # tagging # Peptides % Coverage Gene product sites
Protein synthesis and protein fate CC33592 argS 2 8/7 22/22 arginyl-tRNA synthetase CC3504 1 5 39 peptidase M13 family protein CC3584 1 4 30 peptidase, M16 family CC3653 ffh 1 7 24 signal recognition particle protein CC3687 1 4 15 prolyl oligopeptidase family protein CC3688 1 4 21 prolyl oligopeptidase family protein CC0464 thrS 0 5 10 threonyl-tRNA synthetase CC0910 flhA 0 4 5 flagellar biosynthesis protein FlhA CC2638 0 5 6 peptidase, M16 family CC3068 secA 0 6 7 preprotein translocase, SecA subunit CC3587 rpsA 0 4 27 ribosomal protein S1 Amino acid biosynthesis and degradation CC0088 1 4 6 NAD-specific glutamate dehydrogenase CC0195 leuD 1 5 36 3-isopropylmalate dehydratase, small subunit CC0253 asd 1 4 28 aspartate-semialdehyde dehydrogenase CC0283 argB 1 4 15 acetylglutamate kinase
CC0804 putA 5 8/8/5/5/5 12/12/21/21/21 proline dehydrogenase/delta-1-pyrroline-5-carboxylate dehydrogenase
CC0957 hutU 1 4 11 urocanate hydratase CC1608 astB 1 4 14 succinylarginine dihydrolase CC1741 glnG 2 5/5 18/19 nitrogen regulation protein NR(I) CC2211 argH 1 4 20 argininosuccinate lyase
CC2274 2 7/7 26/25 methylmalonate-semialdehyde dehydrogenase, putative
CC3044 ilvD 2 4/4 13/13 dihydroxy-acid dehydratase CC3589 aroA 2 5/4 31/28 3-phosphoshikimate 1-carboxyvinyltransferase CC3606 gltD 2 5/5 24/20 glutamate synthase, small subunit CC3607 gltB 1 4 4 glutamate synthase, large subunit
CC0482 metE 0 6 10 5-methyltetrahydropteroyltriglutamate-homocysteine methyltransferase
CC1116 0 4 44 chorismate mutase, putative CC3511 hisG 0 5 35 ATP phosphoribosyltransferase CC3574 ald 0 4 23 alanine dehydrogenase Cell envelope and transport proteins CC0131 kup 2 5/4 21/18 Kup system potassium uptake protein CC0335 2 5/5 25/52 TonB-dependent receptor CC0712 feoB 1 4 33 ferrous iron transport protein B CC0808 1 8 30 HlyD family secretion protein CC0932 2 4/4 23/19 ABC transporter, ATP-binding protein CC0964 1 7 22 copper-binding protein CC0983 1 4 26 TonB-dependent receptor CC0991 1 5 12 TonB-dependent receptor CC1052 2 4/4 29/29 glycosyl transferase, group 1 family protein CC1276 1 4 42 ABC transporter, ATP-binding protein CC1516 1 4 18 penicillin-binding protein, 1A family CC1517 3 5/4/5 13/10/12 TonB-dependent receptor
97
Table 3.2. Continued. Gene # tagging # Peptides % Coverage Gene product sites
Cell envelope and transport proteins CC1623 1 5 27 TonB-dependent receptor CC17501 1 5 12 TonB-dependent receptor CC1875 1 5 25 penicillin-binding protein, 1A family
CC1985 lpxC 1 4 32 UDP-3-O-3-hydroxymyristoyl N-acetylglucosamine deacetylase
CC2091 1 4 25 ABC transporter, ATP-binding protein CC2148 1 4 24 ABC transporter, ATP-binding protein CC2299 6 10/4/4/4/4/4 25/18/40/34/26/44 fatty acid transport protein, putative
CC2378 1 4 30 NAD-dependent epimerase/dehydratase family protein
CC2832 2 4/4 39/4 TonB-dependent receptor CC2868 1 4 19 neuB protein, putative CC2924 1 4 11 TonB-dependent receptor CC2928 1 4 27 TonB-dependent receptor CC3280 1 4 30 ABC transporter, ATP-binding protein CC3500 1 4 16 TonB-dependent receptor CC3640 rfaE 2 4/4 32/38 rfaE protein CC3696 1 4 31 ABC transporter, ATP-binding protein
CC0859 0 5 19 sugar ABC transporter, periplasmic sugar-binding protein
Metabolism and energy production CC00761 fadB 2 5/6 10/12 fatty oxidation complex, alpha subunit CC02122 1 4 30 hydrolase, carbon-nitrogen family CC0257 achY 1 4 15 adenosylhomocysteinase CC0337 sucC 1 5 22 succinyl-CoA synthetase, beta subunit
CC0339 sucA 1 4 5 2-oxoglutarate dehydrogenase, E1 component
CC0365 atpF 3 5/4/5 36/40/40 ATP synthase F0, B subunit CC0672 1 4 9 cobalamin biosynthesis protein CC09721 1 4 25 tryptophan halogenase, putative CC1057 kdgK 1 4 24 2-dehydro-3-deoxygluconokinase CC1151 lldD 1 5 28 L-lactate dehydrogenase CC1224 gnl 1 4 21 gluconolactonase
CC1327 1 4 26 ubiquinone/menaquinone biosynthesis methlytransferase family protein
CC1352 2 4/4 35/34 enoyl-CoA hydratase/isomerase family protein
CC1471 ppdK 2 5/5 7/7 pyruvate phosphate dikinase CC1493 ppc 1 4 33 phosphoenolpyruvate carboxylase CC1638 1 5 11 oxidoreductase, GMC family CC1733 1 4 24/ Aas bifunctional protein, putative CC1900 1 4 30 acyltransferase family protein CC1906 gltA 1 5 18 citrate synthase CC2057 zwf 2 4/4 14/15 glucose-6-phosphate 1-dehydrogenase CC2153 1 4 19 polysaccharide deacetylase CC2162 1 4 34 aminotransferase, class I CC2355 2 5/5 20/21 sal operon transcriptional repressor SalR
98
Table 3.2. Continued. Gene # tagging # Peptides % Coverage Gene product sites
Metabolism and energy production CC2494 1 4 26 cytochrome P450 family protein CC2966 1 4 51 short chain dehydrogenase family protein CC2999 1 4 28 citrate lyase, beta subunit, putative CC3031 1 5 20 phospholipase C CC30811 mutB 1 8 23 methylmalonyl-CoA mutase, alpha subunit CC30831 1 5 20 FMN oxidoreductase CC3140 gabD 1 4 46 succinate-semialdehyde dehydrogenase CC3272 glpQ 1 5 26 glycerophosphoryl diester phosphodiesterase CC33801 kduD 2 4/4 26/35 2-deoxy-D-gluconate 3-dehydrogenase CC3527 sdhA 1 11 26 succinate dehydrogenase, flavoprotein subunit CC05441 0 5 9 nuclease, putative CC1733 0 5 27 Aas bifunctional protein, putative Transcription and regulatory proteins CC0652 3 4/4/4 10/10/18 sensory box histidine kinase/response regulator CC0921 3 4/4/4 18/18/14 sensor histidine kinase/response regulator
CC0981 1 4 32 RNA polymerase sigma-70 factor, ECF subfamily
CC1185 2 4/4 20/15 helicase, putative CC1293 1 5 41 DNA-binding response regulator CC1294 1 4 32 sensor histidine kinase CC1345 1 4 43 transcriptional regulator, TetR family CC1420 1 4 40 transcriptional regulator, GntR family CC2284 1 5 20 transcriptional regulator, LacI family CC2521 1 4 13 sensor histidine kinase/response regulator CC2661 1 4 36 transcriptional regulator, GntR family CC30751 1 5 13 sensor histidine kinase/response regulator CC3149 2 4/4 22/22 transcriptional regulator, MarR family CC3192 1 4 40 transcriptional regulator, TetR family CC3560 1 5 21 response regulator/sensor histidine kinase CC0502 rpoB 0 5 4 DNA-directed RNA polymerase, beta subunit CC0503 rpoC 0 6 6 DNA-directed RNA polymerase, beta subunit Cellular processes
CC0542 appA 1 4 12 periplasmic phosphoanhydride phosphohydrolase
CC0899 1 4 12 flagellar hook-associated protein FlaN, putative CC0909 flbD 1 4 28 transcriptional regulator FlbD CC1078 cckA 2 4/5 15/15 cell cycle histidine kinase CckA CC1281 1 4 39 tetracycline resistance protein CC1458 flbT 1 4 37 flbT protein CC1573 motB 1 4 23 chemotaxis MotB protein CC1615 ftsJ 1 4 29 cell division protein FtsJ CC2484 1 4 35 glutathione S-transferase family protein CC3043 katG 1 4 8 catalase/peroxidase CC3226 ftsH 2 7/8 20/20 cell division protein FtsH CC1553 spoT 0 4 8 guanosine-3,5-bis(diphosphate) 3-pyrophosphohydrolase
99
Table 3.2. Continued. Gene # tagging # Peptides % Coverage Gene product sites
Cellular processes CC2724 0 5 6 metal ion efflux RND protein family Unknown function CC0073 1 4 22 hypothetical protein CC0087 1 4 35 hypothetical protein CC0911 1 5 40 hypothetical protein CC1085 1 7 16 hypothetical protein CC1102 1 5 16 hypothetical protein CC1413 1 11 33 hypothetical protein CC1604 1 4 4 hypothetical protein CC1789 2 5/5 36/36 hypothetical protein CC2030 1 4 30 hypothetical protein CC2035 2 4/4 12/10 hypothetical protein CC2060 1 4 43 hypothetical protein CC2193 1 4 24 hypothetical protein CC2442 1 4 28 hypothetical protein CC2475 1 4 22 hypothetical protein CC2476 1 4 15 hypothetical protein CC2704 1 4 55 hypothetical protein CC2749 1 4 23 hypothetical protein CC3004 1 4 24 hypothetical protein CC3053 1 5 21 hypothetical protein CC3055 2 4/4 20/19 hypothetical protein CC3251 1 4 32 hypothetical protein CC3318 2 4/4 29/29 hypothetical protein CC3654 1 4 51 hypothetical protein CC0572 2 4/7 12/23 conserved hypothetical protein CC0623 1 9 19 conserved hypothetical protein CC0643 1 4 38 conserved hypothetical protein CC0645 1 4 25 conserved hypothetical protein CC0748 1 5 14 conserved hypothetical protein CC0757 1 4 21 conserved hypothetical protein CC0944 1 5 21 conserved hypothetical protein CC1423 1 4 10 conserved hypothetical protein CC1624 2 4/4 25/28 conserved hypothetical protein CC1766 2 4/4 31/29 conserved hypothetical protein CC1967 1 6 15 conserved hypothetical protein CC2105 1 4 21 conserved hypothetical protein CC2577 1 4 34 conserved hypothetical protein CC2671 1 4 33 conserved hypothetical protein CC2790 1 4 10 conserved hypothetical protein CC2849 1 4 27 conserved hypothetical protein CC3416 1 4 22 conserved hypothetical protein CC3418 1 6 25 conserved hypothetical protein CC3575 2 4/4 27/27 conserved hypothetical protein
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Table 3.2. Continued. Gene # tagging # Peptides % Coverage Gene product sites
Unknown function CC3666 2 4/4 28/25 conserved hypothetical protein CC3684 2 4/4 13/13 conserved hypothetical protein CC3761 1 4 34 conserved hypothetical protein CC0140 1 5 10 ComM protein CC0587 1 6 30 pentapeptide repeat family protein CC0766 cgpA 1 4 35 GTP-binding protein CgpA CC1654 5 4/4/4/4/4 11/9/10/10/10 GTP-binding protein CC2333 1 4 38 phage SPO1 DNA polymerase-related protein CC2823 1 5 12 TldD/PmbA family protein CC3010 1 4 28 DnaJ-related protein CC3121 1 5 18 phytoene dehydrogenase-related protein CC3128 2 4/4 22/20 TPR domain protein
Proteins are grouped into functional categories based on annotations from GenBank (Nierman, et al., 2001). For each protein, the gene number, the gene name if available, number of peptides identified by MALDI-TOF analysis, and protein name are listed. 1. Genes that are upregulated in microarray analysis (The relative mRNA ratio of ∆ssrA/wild type ≥ 1.6). 2. Genes that are downregulated in microarray analysis (The relative mRNA ratio of ∆ssrA/wild type ≤ 0.6).
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Table 3.3. Functional distribution of identified substrates.
Functional category No. of identified substrates
No. of total proteins in functional category
% of identified substrates1
DNA metabolism 13 84 15.5
Protein synthesis and protein fate 34 261 13.0
Amino acid biosynthesis 11 93 11.8
Cell envelope, transport and binding proteins
32 397 8.1
Metabolism
39 619 6.3
Purines, pyrimidines, nucleosides, and nucleotides
3 48 6.3
Cellular processes
12 209 5.7
Transcription and regulatory functions
18 324 5.6
Unknown function
55 1703 3.2
Biosynthesis of cofactors, prosthetic groups, and carriers
2 83 2.4
Total 219 1. Values are calculated by:
(No. of identified substrates/No. of total proteins in functional category) × 100.
102
Table 3.4. Occurrence of a putative motif inducing SsrA tagging. Gene Strand1 Distance from Motif sequence tagging site (bases)2 CC0010 + 177 C G A C G C C G A G T T C G A CC0010_1 + 144 C G A C G C C G A G T T C G A CC0010_2 + 39 G G A C A G C G A C A T C G A CC0010_3 + 90 G A A G A C C G C G A T C G A CC0073 - 92 C G C C A A C G G C G T C G A CC0076 + 90 C G G C G A C A A G T T C G A CC0076_1 + 120 C C A G G C G C T G A T C C A CC0087 - 47 C G C G G T C A T C G T C G A CC0088 + 69 G A A C G C G G T C A T C G T CC0131 + 75 G G C G A C G G T G A T C G C CC0131_1 - 109 G G T G A C G A T C A T C G T CC0140 - 2 C G A C G T C T T C G T C G C CC0174 + 6 G A C C G C C C A G A T C G C CC0174_1 - 113 C A T G A T C T T G A C C G G CC0195 - 181 C G C G A T G C G G A T C G A CC0197 - 173 C G A A A T C C T C A T C C T CC0212 + 114 C G C C A A C C G C A T C G G CC0253 + 180 C A T C G T C G T G G T C G A CC0257 + 162 A G A C G C C C T G T T C A A CC0283 + 168 C G C C A T G C T G G T C G A CC0335 + 177 C A A G T A C G T G A C C A A CC0335_1 + 177 C A C G G A C A T C A C C G T CC0337 + 171 C G A C A C C G T C G C C G A CC0339 + 120 C G A C G A C A A G A T C A A CC0346 - 7 C A T G G C C T T G A T C A G CC0346_1 - 175 G G G C T T C T T C A T C G A CC0346_2 - 64 C A T G G C C T T G A T C A G CC0365 + 152 G A C C G C G G A C A T C G C CC0365_1 + 72 G G A C G C C G C G A T C G G CC0365_2 + 69 G G A C G C C G C G A T C G G CC0484 - 129 C G A G A A C A T C G T C A G CC0542 + 105 C G C G G C G G T C T T C G C CC0572 - 76 C T A A A A C G T C A T C A A CC0572_1 + 27 G G T C G A G G A C A T C A A CC0587 - 155 C G C G G C C C T G A T G G A CC0623 + 141 G G C C A A C A A G T T C C A CC0643 - 158 C C T C G A C C T G G C C G A CC0645 + 143 C G C C C A C A G G A T G G A CC0652 + 51 C A A G G G G C T G G T C G A CC0652_1 + 183 C G C C G A G G T G G T C G A CC0652_2 + 15 C G C C G C C C T G G T C G A CC0672 + 129 G A A G G A G A A C A T C G A
103
Table 3.4. Continued. Gene Strand1 Distance from Motif sequence tagging site (bases)2 CC0701 - 122 T G C G A A G C A G G T C G G CC0712 + 3 C G A G G C C T T G G T C G A CC0721 + 48 T T T C G A C C T G T C C G A CC0748 - 26 C T T C T C G A A G A T C G G CC0757 - 101 C G G C G A C C T G T T C G A CC0766 + 15 G G T C G A C A A G G T C G T CC0804 + 117 C G T C A A C A T C A C C G C CC0804_1 + 21 C G T C A A C A T C A C C G C CC0804_2 + 66 C G G C G T C A A G A T C G A CC0804_3 + 69 C A A G G C G C T G A T C G A CC0804_4 + 105 C A A G G C G C T G A T C G A CC0808 + 126 C G C C A T C G T G A T C A T CC0897 - 129 G C C G A C G C T G A T C A G CC0899 - 61 C G T G T T G G T G T T C G A CC0909 + 3 C A T C G G C A A G T T C G A CC0911 + 63 C G T C A A G C T G G T C G A CC0917 - 146 C G G G A T C C A G A T C G A CC0921 - 34 G G C G A C G G T G A T C G A CC0921_1 - 64 G G C G A C G G T G A T C G A CC0921_2 + 171 G C T C A C C G T G G T C G A CC0932 - 131 C C A C A C C G T C A T C A T CC0932_1 - 26 C C A C A C C G T C A T C A T CC0944 + 123 C G T G G C G G T G A T C C A CC0957 + 95 G G T C G T C C C G T T C G C CC0964 + 102 C G T G G T G C T G A T C A A CC0972 + 63 C G C C A A G G T G A T C A T CC0981 - 178 C G G C T T C G T G T T C G T CC0983 + 120 G G T C A C G G T G G T C G G CC0991 + 51 C C T C G C G G T G T T C A A CC1009 - 58 C T C C T C G C G C A T C G T CC1009_1 + 33 G C A G A A C A A G C T C G A CC1048 + 111 G A A G A T G C T G A C C G A CC1052 + 114 C G T C A A G C T G G T C G G CC1052_1 + 183 C G T C A A G C T G G T C G G CC1057 - 104 G A A C A A C G C C A T C G A CC1078 + 96 C G T C A A G C T G A T C A C CC1078_1 + 126 G G G C A A G A T C T T C G A CC1085 + 84 C G A C T A C A T C G T C A G CC1102 + 105 G G T G G C G C T G G T C G G CC1151 + 83 C G A C G T C G C G A T C G G CC1185 + 177 C G C C G C C T T C G C C G A CC1185_1 + 138 C A T G G C C C T G G T C G A
104
Table 3.4. Continued. Gene Strand1 Distance from Motif sequence tagging site (bases)2 CC1224 + 18 G G T C G C C C T G G T C G A CC1240 + 75 C G T C G A G C T G A T C A C CC1248 + 173 A C T C G A C G T C A T C A A CC1248_1 + 30 C C T G A T C C T G G T C A A CC1260 + 90 C A T G G A C A T C A T C G T CC1276 + 174 C G T C G A G A T C G T C G A CC1281 - 24 A G C C G C G A T G C T C G A CC1293 + 30 G A C G A A C G T G G T C G A CC1294 + 162 C G C C A A G G C C T T C G A CC1320 - 158 A A A C A C C G T C A T C C C CC1320_1 + 114 G C A G A A G G A G G T C G A CC1327 + 21 C T A C G C C G A G A C C A A CC1345 - 86 C G A C G A C C T G A T C G A CC1352 + 180 G A A G G C G A T C A T C G C CC1352_1 + 162 G A A G G C G A T C A T C G C CC1376 - 181 G T T G A T C G T G A T C G A CC1413 + 69 G C T G G A C T T C A T C G A CC1420 + 22 C G A G A C G C T C G C C A C CC1423 + 66 C G T C G A C G G C A T C A A CC1458 + 172 C G C C A C G C G G A T C G A CC1471 + 27 G G C C A A C A A G G T C G A CC1471_1 + 150 G G C C A A C A A G G T C G A CC1493 + 165 G C C G G C G A T C A T C G A CC1516 + 39 C G A G A C C A T C G T C A A CC1517 + 45 C G A G A A C C T G T T C G A CC1517_1 + 39 C A C C A A C A T G A T C G G CC1517_2 + 138 C A A C A C C G G C T T C A A CC1522 + 99 G C C C G A C G T G A T C G A CC1573 + 144 C A A G G C C A A C T C C G A CC1604 - 137 A G T C A A G A T A A C C A A CC1608 + 125 C G C C A C G G T C G C C G A CC1615 + 71 C G G C G G G C T G A T C G C CC1623 + 45 C G T G A A G C T G A C C G C CC1624 - 94 C A T C A C G A A G T T C G G CC1624_1 - 4 G A T C A C G A A G A T C G C CC1638 + 170 G A A G A C G C T G G T C G C CC1654 + 111 C C A G A A G A A G G T C G C CC1654_1 + 161 C C C C C A C G G G A T C A G CC1654_2 - 7 G G C C G T C T T G G T C C C CC1654_3 + 3 C A A G G T C G G G A C C A A CC1654_4 + 30 C C A G A A G A A G G T C G C CC1733 + 45 C G C G G C G C A C A T C G A
105
Table 3.4. Continued. Gene Strand1 Distance from Motif sequence tagging site (bases)2 CC1741 + 105 C G T G G A G A T G A C C C G CC1741_1 - 50 C G T C A G G C T G A T C G G CC1750 + 125 C G C C T T G A T G A T C C A CC1766 + 114 C T T C A C G C G G T T C G A CC1766_1 + 157 C G G C A T C G T G A T C G G CC1789 - 98 C A C C A C G C T G T T C A A CC1789_1 - 152 C A C C A C G C T G T T C A A CC1844 - 140 C G A A A C C C A C A T C G C CC1844_1 + 57 C C T G A T C C A G A T C G C CC1875 + 144 C G T G G C G C T G G T C G G CC1892 - 7 C T A C A C C T T G A T C G G CC1900 - 113 G G C C T A G A T C A T C G A CC1906 + 12 G A C C A A C A T G T T C A C CC1926 + 165 C G C C A C C C A G T T C A A CC1963 - 53 C G T A G A C G T G G T C G A CC1963_1 - 77 C G T A G A C G T G G T C G A CC1967 + 56 C G C C A T G A A G A T C G T CC1985 + 45 C G C C G T G G T C A T C G A CC2030 - 148 C A T G A C C C G G A T C A G CC2035 + 18 C G C C A G C C T G A T C G C CC2035_1 + 150 G G C G G C G C T G A T C A A CC2057 - 56 G C T C G A C C T C A T C G C CC2057_1 - 38 G C T C G A C C T C A T C G C CC2060 - 44 A C A G G T C A T C A T C A A CC2091 + 27 C G T C A T C C A C A T C G A CC2105 + 180 C T C C A A C G T G A C C G G CC2148 + 108 C A A G G T G C T G T T C A A CC2153 + 150 C A A G A C C G A C G T C G A CC2162 + 183 C C T G A C C C T G G C C G A CC2193 + 34 C G A G G T G C T G T T C G A CC2211 - 160 C G T G G C G G T G A T C G A CC2246 - 143 C C A G G C C C T C G T C G T CC2274 + 87 C G T C G C C A T C T T C A C CC2274_1 + 105 C G T C G C C A T C T T C A C CC2284 - 166 C T G G A C G C T G A T C G A CC2299 + 6 C C T G A T G A A G G T C G A CC2299_1 + 153 G T T C T T C A A C T T C G A CC2299_2 + 9 A A C C G C C T T G A T C A A CC2299_3 - 167 G G T C G C G C T G G T C A T CC2299_4 - 41 G G T C G C G C T G G T C A T CC2299_5 + 111 C G T C G C G C T C T T C A T CC2333 - 55 G A T G G T C C A G T T C A T
106
Table 3.4. Continued. Gene Strand1 Distance from Motif sequence tagging site (bases)2 CC2353 + 185 C A C G A T G A A G A T C A G CC2355 + 33 C C A C G C G C T C A T C G A CC2355_1 + 27 C C A C G C G C T C A T C G A CC2378 + 183 G A T G G A C A T G A T C A A CC2442 - 178 G A T C C A G C A G A T C G G CC2451 + 126 C G T C G A C C T G A C C A G CC2475 + 70 A G A C T T G C T C A T C G A CC2476 - 105 A G A C T T G C T C A T C G A CC2481 - 82 C T C C A C C G T G G T C G C CC2481_1 + 105 G A T C G A C A A C T T C T A CC2481_2 - 91 C T C C A C C G T G G T C G C CC2481_3 + 39 C A G C G T C C G G A T C G A CC2484 - 141 C G A C A G C C T C A T C C C CC2511 - 133 C G C G A T C C G G T T C A A CC2521 + 66 G A A C G G C G T C A T C G C CC2529 + 111 C G A C G C C C T G A T C G C CC2529_1 + 72 C A C G G C C C T G A T C G C CC2544 + 3 C G T G C C G G T C A T C A A CC2544_1 - 118 G G A C G T C G A G A T C G A CC2577 + 137 G A A G A A G C T G T T C G C CC2661 - 184 C T T G A C C T T C G T C A A CC2671 + 153 C G A G G C G C T G T T C G A CC2672 - 55 C A T C A C G A A G A C C G G CC2704 - 31 C G C A G C C G A C A C C A G CC2749 + 108 C A A C G A C G T C A C C G A CC2790 - 151 C A C C G C C C A G A C C A G CC2809 + 81 C A A G G C C A A G G C C G A CC2823 + 51 C A T C A A C G T C A T C A A CC2832 + 183 C G G C G T C C T G T T C G G CC2832_1 - 155 C G T C A C G G A C G C C G C CC2849 - 61 T G A G A A G G T C A T C A A CC2868 - 70 C A T C G A C C A G T C C A G CC2881 + 39 G C T C T A C C A G A T C A A CC2924 + 171 C C C G A C G G T G G C C A A CC2928 - 112 C G T C G A C A G G A T C G A CC2966 - 121 C G A C T C C G A C A T C G A CC2999 + 102 C G C C A A G A T G G T C G A CC3004 + 30 C G A C A A G A T G A C C G A CC3010 - 16 A T A G G C C T T G A T C A C CC3031 + 27 C G A C G A C C T G A T G G G CC3043 + 147 C G T C G A C C T G A T C T T CC3044 + 72 C C C G A T C C T G A T C G A
107
Table 3.4. Continued. Gene Strand1 Distance from Motif sequence tagging site (bases)2 CC3044_1 + 117 C C C G A T C C T G A T C G A CC3049 + 126 G T A C G G C C T G T T C G A CC3053 - 41 C G C G G C C C T C G T C G G CC3055 - 143 G G G C G A G G T G A T C A A CC3055_1 - 26 C A A G A C C G A G G T C G A CC3075 + 117 C G A C A C C G G C A T C G G CC3081 + 99 C A T G G A G G T C G T C T A CC3083 + 88 C C T C G G C G T G A T C G G CC3121 - 111 C G G G G C C C A G A T C G T CC3128 + 186 C T C G A C C C T G G T C G A CC3128_1 + 123 C T C G A C C C T G G T C G A CC3140 + 51 G C T C A A C A T C G T C A C CC3149 + 166 C G A C G C C C A G A T C A C CC3149_1 + 163 C G A C G C C C A G A T C A C CC3192 + 165 C G A C A T C C T G C T G A A CC3199 + 75 C G T C G A G C T G A T C A C CC3211 + 171 C C A G G A G G T G G C C G A CC3211_1 - 184 G G C G G C C A G C A T C G A CC3211_2 + 6 C G A C A A C G T G A T C C A CC3226 + 138 C G A C A G C G A G G T C A A CC3226_1 + 54 G A C C G A C A A G A T C G A CC3246 - 7 C G T C G C C A G C T T C A A CC3251 - 23 C T G G A T C A A G A T C G G CC3272 + 15 C G A C G G C G T G T T C A G CC3280 + 144 G A T C G A G C A G A C C G C CC3318 - 181 C A T G G A G C A G A T C A A CC3318_1 - 163 C A T G G A G C A G A T C A A CC3359 + 6 C C T G A C C C T G G C C G A CC3359_1 + 174 C A A C T T C G T G G C C G A CC3380 - 2 C G A C T T G A T C A T C G A CC3380_1 - 67 C G A C T T G A T C A T C G A CC3416 + 54 G G A C G A C T A C A C C A A CC3418 + 66 C G A C G C G A T C A T C G C CC3492 - 5 G C T G A T C C A G T T C G A CC3500 + 36 G A T C G A C G T G T T C G G CC3504 + 45 C A A G G A C G A G T T C A C CC3527 + 60 C A A G G T G A A G A T C G A CC3538 + 117 C G A G G C G A T G T T C G C CC3560 + 153 C G C C A C C C T G A T C G A CC3575 - 65 G A T C G A G C A C G T C G C CC3575_1 - 104 G A T C G A G C A C G T C G C CC3584 + 162 C G C C G C C C T G A T C A T
108
Table 3.4. Continued. Gene Strand1 Distance from Motif sequence tagging site (bases)2 CC3588 + 65 G A T C A A G A A C A T C G C CC3589 + 105 C G T C G A G G T C A T C G A CC3589_1 + 129 C G T C G A G G T C A T C G A CC3606 + 156 C G A C G G C G T G T T C G C CC3606_1 + 81 C T C G A C G C T G A T C G C CC3607 + 21 C A C C A C G G T G G T C G A CC3630 + 51 C G T G A A G C T G A T C G G CC3630_1 + 90 C G T G A A G C T G A T C G G CC3640 + 72 C G C G A C C C T G A T C A A CC3640_1 + 48 C G C G A C C C T G A T C A A CC3653 + 186 G G C C G A C A T G T T C A A CC3654 - 103 G A A G A A G A A G T T C G G CC3666 + 90 G G T G G T G C T G G T C C A CC3666_1 + 72 G G T G G T G C T G G T C C A CC3684 + 57 C T T C A C C G A G A T C G C CC3684_1 + 117 C T T C A C C G A G A T C G C CC3687 + 120 G C C C G C C C A G T T C G A CC3688 + 180 C G T C A C C T T C T T C G A CC3696 - 46 C G C C G C C G T G A T C G G CC3761 - 31 G G T C A C C G A C A T C A C
1. DNA strand where the motif is found. 2. Distance from the first base of codon encoding the SsrA-tagged amino acid of each substrate. 3. The consensus sequence is generated by WebLogo (Crooks et al., 2004).
Multilevel consensus sequence3
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Chapter 4
Caulobacter crescentus requires SsrA activity for plasmid
replication
110
ABSTRACT
To colonize different species of bacteria, plasmids must have an efficient
replication machinery. Some components of this machinery are encoded by the plasmids
themselves while others are encoded by the host chromosome. Here we show that one
host-encoded factor is a small regulatory RNA, SsrA, involved in bacterial gene
regulation by a trans-translational mechanism. In Caulobacter crescentus strains lacking
SsrA RNA or its protein cofactor, SmpB, broad-host-range plasmids that belong to
pBBR1 family are not maintained due to a specific defect in DNA replication. A mutant
plasmid, pPT1 was isolated from a genetic screen for plasmids that can bypass the
requirement for SsrA activity. pPT1, containing a truncated gene for the replication
initiator, Rep, can be stably maintained in the absence of ssrA or smpB. In addition, Rep
is tagged by SsrA at its C terminus. These results suggest a model in which SsrA tagging
may be required for plasmid replication by affecting physical properties of Rep.
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INTRODUCTION
Plasmids are self-replicating extrachromosomal DNA elements that are present in
most species of bacteria (Lederberg, 1998). Plasmids contribute to bacterial genetic
diversity and adaptability by encoding a variety of genetic determinants. For example,
some plasmids harbor resistance genes that allow their hosts to survive contact with
bactericidal agents such as antibiotics and heavy metals. In pathogenic bacteria, genes
encoding important virulence factors are present on plasmids. In addition, plasmids can
incorporate and deliver genes, thus favoring genetic exchanges between bacterial
populations (Actis et al., 1999). The ability of some plasmids to propagate in many
species of bacteria has raised questions about the nature of the determinants of plasmid
host range.
The initiation of DNA replication is molecule specific, and thus this step is of
great importance for the propagation of a plasmid in a specific host (Kues and Stahl,
1989). In addition to plasmid-encoded cis and trans elements, replication initiation is
dependent on various host-specific replication factors. Therefore, many plasmids can
replicate only in one or a few closely related hosts (narrow host range). In contrast, some
plasmids are able to replicate and maintain themselves in many distantly related bacterial
species (broad host range) (Kues and Stahl, 1989).
The origins of plasmids that replicate via the theta mechanism possess several
characteristic functional elements: multiple direct repeats called iterons which serve as
specific binding sites for the plasmid-specific replication initiation protein, Rep; one or
more elements (DnaA boxes) for the host replication initiation protein, DnaA; and an AT-
rich region where DNA duplex destabilization and assembly of host initiation factors
112
occur (del Solar et al., 1998). In the theta-replicating plasmids, initiation of replication
generally starts with the Rep protein binding to the iterons forming a preinitiation
complex. Subsequently, the Rep-DNA complex in combination with DnaA facilitates the
opening of the strands in the AT-rich region. These initiation proteins also promote the
sequential assembly of components of the replisome complex including DnaB helicase
and DNA polymerase III holoenzyme at the origin of replication (del Solar et al., 1998).
At present, with the exception of DnaA, DnaB, and DNA polymerase, the
knowledge concerning host factors involved in plasmid replication and reasons for
variation in host ranges are incomplete. It can be assumed that host factors which have
not been discovered could, in part, be responsible for the propagation of plasmids in
various hosts. Here, we report the identification of a host-encoded RNA, SsrA, that is
required for plasmid propagation in Caulobacter crescentus. SsrA is a regulatory RNA
that is capable of intervening in expression of selected genes (Chapter 3). SsrA activity
on stalled ribosomes during translation results in the addition of a short peptide to the C
terminus of the nascent protein, and in many cases this peptide targets the protein for
rapid degradation (Karzai and Sauer, 2001; Keiler et al., 1996). SsrA acts in combination
with a protein cofactor, SmpB, which is required for stable association of the RNA with
the ribosome (Karzai et al., 1999) and for protection of the RNA from degradation by a
ribonuclease (Hong et al., 2005). C. crescentus strains deficient in SsrA activity have a
delay in the normal timing of initiation of chromosomal DNA replication resulting in a
defect in cell cycle progression (Keiler and Shapiro, 2003). We report here that SsrA is
essential for plasmid replication in C. crescentus, and that mutations in the origin-binding
protein Rep can bypass the requirement for SsrA.
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RESULTS
Plasmid maintenance in ssrA deficient strains
Plasmids classified to three different incompatibility groups have been identified
that will be maintained by wild-type C. crescentus. pBBR1 and pRK2 (IncP group), and
pRSF1010 (IncQ group) can replicate and be stably maintained in C. crescentus. We
tested the ability of strains lacking SsrA activity due to a deletion of either ssrA or smpB
to maintain pBBR1- or pRK2-based plasmids transferred by electroporation or
conjugation (Figure 4.1). Wild-type cells transformed with pBBR1-based plasmids
carrying antibiotic resistance genes for chloramphenicol (pJS14) or kanamycin (pKJS2)
produced >100 colonies per nmol of DNA, but transformation into the ∆ssrA and ∆smpB
strains produced no colonies on selective medium, indicating a decrease of more than five
orders of magnitude in plasmid acquisition. Similar results were obtained for pBBR1
derivatives carrying resistance genes for spectinomycin and gentamycin (data not
shown). pBBR1-based plasmids that contained a copy of the ssrA gene (pKK838) could
be efficiently transformed into both wild-type and ∆ssrA cells, but not into the ∆smpB
strain. Conversely, a similar plasmid that contained a copy of the smpB gene (pKK840)
could be efficiently transformed into wild-type or ∆smpB cells, but not into the ∆ssrA
strain. Because cells lacking SsrA activity could acquire a plasmid bearing a
complementing gene, these cells are competent for transformation, and the lack of growth
under selective conditions after transformation with non-complementing plasmids must
be the result of a defect in plasmid maintenance. We also mobilized plasmids into C.
crescentus strains by conjugation from E. coli strain S17-1, which contains the IncP
group transfer functions of RK2 integrated into the chromosome (Simon et al., 1983). As
114
found for the transformation experiments, colonies were produced after conjugation for
all plasmids in wild-type C. crescentus, but only for plasmids carrying an ssrA gene in
the ∆ssrA strain and only for plasmids carrying an smpB gene in the ∆smpB strain.
To test if these results were unique to pBBR1 plasmids, we repeated the
transformation and conjugation experiments using RK2-based plasmids. A plasmid
derived from RK2 (pRK290) could be transformed into wild-type cells and strains
deleted for ssrA or smpB (Figure 4.1). This result indicates that ssrA- or smpB-deficient
strains are specifically defective in maintaining plasmids with the pBBR1 origin.
Furthermore, this result demonstrates that plasmids can replicate in ssrA-deficient strains
and that plasmid segregation is not completely abrogated. Therefore, it is likely that there
is a problem with regulation of DNA replication from pBBR1 plasmids. We tested the
ability of Escherichia coli MG1655 lacking ssrA to maintain pBBR1 plasmids, and found
that there was no difference in transformation efficiency between wild-type and ∆ssrA
strains (data not shown). Therefore, E. coli MG1655 does not require SsrA for
maintenance of pBBR1 plasmids.
Selection for mutants that can maintain plasmids in the absence of SsrA activity
To identify sequences on pBBR1 that may cause the ∆ssrA phenotype, we
conducted a genetic selection for mutations that would by-pass the requirement for SsrA
(Figure 4.2). Mutations were introduced in plasmid pKJS2 by replication in the E. coli
mutD strain. This strain has a mutation in the mutD (dnaQ), which encodes the ε subunit
of DNA polymerase III holoenzyme (3’ to 5’ exonuclease) and thus has an increased
spontaneous mutation frequency relative to wild-type E. coli (Speyer et al., 1966).
115
Plasmid DNA was purified from this strain and transformed by electroporation into an
ssrA-deficient Caulobacter crescentus strain. Cells bearing a mutant plasmid that could
be maintained in the absence of ssrA were selected by growth on kanamycin. One mutant
plasmid, pPT1 was recovered, and this plasmid could subsequently be transformed into
wild-type, ∆ssrA, and ∆smpB strains with high efficiency (Figure 4.1). Sequencing of the
pPT1 plasmid revealed a 1.8 kb deletion that eliminated the 3’ end of the open reading
frame of the rep gene (Figure 4.3A and B). The other two known genes on pKJS2, nptII
encoding a kanamycin-resistant gene and mob involved in the mobilization of the plasmid
(Antoine and Locht, 1992) were not affected by this deletion. The maintenance of pPT1
in ∆ssrA strains could be due to the truncation of the Rep protein, the sequences removed
by the deletion, or both. To distinguish among these possibilities, different mutant pKJS2
plasmids were made (Figure 4.3C) and tested for replication in ∆ssrA strain. First, we
engineered a nonsense mutation in pKJS2 that truncates the Rep protein by 61 amino
acids but leaves the rest of the plasmid unchanged (Figure 4.3C panel 3). We also
engineered point and nonsense mutations in rep on pKJS2 such that the mutant pKJS2
expresses the same Rep as pPT1 but does not have any deletions (Figure 4.3C panel 4).
Second, a deletion was engineered to remove sequences downstream from Rep, but leave
the rep gene largely intact (Figure 4.3C panel 5). This excision removes 67% (= 1.2
kb/1.8 kb) of the sequences deleted in pPT1 and results in a change in the last seven
amino acid sequences of Rep from RVTLPRR to LKTVVIH. Third, the two mutations
described above were combined to produce a plasmid with truncated Rep and a large
downstream deletion (Figure 4.3C panels 6 and 7). Each of these mutant pKJS2 plasmids
were efficiently transformed and maintained in wild-type but not in the ∆ssrA strain (data
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not shown), suggesting that either further excision of the other 33% of the sequences
deleted in pPT1 or precise construction of pPT1 is required for pPT1to be maintained in
the ∆ssrA strain.
SsrA tags Rep at the C terminus
In an independent line of investigation, we identified Rep as a substrate for SsrA
in C. crescentus. A global survey of SsrA substrates was conducted as described in
Chapter 3. In this survey, the last six codons of the SsrA tag reading frame were changed
to histidine codons (Figure 3.1), so that proteins tagged by SsrA would bear a C-terminal
six-histidine sequence. These histidine residues both protected the tagged substrates from
rapid proteolysis and facilitated purification. The substrate proteins were purified by Ni2+-
NTA affinity chromatography, separated by SDS-polyacrylamide gel electrophoresis
(PAGE), and identified by mass spectrometry.
Rep was identified in the survey because it was on the plasmid used to deliver the
SsrA-His6 variant (pJS14-ssrA-His6). To confirm that Rep is a substrate for SsrA, we
generated a version of the rep gene encoding a Strep-tag II at the 5’ end of the open
reading frame. This gene was integrated into the xylX locus of a strain bearing the ssrA-
His6 gene at the ssrA locus such that expression of Strep-Rep could be induced by xylose.
These cells were grown in the presence of xylose, Strep-Rep-SsrA-His6 was purified by a
tandem chromatography protocol using Ni2+-NTA followed by Strep-tactin resin. The
most abundant Strep-Rep-SsrA-His6 band on a SDS-PAGE was digested with trypsin and
analyzed by mass spectrometry to identify SsrA tagging sites (Figure 4.4). By searching a
database containing Rep protein sequence with the SsrA-His6 tag after each amino acid,
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we identified eight tryptic peptides including the junction peptide containing both Rep
and the SsrA-His6 tag sequences. In all, 58% of the intact mass of Rep was recovered. In
the junction peptide, the SsrA-His6 is fused to the C terminus of Rep, indicating that Rep
is tagged by SsrA at the C terminus. Furthermore, because Rep was expressed from an
ectopic locus in the chromosome in this experiment, tagging requires only the coding
sequence, and not 3’ untranslated sequences or other factors unique to the normal genetic
locus of rep.
Regulation of Rep by SsrA tagging
The requirement for SsrA activity could be due to properties of the tagged Rep
protein. To investigate whether addition of the wild-type SsrA tag was required for
plasmid replication, we transformed ∆ssrA cells with a variant of ssrA that can release
stalled ribosomes and tag the nascent proteins but does not target these tagged proteins
for degradation. In this variant, SsrA-DD, the final two codons of the tag reading frame
have been changed from Ala-Ala to Asp-Asp (Keiler et al., 2000). We found that pJS14
carrying the ssrA-DD variant (pKK839) is transformed into the ∆ssrA strain with the
same efficiency as pJS14 carrying wild-type ssrA (Figure 4.1), indicating that the wild-
type SsrA tag is not important for plasmid maintenance. It is possible that tagging of Rep
with the SsrA-DD peptide may have the same effect on Rep activity or stability as
tagging with the wild-type SsrA peptide. Such an effect might occur if the C terminus of
Rep is required for activity or folding, such that tagging with any peptide will activate
Rep or promote proper folding.
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Alternative possible role for SsrA in the regulation of Rep is to limit the protein
concentration. If over-production of Rep was lethal to the host cell, then expression of
Rep in the absence of ssrA might kill the host cells directly. To determine if this model is
possible, we placed the rep gene under control of the PxylX promoter at the xylX locus on
the chromosome of both wild-type and ∆ssrA C. crescentus and expressed rep
constitutively by growing cells in the presence of xylose. In the presence of xylose, both
strains produced significantly more Rep protein than wild-type cells bearing pKJS2
(Figure 4.5A), but there were no changes in cellular morphology when examined by light
microscopy. In addition, the growth rate of the rep-expressing strains during log-phase in
defined medium with xylose was not significantly different from that of C. crescentus
which does not have rep on the chromosome (Figure 4.5B). To further test whether the
ability of pPT1 to bypass the requirement for SsrA is due to changes in the expression
level of rep, we examined the amount of Rep protein in wild-type and ∆ssrA cells
containing pKK838, pKK839, or pPT1 (Figure 4.6). There were no significant
differences in the levels of Rep in any of these strains. The small increase observed in the
∆ssrA strain containing either pKK838 or pKK839 could arise from an increase in the
plasmid copy number due to ssrA deletion. Although formally possible, this is unlikely,
because the two strains express the same steady-state levels of mature SsrA as the wild-
type (Keiler and Shapiro, 2003). If the observed increase is real, however, this amount of
Rep should not be deleterious because cells are tolerant to significantly higher levels of
Rep. Taken together, these results indicate that the expression of rep in pPT1 is similar to
that in a pBBR1-based plasmid with wild-type rep. Thus, the ability of pPT1 to bypass
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the requirement for SsrA activity is not due to changes in the expression level of the rep
allele.
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DISCUSSION
In this chapter, we demonstrated that a plasmid replication initiator, Rep is tagged
in wild-type C. crescentus. The use of a variant ssrA gene encoding SsrA-His6 allowed us
to purify the tagged Rep and identify the tagging site by mass spectrometry. The
identification of the junction peptide containing the last six residues of Rep and the SsrA-
His6 tag indicated that tagging occurred at the last amino acid of Rep. This tagging event
is required for the plasmid replication and maintenance in ∆ssrA strain, but the C-
terminal truncation of Rep removes the requirement of SsrA. Furthermore, we
demonstrated that neither SsrA tagging nor truncation of Rep altered the levels of Rep
expression, suggesting that the primary function of SsrA tagging may not be to control
Rep protein level.
Given that SsrA-mediated tagging promotes degradation of potentially toxic
peptide products by marking them with a degradation tag, and rescues ribosomes stalled
on aberrant mRNAs (Karzai and Sauer, 2001; Keiler et al., 1996), a model that may
explain the observed experimental results can be proposed. The mRNA encoding Rep
may have a sequence or structure that causes a ribosome to become stalled at some
position upstream or near the stop codon. Truncated forms of Rep accumulate in ∆ssrA
due to the ribosome stalling and prevent plasmid replication by competing with full-
length Rep for DNA-binding sites or other replication factors and repressing replication.
Under wild-type conditions, these dominant forms of Rep are tagged by SsrA and
degraded, resulting in plasmid replication. In pPT1, the unidentified determinant causing
the ribosome stalling near the 3’ end of rep mRNA may be removed by the deletion.
Therefore, there will be no ribosome stalling that needs to be rescued by SsrA. This
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unique feature of pPT1 may allow it to bypass the requirement of SsrA activity. In
support of this model, Western analysis of Rep overexpressed from the xylX locus in
∆ssrA strain displayed a smear below the full-length Rep (Figure 4.5A), indicating that
truncated forms of Rep are indeed produced in the absence of SsrA activity.
A similar mechanism has been observed in SsrA-mediated regulation of Mu
repressor for prophage induction in E. coli (Ranquet et al., 2001). Lysogenic development
of bacteriophage Mu is established by an immunity repressor inhibiting phage
transposition functions required for phage replication (Symonds et al., 1987). In the
absence of SsrA activity, truncated forms of Mu repressor resembling dominant, heat-
stable repressor mutants are observed (Ranquet et al., 2001). These truncated repressors
prevent induction by maintaining binding to the operator even under inducing conditions.
Like the case for Rep in C. crescentus, normal Mu induction can be restored in ∆ssrA by
SsrA-DD, suggesting that without the need for proteolysis addition of SsrA tag may
abrogate the DNA-binding function of the Mu repressor.
Although the proposed model is plausible, a couple of questions remain to be
addressed. First, if SsrA acts to remove partially synthesized Rep from the cell by tagging
it for degradation, why does ssrA-DD, encoding a protease-resistant tag rescue the ∆ssrA
phenotype related to plasmid replication and aforementioned phage Mu induction? One
possible explanation is that the SsrA-DD tag sequences, at some low level, signal
proteolysis. Accordingly, this level of proteolysis would be sufficient to support plasmid
replication or phage induction. Although formally possible, the stability of SsrA-DD-
tagged Rep needs to be tested to validate this scenario. Alternatively, SsrA-DD tagging of
the truncated forms of Rep may induce conformational changes rendering them incapable
122
of DNA binding or other replication factors. This inactivation of the dominant forms of
Rep will allow plasmid replication. Although intriguing, the details of this mode of SsrA-
DD action are speculative at this stage.
Secondly, why does the ssrA-DD allele not complement the cell cycle delay
phenotype observed in the ∆ssrA C. crescentus (Keiler and Shapiro, 2003)? It is possible
that the mode of SsrA action in control of plasmid replication is distinct from that for cell
cycle progression. As described before, SsrA may regulate plasmid replication by
controlling Rep activity. In the case of cell cycle control, regulation of a specific gene(s)
by SsrA may be required. Therefore, tagging with the wild-type SsrA peptide and
subsequent proteolysis of the tagged substrate(s) may be essential for this process (Keiler
and Shapiro, 2003). Alternatively, some dominant negative proteins involved in the cell
cycle control may not be inactivated by SsrA-DD tagging whereas Rep is.
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MATERIALS AND METHODS
Bacterial strains and plasmids
The wild-type C. crescentus strain used in this study is CB15N (Evinger and
Agabian, 1977). C. crescentus strains were grown at 30oC in M2G, M2X, or PYE
medium (Ely, 1991) supplemented with 1-2 µg/ml chloroamphenicol, 2.5-5 µg/ml
gentamycin, 5-20 µg/ml kanamycin, 25-50 µg/ml spectinomycin, or 1-2 µg/ml
tetracycline as necessary, and monitored by optical density at 660 nm. The ΔssrA and
∆smpB strains have been described previously (Keiler and Shapiro, 2003). Strain
CB15N::rep (∆ssrA::rep) in which rep is integrated into the xylX locus on the
chromosome was generated by transforming pRepb into wild-type C. crescentus (or
∆ssrA) and selecting recombinants by growth on kanamycin. Insertion of pRepb at the
correct locus was verified by PCR. Strain D850-1 in which wild-type ssrA gene was
replaced by a variant ssrA-His6 encoding a tag-peptide with six histidines at the C
terminus was constructed as follows. A DNA fragment containing the ssrA-His6 and
promoter sequences were digested out from pSsrA-His6 (Chapter 3) and cloned into p850
(pNPTS138 with DNA sequences flanking both ends of the ssrA gene) (Keiler and
Shapiro, 2003). The resulting plasmid was transformed into the ∆ssrA strain to generate
the D850-1 strain using the two-step recombination method as previously described (Gay
et al., 1985), and verified by PCR analysis. To construct a strain containing chromosomal
copies of both ssrA-His6 and rep, pRepc was introduced into the xylX locus of strain
D850-1 and integrants was selected by growth on kanamycin. The incorporation of
pRepc was verified by PCR. E. coli strains were grown at 37°C in Luria-Bertani broth
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(Sambrook et al., 1989) supplemented with 50-100 µg/ml ampicillin, 20-30 µg/ml
chloramphenicol, 15-20 µg/ml gentamycin, 30-50 µg/ml kanamycin, 50 µg/ml
spectinomycin or 12-12.5 µg/ml tetracycline as necessary, and monitored by optical
density at 600 nm. The wild-type E. coli strain used in this work was MG1655. Strain
MG1655 ssrA::kan was constructed by phage P1 transduction from strain W3110
ssrA::kan (Karzai et al., 1999). pRepa was constructed by amplifying the coding
sequence of the rep gene from plasmid pJS14 by PCR and cloning the product into the
plasmid pET28a (Novagen) to produce a gene encoding N-terminal His6-tagged Rep. To
generate pRepb, the rep gene was amplified by PCR from pJS14 and cloned into plasmid
pRW432 under control of the xylose-inducible promoter (Meisenzahl et al., 1997).
Plasmid pRepc was constructed by amplifying the coding region of rep by PCR from
pJS14 using primers to add a Strep-tag II (Trp-Ser-His-Pro-Gln-Phe-Glu-Lys) at the 5’
end of the rep coding sequence, and the product was cloned into plasmid pRW432.
A genetic screen to isolate plasmids that can by-pass the requirement for SsrA
Plasmid pKJS2 was electroporated into E. coli mutD strain. Transformants were
recovered on LB agar plate supplemented with 50 µg/ml kanamycin. Mutated pKJS2
plasmids were isolated from all the transformants, electroporated into ∆ssrA C.
crescentus strain, and plated on PYE agar plate supplemented with 20 µg/ml kanamycin.
Plasmids from kanamycin-resistant C. crescentus were prepared and retransformed into
∆ssrA C. crescentus strain. The mutations were identified by direct sequencing of the
recovered plasmid.
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Transformation of C. crescentus strains
To test the maintenance of various plasmids, 100 ng of each plasmid was used to
transform wild-type, ∆ssrA, and ∆smpB C. crescentus strains by electroporation or
conjugation. Transformants were recovered on PYE agar plates supplemented with
appropriate antibiotics at 30°C and counted.
Protein purification and antisera production
His6-Rep for antibody production was produced from log-phase cultures of E. coli
strain BL21(DE3)/pLys (Novagen) bearing plasmid pRepa by growth in the presence of 1
mM isopropyl-β-D-thiogalactopyranoside for 3 h. The culture was quickly cooled on ice,
cells were harvested by centrifugation, and the cell pellet was suspended in 5 ml buffer A
(10 mM Tris-HCl (pH 8.0), 100 mM NaH2PO4, 8 M urea, 5 mM imidazole) per gram wet
weight and lysed by sonication. The lysate was cleared by centrifugation at 10,000 x g for
30 min and the supernatant was added to 1 ml Ni2+-nitrilotriacetic acid (NTA) resin
(Qiagen) equilibrated in buffer A. After mixing for 1 h at room temperature, the resin was
packed into a column, washed with 100 ml buffer A, 100 ml buffer A20 (A buffer with
20 mM imidazole), and eluted with 10 ml buffer A containing 250 mM imidazole.
Fractions were separated on a preparative SDS-polyacrylamide gel and the band
corresponding to the His6-Rep protein was excised and used to immunize rabbits.
Immunization and sampling of the sera were performed by Josman LLC. (Napa, CA).
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Western analysis
For Western blot analysis of Rep protein expression, C. crescentus bearing
various plasmids were grown to log-phase in PYE broth and lysed in SDS sample buffer
(Sambrook et al., 1989). Log-phase cultures of strain CB15N, CB15N::rep, and
∆ssrA::rep were induced with 0.1% xylose, grown for 3 h in PYE broth and lysed in SDS
sample buffer (Sambrook et al., 1989). Equal volumes (corresponding to equal OD660/ml
of culture) of total cellular proteins were electrophoresed on 15% SDS-polyacrylamide
gels and transferred to PVDF membranes (Amersham Biosciences). These blots were
probed with anti-Rep antisera followed by goat anti-rabbit IgG, and detected with ECL
Western blotting detection reagents (Amersham Biosciences) and X-ray film.
Growth curve
Cultures of strain CB15N, CB15N::rep, ∆ssrA, and ∆ssrA::rep were grown in
PYE medium to an OD660 of 0.5-0.6. Cells were then washed in M2G or M2X and diluted
into fresh M2G or M2X at an OD660 of 0.05. Cultures were grown at 30°C to an OD660 of
0.5-0.6 and samples were taken periodically to monitor the changes in optical density at
660 nm. The absorbance values were fit to an exponential function for determination of
growth rates.
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Purification and characterization of SsrA tagged Rep
To purify the principle SsrA-His6 tagged Rep, log-phase cultures of C. crescentus
strain D850-2 were induced by the addition of xylose to 0.1%, and grown for 3 h at 30°C
in PYE broth. Cells were harvested by centrifugation, resuspended in 5 ml buffer B (50
mM NaH2PO4 (pH 8.0), 300 mM NaCl, 10 mM imidazole) per gram wet weight and
lysed by sonication. The lysate was cleared by centrifugation at 10,000 x g for 30 min
and added to 0.5 ml Ni2+-NTA resin equilibrated in buffer B. After mixing for 1 h at 4°C,
the resin was packed into a column, washed with 100 ml buffer B, 100 ml buffer B20 (A
buffer with 20 mM imidazole). SsrA-His6 tagged Rep was eluted with 10 ml buffer B
containing 250 mM imidazole and applied to a Strep-Tactin column (IBA) equilibrated in
buffer C (100 mM Tris-HCl (pH 8.0), 100 mM NaCl, 5 mM EDTA). The column was
washed with 100 ml buffer C and Strep-Rep-SsrA-His6 was eluted with 10 ml buffer C
containing 2.5 mM desthiobiotin. Purified tagged Rep was separated on a SDS-
polyacrylamide gel and identified by Coomassie Brilliant Blue staining. Gel slices were
excised and processed for peptide mass fingerprinting by matrix-assisted laser desorption
ionization time-of-flight (MALDI-TOF) mass spectrometry (Penn State Mass
Spectrometry Facility) as described (Chapter 3). A database containing Rep protein
sequence with the SsrA-His6 tag (AANDNFAEHHHHHH) after each amino acid was
created and searched using MS-Fit (Clauser et al., 1999) as described (Chapter 3). An
expected mass accuracy of 500 ppm was used. A maximum of two missed enzymatic
cleavage and modification of cysteines by carboxyamidomethylation were considered
during the searches. For protein identification, the requirement of successful matches
128
entailed the junction peptide and a minimum of five peptides mapping 50% of the intact
mass of a protein.
129
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Figure 4.1. Colony forming units of plasmids in C. crescentus strains. The values presented are the averages of three separate experiments, and error bars are included where applicable.
133
Figure 4.2. Schematic representation of a genetic selection for plasmids bypassing the requirement of SsrA activity. Mutated plasmid KJS2 (pKJS2) via replication in mutD mutant E. coli was transformed into ∆ssrA C. crescentus and selected by growth on kanamycin.
134
Figure 4.3. Properties of plasmid pPT1 and construction of variant pKJS2 plasmids. (A) pPT1 is generated by a 1.8 kb deletion (red line) of pKJS2 that removed the 3’ end of the open reading frame of the rep. (B) Schematic depiction of wild-type (in pKJS2) and truncated rep (in pPT1) is shown. The size of open reading frame (bp) and total number of amino acids (aa) are shown above and below the bar, respectively. The altered C-terminal amino acid sequences in truncated Rep (aa 155, and aa 158-160) are shown in red. (C) Schematic drawings (not to scale) of the rep (blue bar)-lacZα (yellow bar)-nptII (green bar) region of pKJS2, pPT1, and several pKJS2 variants. For each construct, mutated amino acid residues in Rep are shown in red and letter Z indicates the end of Rep protein sequence. Restriction enzymes, PshAI and NsiI are used to delete sequences from pKJS2 (panels 5-7).
135
Figure 4.4. MALDI-TOF analysis of SsrA-tagged Rep. (A) Experimentally obtained and expected monoisotopic peptide masses are shown. A search was conducted against the Rep database as described in Materials and Methods. The fragments matching the protein sequence are highlighted in bold (B). The Strep-tag-II and SsrA-His6-tag are singly and doubly underlined, respectively.
136
Figure 4.5. Overexpression of Rep does not harm cells. (A) Wild-type cells (lanes 1 and 2), wild-types cells containing rep integrated into the xylX locus on the chromosome (lanes 3 and 4), ∆ssrA cells containing rep integrated into the xylX locus on the chromosome (lanes 5 and 6) were cultured in minimal media with glucose (M2G which suppresses the xylX locus) or xylose (M2X which induces the xylX locus). Total proteins were prepared from these cells and analyzed by Western blotting using α-Rep antibody. The arrow indicates the full-length Rep. (B) The doubling time (min) during exponential growth of wild-type (or ∆ssrA) strain, and an isogenic strain containing rep integrated into the xylX locus on the chromosome was measured.
137
Figure 4.6. Expression of Rep in wild-type and ∆ssrA C. crescentus. Equivalent amounts of total proteins were prepared from wild-type cells harboring no plasmid (lane 1), pKK838 (lane 2), pKK839 (lane 3), pKJS2 (lane 4), and pPT1 (lane 5), or ∆ssrA cells harboring, pKK838 (lane 6), pKK839 (lane 7), and pPT1 (lane 8), were prepared and analyzed by Western blotting using α-Rep antibody. The solid arrow and the dashed arrow indicate the full-length Rep and the truncated Rep, respectively.
CURRICULUM VITAE
Sue-Jean Hong EDUCATION 1991-1996 Hon. B.Sc. Biochemistry and Chemistry University of Toronto Toronto, Canada 1997-1999 M.S Life Science Gwangju Institute of Science and Technology Gwangju, Korea 1999-2005 Ph.D. Biochemistry, Microbiology and Molecular Biology The Pennsylvania State University University Park, PA. AWARDS, HONORS AND FELLOWSHIPS 1991-95 Canada Scholarship University of Toronto Open Admission Scholarship University of Toronto Erindale College Admission Scholarship 1999 The Braddock Graduate Fellowship 2000 The Althouse Outstanding Teaching Assistant Award 2001 Paul Berg Prize in Molecular Biology 2002 Student Travel Award for Society for Developmental Biology 61st Annual
Meeting 2003 Braucher Fund Award 2004 R. Adams Dutcher travel fund Award PUBLICATIONS 2003 Abmayr, S. M., Balagopalan L., Galletta, B. J. G., and Hong, S-J. 2003. Cell and
Molecular Biology of Myoblast Fusion. Int. Rev. Cytol. 225: 33-90. 2005 Hong, S-J., Tran, Q. A., and Keiler, K. C. 2005. Cell-cycle regulated degradation
of tmRNA is controlled by RNase R and SmpB. Mol Microbiol. 57: 565-575. ORAL PRESENTATIONS AT INTERNATIONAL CONFERENCES 2003 Hong, S-J., Tran, Q-A., and Keiler, K.C. Regulation of SsrA RNA in
Caulobacter crescentus. The 2003 Molecular Genetics of Bacteria and Phages Meeting. Madison, WI. August 05-10, 2003.
