SciP: A novel inhibitor of CtrA transcriptional activity in
Caulobacter crescentus
N
MASSACHUSE
I SEPO
by
i
201
Kasia G. Gora
LIBRAR'ES
B.S. Biology
California Institute of Technology, 2006
ARCHNVES
SUBMITTED TO THE DEPARTMENT OF BIOLOGY IN PARTIAL FULLFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY IN BIOLOGY
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MASSACHUSETTS INSTITUTE OF TECHNOLOGY
August 8, 2011
© Kasia G. Gora. All rights reserved.
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SciP: A novel inhibitor of CtrA transcriptional activity in
Cauobacter crescentus
by
Kasia Gora
Submitted to the Department of Biology
on August 8, 2011 in partial fulfillment of the requirements of the degree of
Doctor of Philosophy in Biology at the Massachusetts Institute of Technology
Abstract
Cells must sense changes in their environment and respond appropriately in order to
survive. A common survival strategy is for cells to translate an environmental signal into
the activity of a transcription factor they effects a change in gene expression. In this way,
cells can express a gene just-in-time for its biological function. In addition, cells can
coordinate the activity of many transcription factors to construct genetic regulatory
networks that integrate many inputs to control complex cellular behaviors.
In this thesis, I use the model system Caulobactercrescentus to examine the regulation of
CtrA, the essential transcription factor at the core of the Caulobactercell cycle regulatory
network. CtrA regulates the activity of approximately 100 cell cycle genes many of
which are critical for cell cycle progression. CtrA activity is regulated at the level of
abundance, post-transcriptional modification, and here I show that CtrA is also regulated
by a novel protein-protein interaction. I identify SciP, a GI specific inhibitor of CtrA
transcriptional activity and show that SciP forms a complex with CtrA at CtrA dependent
promoters. The SciP/CtrA interaction likely prevents CtrA from recruiting RNA
polymerase thereby blocking the activation of transcription. In addition, I show that SciP
is restricted to G1 by regulated proteolysis. I identify the Lon as the SciP protease and
show that Lon is required for SciP proteolysis in vivo and that E. coli Lon degrades SciP
in vitro. Finally I engineer a stable allele of sciP and show SciP proteolysis is critical for
proper cell cycle progression.
Thesis Advisor: Michael T. Laub
Title: Associate Professor of Biology
Acknowledgements
I would like to thank Mike Laub for taking me on as his graduate student and giving me the
opportunity to work in his lab. Mike has been a wonderful mentor. He is wicked rigorous and
wicked smart. Thank you for investing so much time in developing my project and
sharpening my communication skills. I hope that I have risen to meet your high expectations.
I would also like to acknowledge Barrett Perchuk for his invaluable scientific and moral
support. Without his technical expertise I could not have been so productive. Barrett is also
my dear friend. Barrett, you made every day in lab a pleasure, even that bad ones. Thank you
for your unwavering friendship and support.
It has been a privilege to work with all the Laub lab members past and present. I would
especially like to acknowledge Christos Tsokos, Erin Chen, and Andy Yuan for helping me
with key experiments and Jeff Skerker for his contributions to building the infrastructure of
the lab. I would like to thank Emily Capra, Andy Fan, Erin Chen, Josh Modell, and Celeste
Peterson for all their invaluable help editing this manuscript. Emily and Andy Fan went
above and beyond to help me edit the introduction. Emily is my two-component fairy
godmother and Andy is the yeast master. In addition Emma Lubin, Kristina Jonas, Chris
Aakre, Anna Podgornaia and Diane Baer have all contributed to making the Laub lab a
wonderful place to work. It has been a true pleasure to work with our honorary lab member
Orr Ashenberg and his fellow Keating lab member Aaron Renkie. I would like to thank Matt
Wohlever for his enthusiastic work performing in vitro degradation.
I would also like to thank the members of my thesis committee Angelika Amon and Alan
Grossman for years of thoughtful comments, helpful advice, and advocacy. Thank you Eric
Alm for being my outside examiner and a generally cool dude. In addition, Frank Solomon
has also been an incredible mentor in science, career, and life. Thank you Tania Baker, Bob
Sauer, Amy Keating, and Steve Bell (and Radar) for taking the time out of your busy
schedules to chat. Finally, a special thank you to Tony Sinskey for putting in a good word
and helping me find my first job.
I would like to thank all my friends who have supported me over the years. Thank you
Jennifer Patton, Alex McCauley, Marya Shiekh, Paul Wiggins, Hila Hashemi and the entire
CMT crew, Michael Betancourt, Matt Walker, Richard Eager, Cindy Chen, Eric Spear, Mary
Lee, Lori Ling, Bernardo Pando, Drew Nager, Mihir Mehta, Emilia Bora, Mihai Bora, John
Allman, Jared Leadbetter, Eric Matson, Alden Waters, Kaveh Milaninia, Paola Rubesco, Jane
& Marty, Zenzie Brooks, and especially Matthieu Couturier and our fluff-nugget Darwin.
Thank you to the cheerful staff of Starbucks for fueling my Ph.D.
Most of all I'd like to thank my family for their unconditional love and support that have
made possible this amazing opportunity. Mom, Dad, and Michael, thank you for holding my
hand through the last few months, I could not have finished without you. Thank you Oscar,
Poosha, and Peanut for all your love and cuteness.
Table of Contents
Ab stract ................................................................................................................................
2
Acknowledgements.......................................................................................................
3
Chapter 1: Introduction...................................................................................................
9
Regulation of transcription ................................................................................
10
Transcriptionand the yeast cell cycle ...................................................
15
Two-component systems........................................................................
18
Regulating two-component systems ....................................................
23
Elaboration of two-component systems in the Caulobactercell cycle..............26
Caulobacteras a cell cycle model ........................................................
27
CtrA, master regulatorof the Caulobactercell cycle............................
30
Regulation of CtrA ................................................................................
35
Identifying missing regulatorsof CtrA ................................................
39
References ........................................................................................................
Chapter 2: A cell-type-specific protein-protein interaction modulates transcriptional
activity of a master regulator in Caulobactercrescentus ............................................
41
46
Abstract .................................................................................................................
47
Introduction ......................................................................................................
47
1
Resu lts....................................................................................................................5
Identification of sciPas a candidate regulatorof CtrA and the cell cycle51
SciP is present only in GJ swarmer cells...............................................
54
sciP is requiredfor proper swarmer cell development and cell cycle
progression...........................................................................................
55
Depletion of SciP affects CtrA -dependentgene expression...................58
Overexpressing sciP disrupts the cell cycle and drives the downregulationof CtrA -dependent genes .....................................................
59
SciP does not affect CtrA degradationorphosphorylation...................63
SciP binds directly to CtrA ..................................................................
65
SciP binds CtrA without disruptingDNA-binding................................
67
Discussion .......................................................................................................
70
Regulation of SciP.................................................................................
72
M echanism of inhibition CtrA ...................................................................
73
Regulating response regulators............................................................
74
Concludingremarks ..............................................................................
75
Experimental procedures ..................................................................................
76
Acknowledgements............................................................................................
87
References..............................................................................................................88
Chapter 3: The Caulobactercell cycle regulator SciP is proteolyzed at the G1 -to-S
transition by Lon ................................................................................................................
91
Introduction............................................................................................................92
2
R esu lts....................................................................................................................9
SciP is regulatedprimarily at a post-transcriptionallevel ...................
92
SciP is an unstable protein, but is not degradedby ClpXP, ClpAP, or
HslUV ....................................................................................................
93
SciP is degradedby Lon .......................................................................
96
Addition of a C-terminal tag interferes with SciPproteolysis...............97
Proteolysis of SciP is necessaryfor proper cell cycle progression........... 99
Discussion............................................................................................................102
Post-translationalRegulation of SciP .....................................................
102
Regulated Proteolysis by Lon ..................................................................
102
ConcludingRemarks................................................................................104
Experim enal Procedure........................................................................................105
Acknowledgem ents..............................................................................................109
References............................................................................................................110
Chapter 4: Conclusions and future work .........................................................................
112
Characterizing the interaction between CtrA and SciP........................................114
Does SciP contactDNA and if so, how? ..................................................
114
Why does SciP requirefree DNA for complex formation?...................... 117
What is the molecular surface that mediates the interactionbetween CtrA
118
andSciP ...................................................................................................
Regulated proteolysis and the Caulobactercell cycle.........................................119
Outstanding questions in cell cycle regulated transcription in Caulobacter....... 121
List of Figures and Tables
Chapter 1: Introduction
Figure 1.1 Three modes of activation of gene expression by transcription factors 1
Figure 1.2 Activation of eukaryotic transcription.............................................
13
Figure 1.3 Regulation of metazoan transcription factor E2F.............................14
Figure 1.4 S. cerevisiaecell cycle.....................................................................
16
Figure 1.5 S. cerevisiae cell cycle transcription network .................................
17
Figure 1.6 Two-component architecture..........................................................
22
Figure 1.7 Connector proteins in two-component systems...............................25
Figure 1.8 Caulobactercell cycle.....................................................................
28
Figure 1.9 Flagellar assembly............................................................................
29
Figure 1.10 CtrA activity and regulation through the cell cycle.......................30
Figure 1.11 Dual function of CtrA in regulation of the origin and cell-cycle
regulated transcription .......................................................................................
32
Figure 1.12 PhoB activation by phosphorylation ............................................
34
Figure 1.13 Activation of transcription by CtrA...............................................35
Figure 1.14 Regulation of CtrA transcription ...................................................
36
Figure 1.15 Differential activation of CtrA at the swarmer and stalked pole........37
Figure 1.16 CtrA proteolysis by ClpXP ...........................................................
38
Chapter 2: A cell-type-specific protein-protein interaction modulates transcriptional
activity of a master regulator in Caulobactercrescentus
Figure 2.1 Identification of sciP as a key regulator of cell cycle progression.......52
Figure 2.2 Multiple sequence alignment of SciP orthologs...............................53
Figure 2.3 Flow cytometry analysis of cells from Figure 2.1E .........................
55
Figure 2.4 SciP depletion phenotype ................................................................
57
Figure 2.5 Effects of sciP overexpression and depletion on CtrA-dependent gene
. 61
expression ........................................................................................................
Figure 2.6 Phenotypic consequences of sciP overexpression............................62
Figure 2.7 Morphology and chromosomal content offtsZ depletion strain...........65
Figure 2.8 SciP and CtrA interact directly in a yeast two-hybrid assay ............
66
Figure 2.9 Model of SciP structure ...................................................................
67
Figure 2.10 Electrophoretic mobility shift assays with CtrA and SciP .............
69
Figure 2.11 Regulation of CtrA-dependent gene expression during cell cycle
p rogression .............................................................................................................
71
T able 2.1 Strains list .........................................................................................
85
Table 2.2 Prim ers list .........................................................................................
86
Chapter 3: The Caulobactercell cycle regulator SciP is proteolyzed at the GI -to-S
transition by Lon
Figure 3.1 Post-transcriptional regulation of SciP............................................
93
Figure 3.2 SciP stability in various protease mutant strains ..............................
95
Figure 3.3 In vitro proteolysis of SciP by E. coli Lon .....................................
97
Figure 3.4 Stabilization of SciP through an C-terminal M2 tag ........................
98
Figure 3.5 Phenotypic consequences of stable SciP ............................................
100
T able 2.1 Strains list ............................................................................................
108
T able 2.2 Prim ers list ...........................................................................................
108
Chapter 4: Conclusions and future work
Figure 4.1. Dispensability of the putative SciP motif in vitro and in vivo...........116
Figure 4.2. Modeling the three-dimensional structure of SciP ............................ 118
Figure 4.3. Electrophoretic mobility assays with mutant CtrA and SciP ............ 119
Chapter 1
Introduction
Introduction
Regulation of transcription in cells
Signal transduction is fundamental to the lives of all cells. In order to sense
environmental stimuli or to internally regulate cellular events, a cell must properly
translate an internal or external signal into an effect. One of the most common effects, or
outputs, of signaling pathways is a change in activity of a transcription factor and,
consequently, a change in gene expression. In this way, cells adaptively regulate the
expression of genes just-in-time for their biological function.
How do transcription factors activate gene expression? In bacteria, genes are transcribed
by a five subunit RNA polymerase complex (a2pp'o) in association with a sigma factor
[1]. The sigma factor typically recognizes DNA sequences near the -35 and -10 positions
relative to the transcriptional start site allowing polymerase to bind to the double stranded
DNA and form a closed-complex [1, 2]. Next the complex undergoes a conformational
change, melting the DNA strands to form an open-complex that can initiate transcription
[2].
Transcription factors can activate transcription in at least three general ways: regulated
recruitment of RNA polymerase, activation of RNA polymerase, and direct promoter
activation [2]. A classic example of the regulated recruitment of RNA polymerase by a
transcription factor comes from studies of the lac operon of E. coli. The lac operon
consists of several genes involved in lactose metabolism including lacZ, a gene encoding
p-galactosidase, an enzyme that cleaves lactose [2]. Although E. coli can metabolize
lactose, glucose is the preferred energy source and the lac operon has evolved to highly
express lactose metabolism genes when glucose is unavailable and lactose is present [3].
The lac operon is normally repressed by the binding of the lac repressor, Lac, to the -10
region of the lac promoter thereby occluding RNA polymerase and preventing
transcription (Fig. 1.1A) [4]. Allolactose, a lactose metabolite, can bind LacI, thereby
relieving repression of the lac operon. In the absence of glucose, the transcription factor
CAP (catabolite activator protein) is binds the signaling molecule cAMP activating CAP
to bind the lac promoter and recruit RNA polymerase that is bound to 070 through a direct
interaction with the a subunit of RNA polymerase (Fig. 1. lA) [5]. In this way E. coli can
induce lactose metabolism genes just when they are needed.
allolactose
.3t
~~~~0~
lacZ
W44IacZ
.35
inerT
Figure 1.1. Three modes of activation of gene expression by transcription factors. (A)
Regulated recruitment of RNAP by CAP. (B) NtrC activation of RNAP. (C) MerT activation of the
mer operon promoter. Adapted from [2].
In contrast to the lac operon, the gin operon of E. coli requires polymerase activation
rather than recruitment [2]. The gin operon encodes three genes important for nitrogen
metabolism including ginA, a gene that encodes glutamine synthetase, an enzyme
involved in glutamine synthesis [6]. The amino acid glutamine is an important
component of proteins and precursor for the synthesis of other nitrogen containing
molecules by the cell [6]. Expression of the gin operon is activated by NtrC, a
transcription factor that binds more than 100 nucleotides upstream of the promoter [6].
NtrC binds DNA in response to nitrogen starvation and the NtrC/DNA complex loops
around to make contact with the closed complex of RNA polymerase bound to the c54
leading to isomerization from the closed to open complex (Fig. 1.1.B) [6]. In this way
NtrC activates the gin operon to express glutamine synthetase and make glutamine when
the cell is starved of nitrogen.
As a final example, the mer operon requires activation of the promoter [2]. In many
Gram-negative bacteria, the transposon encoded mer operon (merTPAD) confers
resistance to the toxic heavy metal mercury [7]. Expression of the mer operon is
regulated by the transcription factor MerR [7]. In the absence of mercury, MerR forms an
inactive closed-complex with RNA polymerase at the mer operon promoter, but when
mercury is present, it binds MerR, causing a conformational change in the protein that
allows it to distort the DNA and thereby activate transcription (Fig 1.1.C) [8]. This
mechanism allows MerR to turn on mer operon only in the presence of mercury.
In many ways, transcriptional activation in eukaryotes is an elaboration of prokaryotic
mechanisms. Unlike prokaryotes, eukaryotes have several different RNA polymerase
enzymes but the majority of genes are transcribed by Poll which is structurally similar to
prokaryotic RNA polymerase [5]. Eukaryotic Poll promoters are often characterized by a
TATA box, an AT-rich region 50-70 base pairs upstream of the transcriptional start site
which plays an analogous role to the prokaryotic -10 and -35 boxes [5]. Unlike bacteria,
yeast require many global transcription factors including TBP and TFIIB to activate
transcription [5]. TBP (TATA binding protein) plays the critical role of recognizing and
binding the TATA box and in association with a host of associated factors forms the
TFIID complex. In a gross simplification, TFIID recruits the holoenzyme consisting of
PolIl and many other factors through TFIIB, a factor that acts as a protein bridge between
TBP and the holoenzyme complex (Fig. 1.2) [9]. In yeast, transcription factors generally
have two separable domains, a DNA binding domain and an activating domain that
together can activate transcription through regulated recruitment of PolII [5]. The
activating domain interacts with the holoenzyme and components of TFIID while the
DNA binding domain binds between 100 and 250 base pairs upstream of the
transcriptional start site (Fig. 1.2) [5].
TATA
Figure 1.2. Activation of eukaryotic transcription. The TFlD complex recognizes the TATA
box and binds the RNA polymerase holoenzyme through TFIlB. Eukaryotic transcription factors
engage in regulated recruitment of the holoenzyme by interacting with DNA through a DNA
binding domain (BD) and the holoenzyme through the activating domain (AD). Adapted from [5].
Transcription factors play a key role in regulating gene expression so it is no surprise that
their activity is strictly regulated. For instance the metazoan transcription factor E2F
drives the expression of hundreds of genes in late G1, many of which are critical for
proper progression of S phase [10]. E2F activity is regulated in at least three ways:
abundance, protein-protein interactions, and post-transcriptional regulation (Fig. 1.3).
First E2F abundance is regulated at the transcriptional level with mRNA accumulating in
GI [11]. Secondly, in GO, E2F is prevented from binding DNA and activating
transcription by a protein-protein interaction with the Rb protein [10]. During G1, Rb
becomes phosphorylated and releases E2F initiating the late G1 transcriptional program
[10]. Finally in S phase, E2F itself becomes phosphorylated which reduces its ability to
bind promoters and ensures that the E2F regulon is only expressed during a brief window
in late GI [12].
GO
GI
Figure 1.3. Regulation of metazoan transcription factor E2F. In GO, E2F is inactivated by a
protein-protein interaction with Rb and is released by Rb phosphorylation in G1 at the same time
E2F mRNA accumulates. E2F is phosphorylated and inactivated in S phase.
Transcription and the yeast cell cycle
Our previous examples have focused on the regulation of individual genes or operons by
individual transcription factors. However, many transcription factors can interact to form
genetic regulatory networks that control complex processes.
For instance, a complex
transcription factor network controls cell cycle gene expression in yeast ensuring that
these genes are expressed at their time of function [13]. This genetic regulatory network
interacts with a biochemical oscillator to drive progression of the cell cycle. Because the
biochemical oscillator was characterized before the genetic regulatory network, I will
begin with a brief overview of its key players and then discuss the role of regulated
transcription in the cell cycle.
In eukaryotes, cyclin-dependent kinases (Cdks) are at the core of the biochemical cell
cycle oscillator. While metazoans have several Cdks, the budding yeast Saccharomyces
cerevisiae has only one, Cdc28, which phosphorylates different substrates based on its
association with 9 different cyclins: Cln3 interacts with Cdc28 in GI, Clnl and Cln2 at
the G1-to-S transition, Clb5 and Clb6 in S phase, and Clbl, Clb2, Clb3, and Clb4 in M
phase (Fig 1.4) [14,
15]. For instance in S phase, the Cdc28/Clb5 complex
phosphorylates key proteins that activate origins of replication to permit the initiation of
DNA synthesis [16]. During M phase, another complex, Cdc28/Clb2, phosphorylates
Pdsl, protecting it from degradation [17]. Pdsl is an inhibitor of Espl, a chromosome
seperase protein. Therefore Cdc28/Clb2 activity prevents premature chromosome
seperation by stabilizing Pds 1 [18]. These are just a few of the many examples of key cell
cycle processes that are regulated by Cdc28 activity.
Clbi,2
3,4
G2
Cdc28
cn
Clb5,6
So
Cini,2
S
assoiatin wih vDNArosinoteisTe
replication
elld ylrtae
r clrcdeensherlvn
cycinsareindcae ngretpanel [1,1]
bud fo
Start
Figure 1.4. S. cerevisiae cell cycle. Key events of the yeast cell cycle are regulated by Cdc28 in
association with various cyclin proteins. The cell cycle stages are color-coded and the relevant
cyclins are indicated in grey panels [14, 15].
In addition to the biochemical oscillator, a group of transcription factors ensures that cell
cycle genes are expressed at the appropriate time. The key players include Mbp 1, Swi4,
Swi6, Mcml, Fkhl, Fkh2, Nddl, Swi5, and Ace2 (Fig. 1.5) [13]. In late GI the SBF
complex (Swil and Swi6) and the MBF complex (Mbpl and Swi6) drive the expression
of 200 genes, the so-called Cln cluster [19]. SBF controls the expression of genes
involved in budding while MBF regulates genes involved in DNA replication and repair
[19]. Next, in G2 phase, Fkhl and Fkh2 activate transcription of the Clb2 cluster, -35
that are involved in mitosis [20]. Activation of Fkh2 requires the phosphorylated
transactivator Nddl [21]. In M phase, Mcml drives the expression of the MCM cluster of
genes involved in DNA synthesis [20]. Finally in late M phase and G1, Swi5 and Ace2
control expression of the Sici cluster [22, 23].
These transcription factors form a regulatory network in which transcription factors
expressed in one stage of the cell cycle activate those expressed in the next stage [13].
For instance, in late GI, SBF and MBF both regulate expression of Nddl, the key
activator of G2 expression by Fkh2 [13]. Then in G2, Fkh2 and Nddl regulate the
expression of Swi5 and Ace2 which act in M phase [13]. Finally, M phase Mbpl
activates the expression of Swi4 that acts in late GI closing the regulatory loop (Figl.5)
[13].
Swi5
Ace2
Mcm1
Fkh2
Nddi
Swl6
G2
S
Swl6
Sw14
Mbp1
start
Figure 1.5. S. cerevisiae cell cycle transcription network. Key transcription factors are
indicated in grey ellipses at their time of activity. General interactions between phase specific
clusters are indicated by arrows. Adapted from [13].
The biochemical oscillator and transcription network driving cell cycle progression in
yeast have many points of interaction. For instance Swi5 and Ace2 regulate the Cln3
promoter [13]. In turn both Swi4 and Swi6 are targets of Cdc28/Cln3 in vitro and Swi6
has been shown to be phosphorylated in a Cdc28/Cln3 dependent manner in vivo [22, 24-
17
26]. In addition, SBF is inhibited by association with the protein Whi5 until Whi5
becomes phosphorylated by Cdc28/Cln3 and releases SBF to activate transcription [27].
SBF and MBF also bind the Clnl,2 and Clb5,6 promoters [13]. Furthermore the
transactivator Nddl is a substrate of Cdc28/Clb2 creating a positive feedback loop that
ensures Clb2 expression occurs only late in S phase [28]. Cdc28 activity clearly
influences cell cycle gene expression; however, recent work shows that the majority of
cell cycle regulated transcription remains periodic in a strain lacking all S and M phase
cyclins suggesting that the transcription factor network is autonomous of the biochemical
oscillator [29].
Two-component systems
A common way of modulating gene expression in bacteria involves two-component
signal transduction systems. These signaling pathways allow cells to sense and respond to
their internal or external environment. At their core, two-component systems consist of a
histidine kinase and a response regulator. In response to a signal, the kinase typically
autophosphorylates a conserved histidine residue and then specifically transfers the
phosphoryl group to a conserved aspartate residue on the response regulator. Many
response regulators contain DNA binding domains, and thus phosphorylation can lead to
downstream transcription by activating DNA binding ability [30]. In addition to
phosphorylating response regulators, many histidine kinases are bifunctional and can act
as phosphatases, dephosphorylating their cognate regulator when not activated to
autophosphorylate [30].
Histidine kinases consist of a modular kinase core coupled to a sensor domain. Sensor
domains vary greatly but a common topology is a periplasmic sensor consisting of two
transmembrane helices [31]. Other kinases are soluble and contain PAS or GAF domains
that allow them to bind and sense small molecules [31]. In contrast to the sensor domain,
the catalytic core contains a highly conserved C-terminal catalytic and ATP binding (CA)
domain and a slightly more variable dimerization and histidine phosphotransfer (DHp)
domain [31]. The CA domain catalyzes the transfer of the phosphoryl group from ATP to
a conserved histidine residue located in the DHp domain [31]. As the name suggests, the
DHp domain also mediates dimerization, which is essential for autophosphorylation,
although the autophosphorylation reaction can occur either in cis or trans [30]. Unlike
many eukaryotic kinases, histidine kinase dimerization is not induced by the signal;
histidine kinases usually exist as dimers even when not active as kinases [31]. Histidine
kinases are modular and functional chimeras can be created by swapping sensor domains
and kinase cores of different proteins [32].
Response regulators are also modular and consist of a receiver domain and an effector
domain [31]. Receiver domains are conserved and contain the active site that receives the
phosphoryl group from the kinase on a conserved aspartate residue. Effector domains are
more variable but most encode DNA binding domains [31]. In these transcription factor
response regulators, phosphorylation of the receiver domain can promote protein-protein
interactions, like dimerization, that activate the response regulator to bind DNA and
change gene expression [30].
One of the best-studied two-component systems is the EnvZ-OmpR system that controls
osmoregulation in E. coli (Fig. 1.6A). In response to osmotic shock, the histidine kinase
EnvZ
is activated
and phosphorylates
its cognate
response regulator OmpR.
Phosphorylation of OmpR induces dimerization allowing OmpR to bind DNA and act as
a transcription factor. OmpR regulates the expression of two porin genes, ompF and
ompC, whose differential expression allows the cell to balance ions between the
cytoplasm and periplasmic space and thus protect the inner membrane from excessive
turgor pressure [31, 33].
An elaboration of the canonical two-component pathway is the phosphorelay. In a
phosphorelay, the histidine kinase transfers a phosphoryl group to a response regulator,
which in turn transfers the phosphoryl group to a histidine phosphotransferase (HPt),
which transfers it to another response regulator [31]. The addition of extra steps in a
phosphorelay has been speculated to allow for more complex regulation of the output. A
classic example of a phosphorelay controls the initiation of sporulation in B. subtilis (Fig
1.6B). In response to physiological cues, KinA and several other kinases phosphorylate
an aspartate residue on the response regulator SpoOF. SpoOF then phosphorylates a
histidine residue on the phosphotransferase SpoOB. In turn, SpoOB phosphorylates the
terminal response regulator SpoQA which increases its affinity for DNA and hence its
ability to activate target genes [34]. SpoOA directly regulates the expression of over 100
genes involved in initiating the sporulation genetic regulatory network [35].
The intermolecular connectivity between components of a phosphorelay can vary. In the
sporulation example, each component comprises its own protein. However different
components are often combined within a single protein. Generally it is the kinase that is
fused to the first receiver domain to form a so-called hybrid kinase [30]; the histidine
phosphotransferase can also be fused to a hybrid kinase. For instance the ArcB-ArcA
system of E. coli contains the same number of elements as the B. subtilis sporulation
phosphorelay, but the first three are fused into a single hybrid kinase, ArcB, with the
terminal regulator, ArcA, a separate, soluble protein (Fig. 1.6C). In response to anoxia,
ArcB autophosphorylates on the conserved histidine in the catalytic core and relays
phosphate to an aspartate on a receiver like domain, then to a histidine on a HPt like
domain, and finally to an aspartate on the response regulator ArcA. ArcA in turn
represses genes involved in aerobic respiration which would be energetically wasteful
under anaerobic conditions [36].
A. E. coi Osmoregulation
P
mechanical stress
froms osmotic
shock?
P
P
regulation of porin genes
ompF, ompC
OmpR
EnvZ
B. B. subtilis Sporulation
P
His
P
P
KinA
SpoOF
SpoOB
* P
~
SpoOA
Induction of the
sporulation
genetic regualtory
network
KinB-D
B. E. coil Anoxic Response
redox state of
eclector carriers
or membrane
potential?
6
ArcB
j P
P
P
represison of aerobic
metabolism genes
"
ArcA
Figure 1.6. Two-component architecture. (A)The EnvZ/OmpR two-component system from E.
coli. Phosphate flow from the kinase core (yellow bullet) to the receiver domain (blue box) is
indicated by the arrow. The input and output domains are represented by the gray and purple
arrows respectively. (B) The SpoQA phosphorelay from B. subtilis schematized as in (A) with the
addition of a receiver domain (teal box) and an HPt (orange ellipse). (C) The ArcB hybrid kinase
from E. coli. Adapted from [30, 37].
Given the utility and flexibility of two-component systems, it comes as no surprise that
most bacterial genomes encode dozens to hundreds kinase-regulator pairs that allow cells
to respond to diverse environmental signals [32]. Unlike eukaryotic kinases that often
phosphorylate many target proteins, histidine kinases are generally specific for their
cognate response regulators and preferentially phosphorylate them over other possible
targets [32]. How then do cells ensure the specificity of so many paralogous signaling
proteins? The answer lies in molecular recognition between the kinase and regulator
rather than contextual strategies such as scaffolds. Histidine kinases typically have a
strong kinetic preference for their cognate regulator over all other regulators in a cell
[38]. Recent studies have identified a small set of residues in the DHp domain of E. coli
EnvZ that confer specificity for its cognate regulator OmpR. Mutating three residues in
the wild-type E. coli EnvZ to those found in RstB was sufficient to change the specificity
of EnvZ from OmpR to RstA, the cognate regulator of RstB [39]. Subsequent work
demonstrated that mutating just four residues in E. coli OmpR to the RstA sequence was
sufficient to change specificity for EnvZ to RstB [40]. Thus, histidine kinases and
response regulators can ensure the specificity of partner recognition through a few key
specificity residues.
Regulating two-component systems
Two-component signaling pathways transduce information by converting various input
signals into the output of a response regulator's phosphorylation level. As described in the
previous section, activation of a histidine kinase by its ligand or signal results in
autophosphorylation and then phosphotransfer to its cognate regulator. However,
additional factors can influence the flow of phosphate and therefore the output of the
system. For example, protein phosphatases exist that dephosphorylate some response
regulators. Other factors can modify the output of a response regulator without affecting
their phosphorylation status, often by binding directly to a response regulator and
preventing its interaction with DNA. Collectively, factors that modulate phosphate flow
through a two component system are called connectors because they can provide links
between different regulatory systems [41]. Connectors can act at any step of the
phosphorelay; below I will discuss a few key examples for connectors whose mechanism
of action has been elucidated.
Some connectors act at the first step of the phosphorelay by regulating the
autophosphorylation of a histidine kinase. One such example is the B. subtilis protein
Sda, which blocks autophosphorylation of KinA (Fig. 1.7A) [42]. Sda is thought to bind
directly to KinA and prevent the CA domain from passing the phosphoryl group to the
conserved histidine in the DHp domain. [43]. The sda gene is activated by the replication
initiation/transcription factor DnaA; Sda protein is highly expressed when DnaA activity
is high as may occur when during the initiation of replication at the beginning of each cell
cycle [44]. Therefore expression of Sda prevents downstream phosphorylation of SpoOA,
preventing the initiation of the sporulation in cells that have not finished replicating their
DNA [44].
Other connectors act as phosphatases and directly dephosphorylate response regulators.
Several examples of phosphatases exist in the B. subtilis sporulation phosphorelay. For
instance members of the Rap family of phosphatases (RapA, RapB, RapE, and RapH) all
directly
dephosphorylate
the
response
regulator
SpoOF
(Fig.
1.7A)
[45-47].
Dephosphorylation of SpoOF results in back transfer of phosphate from SpoOA to SpoOB
back onto SpoOF, effectively preventing sporulation by dephosphorylating SpoOA [45].
The expression of some Rap phosphatase genes is induced during competence, an
alternative differentiation pathway where the bacteria can take up foreign DNA, and
others are induced during vegetative growth [45-47]. Thus the Rap phosphatases allow B.
subtilis to prevent inappropriate sporulation when the cells are growing vegetatively or
initiating competence. The SpoOE family of phosphatases, including SpoOE, YisI, and
YnzD, directly dephosphorylate SpoOA and are also differentially regulated by
alternative differentiation and growth pathways (Fig 1.7A) [48, 49].
RapA/BIHIE
SpoOE/YisIlYnzD
Sdax p
ilm
SpoOF
SpoOB
SpoOA
KinB-D
SixA
V!
P
P
ArcB
ArcA
A
\w RapH
ComA
Spx
R
\Y
!I
spx
ComA
Figure 1.7. Connector proteins in two-component systems. (A) Inhibition of KinA
autophosphorylation by Sda (red pentagon). Dephosphorylation of the receiver domains of SpoOF
and SpoQA by the Rap and SpoOE family of phosphatases (green crescent) respectively in B.
subtilis. (B) Dephosphorylation of the HPt of ArcB by SixA in E. coli. (C) Inhibition of ComA DNA
binding by RapH in B. subtilis. (D) Inhibition of transcriptional activation by ComA by the RNA
binding protein Spx (red half circle) in B. subtilis [30, 37].
Another target for phosphatases may be histidine kinases and phosphotransferases
although to date there has only been one report of a phosphatase for these types of
molecules. The E. coli protein SixA was suggested to dephosphorylate the histidine
phosphotransferase domain of the hybrid kinase ArcB, although the in vivo relevance of
SixA remains unclear (Fig. 1.7B) [50].
In addition to affecting the flow of phosphate in two-component systems, connectors can
also modulate the output of response regulators in other ways. In the case of response
regulators that are transcription factors, connectors can inhibit DNA-binding without
affecting phosphorylation. For example, although RapH acts as a phosphatase for SpoOF,
it has a different mechanism for regulating the ComP-ComA two-component system
which controls genetic competence in response to cell density signals [51].
RapH
directly binds ComA to prevent it from binding to target promoters (Fig. 1.7C) [46].
While RapH acts directly on ComA, another B. subtilis protein, Spx, blocks the activity
of ComA by interfering with its ability to recruit RNA polymerase (Fig. 1.7D). Spx binds
to the c-subunit of RNA polymerase, preventing the formation of a ComA-RNAPpromoter complex [52].
While these examples are by no means exhaustive, they give a flavor of the diverse
mechanisms that have evolved to modulate the activity of two-component systems.
Elaboration of two-component systems in the Caulobacter cell cycle
In the examples above I have focused on two-component signaling pathways that regulate
environmental and stress responses. However, these signaling proteins also play a critical
role in regulating the cell cycle and the development of Caulobactercrescentus, the focus
of my thesis work.
Caulobacter as a cell cycle model
Caulobactercrescentus is a fresh water oligotrophic alpha-proteobacterium. Members of
the Caulobacter genus were first described at the turn of the
20 'b century
and were
characterized by their long stalks and tendency to form clumps of cells called rosettes
[53]. The lab strain Caulobacter crescentus (CB15) was not isolated until 1960 [52].
However, because of its unique cell cycle, Caulobactercrescentus quickly emerged as a
system for studying differentiation and the bacterial cell cycle [53].
The Caulobacter cell cycle is characterized by an asymmetric cell division and cellular
differentiation. Each cell division yields a distinct mother cell (stalked cell) and daughter
cell (swarmer cell) (Fig. 1.8) [54]. The stalked cell is sessile and marked by a polar stalk
while the swarmer is motile and marked by a single polar flagellum and several pili [54].
Stalked cells are competent to immediately initiate DNA replication while swarmer cells
are in a G 1-like phase and must differentiate into stalked cells to initiate DNA replication
and proceed through the cell cycle. Caulobacter has proven to be an attractive model
system for understanding the bacterial cell cycle and cellular differentiation as each cell
cycle transition is coordinated with polar morphogenetic events [54]. The G1-to-S
transition is marked by the loss of the flagellum and pili and the development of a stalk at
the same pole, while the end of S phase is marked by the synthesis of a new flagellum at
the nascent swarmer pole opposite the stalked pole [54].
swarmer cell
late predvisional
N_=
-
stalked cell
S
chromosome
segregation
DNA replicati n
flagellar synthesis
pili synthesis
early predivisional
Figure 1.8. Caulobacter cell cycle. Asymmetric cell division yields one swarmer cell and one
stalked cell. Stalked cells immediately initiate DNA replication while swarmer cells differentiate
into stalked cells to proceed through the cell cycle. Key events are indicated in text while visible
polar morphogenetic events are highlighted in red. Cell cycle phases are color-coded.
Much of the early work on Caulobacter focused on dissecting these visible polar
morphogenetic events. Flagellar morphogenesis was especially amenable to genetic
analysis through the isolation of non-motile mutants [55]. Through detailed epistatic
analysis of these mutants, a specific hierarchy of flagellar assembly was established (Fig.
1.9). This hierarchy consists of four classes of genes that are expressed in an order that
corresponds to the order of flagellar assembly [55]. Expression of the next level in the
hierarchy requires proper expression of the previous level [55]. The flagellum is
assembled from the inside out with class II genes encoding for proteins that make up the
28
internal membrane structures including the MS ring, switch, and a secretion apparatus
that allows for the export of the extra-cytoplasmic components [55]. The class III genes
encode proteins that constitute the periplasm spanning basal body as well as the
extracellular hook while class IV genes encode the proteins of the flagellar filament [55].
The class I designation was reserved for the initiator(s) of flagellar biosynthesis, which
was ultimately identified as the response regulator CtrA [56].
class IV
I
fljJ
fijM
fljK
fljL
fjN
fijO
filament
flgF*
flgBC
flgi
hook
class Ill
flgK
hook operon
basal body
..
fihA
class 11
class I
fliF*
jffiQ*
fLM
SfliOP
ctrA
Figure 1.9. Flagellar assembly. Flagellar assembly requires the expression of four classes of
genes. The flagella is synthesized from the inside out with the class 11,111,
and IV genes coding for
structural components of the basal body, hook, and filament respectively. CtrA is the only class I
gene and initiates the hierarchy. Asterisks indicate operons. Adapted from [55, 57].
In addition to regulating flagellar biogenesis, CtrA is an important regulator of the cell
cycle and is essential for viability [55, 56]. Initially, CtrA was found to bind a motif in
class II promoters that had previously been identified in the promoter of the gene for an
essential methyltransferase,
ccrM, and in the origin of replication [56]. These
observations gave the first clues to the importance of CtrA for the regulation of many
different developmental processes.
CtrA, master of the Caulobacter cell cycle
Subsequent work established CtrA as a master regulator of Caulobacter cell cycle
because it inhibits the initiation of DNA replication and regulates the transcription of a
large set of cell cycle genes (Fig. 1.10). Like most response regulators, CtrA is activated
by phosphorylation [56] and regulated changes in CtrA phosphorylation (CtrA-P) drive
progression of the cell cycle. CtrA-P is abundant in G1 swarmer cells, where it inhibits
the initiation of DNA replication by binding the origin [58, 59]. The cell then clears CtrA
at the G1-to-S transition through a combination of dephosphorylation and degradation,
thereby releasing the origin and permitting initiation in stalked cells [58]. After being
cleared from the cell, CtrA is subsequently transcribed and phosphorylated in
predivisional cells, where it directly activates the expression of approximately 100 genes
involved in cell cycle progression and morphogenesis [60]. Thus, CtrA controls the
asymmetric morphogenesis of Caulobacter as well as the asymmetry in replicative
capacities of the two daughter cells.
CtrA-P
GI
G2
S
CtrA-actIvated
Origin repression
CtA rotoyiA
one ex rslon
tAphshrlto
Figure 1.10. CtrA activity and regulation through the cell cycle. The presence of CtrA-P is
indicated in blue. Cell cycle specific activities of CtrA are indicated in white boxes and regulation
of CtrA is indicated in grey boxes.
In Caulobacter both CtrA and the widely conserved replication initiation factor DnaA
regulate the initiation of DNA replication [61]. DnaA binds at least four sites in the origin
of replication and promotes origin melting (Fig 1.1 1A) [59]. Although the mechanisms
are not yet clear, changes in DnaA activity set the periodicity of the Caulobacter cell
cycle [62]. DnaA must be activated to initiate each round of replication then deactivated
to prevent over-replication. On top of the regulation by DnaA, CtrA blocks DNA
replication specifically in swarmer cells through the occupancy of five binding sites in
the origin of replication (Fig. 1.11 A) [59, 62]. Sites D and E are immediately adjacent to
two of the conserved DnaA boxes suggesting that CtrA binding may occlude DnaA (Fig)
[59]. In addition, sites A and B overlap and repress the hemE promoter, expression from
which is also thought to play a role in origin melting and initiation (Fig 1.11 A) [63].
Mutating three of the CtrA binding sites (sites B,C, and D) in the origin of replication
leads to premature initiation, i.e. shorter periods between replication events, and thus
chromosome accumulation emphasizing the importance of CtrA silencing of the origin
[62].
Progression of the Caulobactercell cycle also depends heavily on regulated transcription
[64]. Developmental genes are expressed just-in-time for their function. For instance, the
flagellar gene hierarchy is expressed in predivisional cells just as the flagellum is
assembled [60]. Expression of 533 genes in the Caulobacter genome is cell cycle
regulated and expression of nearly a quarter of these genes depends on CtrA [64].
Chromatin-immunoprecipitation and microarray experiments have shown that CtrA
directly regulates approximately 100 of these genes (Fig. 1.11B)[64]. CtrA activates
expression of the majority of its regulon in predivisional cells. About a dozen genes are
A.
C
D
E
RB
Orgin of Replication
B.
*
,1
CtrA reaulon
CC#
gene
CC396
CC2700
CC70
CC2704
CC1211
CC3474
CC3445cigT
CC3476
CSNW
p
protein function
L-lactata 2-monooxygenaea
reguato
n
potllc
hyote protein
cIe protein
CCV02
olgupr
hpthicl rlt
a
i
RNA potynsrease gator
in
tlwpto prote
h
fin
r
ra
ily
hprota
gaWrm
e
= Ionrepresdor
CC363 cfA cg dl g tcranscrnptional regulator
CC0N6Omne@( Sadenoeytelornneeyntiietase
transcriptionalregulator,LaGIaml
C231CC2317 mrphl melluylacepllng c-eoti
prote=
CC2641 ft@Z cell divielonloj ldeet p p rotein
tein
flalin
modca
CCO23
R
CCO234 NmO fliin modification protein
CCO378on
D
metsylnratalaa
ypoeptng ohearotaxte protein
mthy
C-430 mh
- chemyottcu protein
CC043i cP
CCp432 doe
heaido mPro
regulator
CCO0 heA cheanotaxlrinaea
cco43 chew chemotaxiaadapterprotein
chaR
chemortexie
protein
mathyitrterae.
CC0436
prote
endncaio
k n
CC0436 citel
chemfotede
giutematemtityetr
ra blenheirgua
T
CC1490
regulator
00043 cheYl chemnotenirapne
CCOM3 citD
chemnotaxieprtn
00042 citeU chemnotabtproee
CC0440 cheYW chetnteae eapne regulator
che;;mxiepoti
CC041 citED
itglr motor switch protein
UG
CONS0
000907 fbS
fla
rti
flagellar mo tch
protei
0C0906 MN
C0009 11W responsaeregulator
COMti fgS flagagar baaiody rodprotein
=aa-bdrod protein
Wig fiegetr
CC0954
hoctaabody compleoprotein
CC09MIRAE flagatier
.1
-0.6
CC14t7 fliG Ilageflin oiiaio rti
fiagelier btoayntiieeiaregulator
CCI458 fnb
CC1469OWP feglrprt
o rti
Ngtrbeibd
COl ffpP
o rti
CC2OS4florG faelrbabd
tledcl ifeetain euao
CC2462 ple
cycle roepone regulator
dIffaretiatonloli
CC2463 rvK
002641 ftA
calldivisionprotein
cell division protein
CC2542 feQ
CC2NIl mnor
peptidoglycan aynthesis enzyme
CC2692 flaW
call
dintelan prolte
CC228 itfa holdfeetsyntheis protein
holdaatyetheetepWat
CC2629 itAll
CC2366 nouSrol
log(rdo)
0
0.6
1
~polaPlled
CC2NO
CC2961
00253 rpoN
CC=22
CC0429
CCO7U1
C00794 130
Ctiti0
CC1102
hy poM.tical- prtein
hyphtical protein
RN.pO1ymwrevesigma-fi factor
hypotheticalProtein
hypothetia protein
lagi
faein
conerad hypothetical protein
011"ieiclprlt
00136 cop
CC2324
ATP-depontM proltee
hyid hslotn kIdas
CC2943 cp5
C02544 cpeD
CC2ll44 cpaD
CC21141cpall
CC2947 cptA
CUMS4 ptA
CC3219 CC3205 rcdA
COl03 aciP
plleeaaebly protin
plus assemblyprotein
plluaassemblyprotein
pus asamubly protei
pllueaseemblyprotein
pilot subunit protein
hybrid hisetc kinae
loial
prtoti
hyp
CtrA tnrnacriltinal Inhibiter
prolytoc subunit
Figure 1.11. Dual function of CtrA in regulation of the origin and cell-cycle regulated
transcription. (A). Organization of regulatory elements in the origin of replication. CtrA binding
sites are indicated in blue and DnaA sites are indicated in green. The developmentally regulated
hemE promoter is indicated by a pink arrow [59]. (B) Expression of the CtrA regulon in two CtrA
loss of function strains ctrAt' and cckAt. Genes repressed by CtrA are in yellow and those
activated by CtrA are in blue [65].
directly repressed by CtrA and only expressed at the G1-S transition when CtrA is
cleared from cells (Fig 1.11 B). The CtrA regulon consists of genes involved in diverse
functions like polar morphogenesis, DNA methylation, cell wall remodeling, proteolysis,
and other two-component proteins and transcription factors [60].
CtrA regulates transcription by binding a specific DNA sequence found in target
promoters, and phosphorylation likely activates CtrA by increasing its affinity for DNA
[66]. Although the specific mechanism of activation of CtrA is unknown, activation by
phosphorylation of related response regulators has been studied in atomic detail [37]. In
general, phosphorylation of the receiver domain, a doubly wound a/p fold, results a
significant displacement of the a4-p5-a5 surface (Fig. 1.12A) [37]. Phosphorylation of
the conserved aspartate changes the orientation of two key switch residues (Ser/Thr on P4
and Phe/Tyr on
p5)
from facing away from (exposed) to toward (buried) the active site,
thus transmitting phosphorylation status to changes in protein structure [37]. In the
related protein PhoB to E. coli, structural changes induced by phosphorylation lead to
homodimerization of the a4-p5-a5 surfaces, likely bringing together the DNA binding
domains of each monomer in a conformation that allows them to bind to respective halfsites on the DNA (Fig. 1.12B) [67]. Although the details of RR activation probably vary
for each specific response regulator, CtrA is likely to be activated by a similar
phosphorylation induced dimerization mechanism.
a4
al
B.
.
active
dimer
DNA
a2
Figure 1.12. PhoB activation by phosphorylation. (A) Crystal structures of the phosphorylated
vs. unphosphorylated PhoB receiver domain. Green indicates the unphosphorylated structure
while purple is phosphorylated. Key switch residues are indicated in orange and red. Image from
[37]. (B) Phosphorylation of PhoB results in homodimerization positioning the DNA binding
domains to interact with DNA [67].
How does CtrA binding to DNA regulate transcription? The short answer is that CtrA
bound to target promoters can recruit RNA polymerase to that promoter and thus activate
transcription [68]. The majority of genes activated by CtrA contain the CtrA binding
motif (TTAA-N7-TTAA) near the -35 position of the promoter and binding of a CtrA
dimer is thought to recruit RNA polymerase (Fig. 1.13A) [68]. In contrast, genes that are
repressed by CtrA typically have a CtrA binding motif overlapping the -10 position of the
promoter (Fig. 1.13B) [69]. In these cases CtrA binding probably occludes RNA
polymerase from binding the promoter thereby repressing transcription. Finally, some
genes contain multiple CtrA binding sites. For instance, pilA has a typical -35 site plus
three additional upstream CtrA sites, the regulatory consequences of which are unknown
(Fig. 1.14C) [70].
B.
P1
-35
0
-35
-10
ctrA
C.
HA
Figure 1.13. Activation of transcription by CtrA. (A) CtrA (blue barbell) may activate
transcription by binding the -35 box of target promoters and recruiting RNA polymerase. (B) CtrA
inhibits transcription by binding the -10 box and occluding RNA polymerase. (C) Multiple CtrA
binding sites regulate genes such as pilA.
Regulation of CtrA
In accordance with the importance of CtrA for cell cycle progression and viability,
Caulobacter has evolved several redundant mechanisms for regulating the activity of
CtrA. These include the regulated transcription of ctrA and cell-type specific
phosphorylation and proteolysis.
Like the expression of the CtrA regulon, the expression of the ctrA gene is cell cycle
regulated. ctrA is first transcribed in stalked cells shortly after the initiation of DNA
replication with expression peaking in late predivisionals (Fig. 1.14A). The ctrA gene has
two distinct promoters, an early P1 and a late P2 promoter, and expression is subject to
both negative and positive autoregulation (Fig. 1.14A) [69]. P1 contains a CtrA binding
site in the -10 box and is repressed by CtrA while P2 has a CtrA binding site in the -35
box and is activated by CtrA (Fig. 1.14B) [69]. Expression from P1 turns on by an
unknown mechanism following the initiation of S phase when CtrA is cleared from the
cell, leading to an initial burst of CtrA protein [69]. The newly synthesized CtrA then
feeds back to inhibit transcription from P1 but activate expression from the stronger P2
promoter, leading to the rapid accumulation of CtrA [59]. Thus the negative and positive
feedback loops ensure CtrA is produced to high levels shortly after being cleared during
the GI-S transition [71].
A
ctrA
P1
P2
relative time of expression
P1
-10
P2
-35
T T
Repressed Activated
Figure 1.14. Regulation of CtrA transcription. (A). Expression of ctrA as a function of the cell
cycle including the contribution of the P1 and P2 promoters. [69] (B). Negative and positive
feedback at the ctrA promoter. Each CtrA monomer is represented by a blue barbell.
CtrA is also activated by phosphorylation via a phosphorelay similar to that used in B.
subtilis sporulation [72]. The hybrid histidine kinase CckA first autophosphorylates and
then passes a phosphoryl group to an attached receiver domain, which can transfer it to
the histidine phosphotransferase ChpT which then directly phosphorylates CtrA (Fig.
1.15) [73]. CckA can also act as a phosphatase and pull phosphate off CtrA by reversing
the phosphorelay [74]. CckA is activated as a kinase by DivL at the swarmer pole and
becomes a phosphatase through localization at the stalked pole where DivL is either
absent or directly inhibited by the phosphorylated, single domain response regulator
DivK (Fig. 1.15) [74]. DivK is phosphorylated at the stalked pole by its cognate kinase
DivJ and dephosphorylated at the swarmer pole by the bifunctional kinase PleC (Fig.
1.15) [75]. Thus the pole specific activity of DivK dictates CckA activity and ensures
CtrA is phosphorylated in nascent swarmer cells and dephosphorylated in the stalked
compartment through the establishment of a gradient of CtrA-P [74, 76, 77].
P
P
P
P1
swarmer pole
stalked pole
Figure 1.15. Differential activation of CtrA at the swarmer and stalked pole. (A) The hybrid
HK CckA exhibits kinase activity at the swarmer pole activates CtrA through the ChpT
phosphorelay. (B) Swarmer pole factors promote CckA phosphatase activity resulting in
dephosphorylation of CtrA at the stalked pole [76].
In addition to transcriptional control and phosphorylation, CtrA activity is also regulated
by proteolysis. CtrA proteolysis is regulated by the same phosphorelay that
phosphorylates CtrA (Fig. 1.16) [73]. In addition to CtrA, ChpT also phosphorylates the
single domain response regulator CpdR [73]. Unphosphorylated CpdR is required for
regulated proteolysis of CtrA by the ClpXP protease during the G1-S transition [78]. In
swarmer cells when CckA is active as a kinase, CpdR is phosphorylated and thus unable
to mediate CtrA proteolysis. When CckA kinase activity is down-regulated by
phosphorylated DivK during the G1-to-S transition CpdR becomes dephosphorylated,
enabling it to somehow stimulate CtrA proteolysis. CtrA is targeted for proteolysis by a
bipartite degradation signal, present in the first 56 amino acids and in the last 15 amino
acids [79]. In addition, the protein RcdA is thought to contribute to proteolysis by helping
bring CtrA to ClpXP although RcdA does not affect CtrA proteolysis by ClpXP in vitro
[80, 81].
stalked pole
oe 00 0'CpdR
CckA
p1
P
CtrA
p
ChpT
p
P
CpdR
CtrA
Figure 1.16. CtrA proteolysis by CipXP. Dephosphorylation of CpdR leads to localization of
CipXP to the swarmer pole and CtrA proteolysis.
Identifying missing regulators of CtrA.
Despite our detailed understanding
of the regulation of CtrA by transcription,
phosphorylation and degradation, several observations suggest that CtrA may be subject
to additional modes of control. For instance, constitutive expression of a non-degradable
allele of ctrA is not lethal while expression of a non-degradable, phosphomimetic allele is
suggesting that dephosphorylation is necessary and sufficient to deactivate CtrA at the
G1-S transition. Although CckA phosphatase activity at the swarmer pole normally
dephosphorylates CtrA, CckA phosphatase activity is also not essential [74]. The mutant
CckA(V366P) lacks phosphatase activity in vitro but can replace wild-type CckA without
severe consequences to cell cycle progression [74].
This raises the possibility of an
unidentified CtrA phosphatase. Given the precedence of multiple phosphatases in the B.
subtilis sporulation pathway, it is possible that CtrA may be dephosphorylated directly or
indirectly by one or more phosphatases.
Another missing piece of the puzzle involves the absence of CtrA transcriptional activity
in swarmer cells. Although CtrA is phosphorylated and abundant in both swarmer and
predivisional cells, the majority of CtrA activated genes are expressed only in
predivisional cells [60]. This raises the possibility that CtrA lacks a co-activator in
swarmer cells or that there exists a swarmer-specific inhibitor of CtrA activity.
In the following chapters I will document our discovery and characterization of a CtrAspecific transcriptional inhibitor called SciP. In chapter 2, I will describe the
computational screen we used to identify SciP as well our characterization of its
expression during the cell cycle. I will also present a series of experiments that lead to a
model in which SciP interacts directly with CtrA at CtrA dependent promoters and blocks
the recruitment of RNA polymerase. In chapter 3, I will describe the post-transcriptional
regulation of SciP itself. Specifically I will demonstrate that SciP is proteolyzed by the
Lon protease and that a failure to degrade SciP during the G1 -S transition has significant
deleterious effects on cell cycle progression in Caulobacter.
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Chapter 2.
A cell-type-specific protein-protein interaction modulates
transcriptional activity of a master regulator in
Caulobactercrescentus
This work was published in Molecular Cell, 2010. 39(3): p. 455-67 by Kasia G Gora,
Christos G Tsokos, Y Erin Chen, Balaji S Srinivasan, Barrett S Perchuk, Michael T Laub
B.S.S. designed and coded the computational screen. K.G and M.T.L analyzed data and
wrote the manuscript. K.G. performed all the experiments with the expection of in vivo
phosphorylation performed by C.G.T, microarrays perfomed by B.S.P, and in vitro
phosphorylation peformed by Y.E.C.
Abstract
Progression through the Caulobactercell cycle is driven by the master regulator CtrA, an
essential two-component signal transduction protein that regulates the expression of
nearly 100 genes. CtrA is abundant throughout the cell cycle except immediately prior to
DNA replication. However, the expression of CtrA-activated genes is generally restricted
to S-phase. Here, we identify the conserved protein SciP and show that it accumulates
during GI where it inhibits CtrA from activating target genes. The depletion of SciP from
G 1 cells leads to the inappropriate induction of CtrA-activated genes and, consequently, a
disruption of the cell cycle. Conversely, the ectopic synthesis of SciP is sufficient to
inhibit CtrA-dependent gene expression, which also disrupts the cell cycle. We
demonstrate that SciP binds to CtrA without affecting stability or phosphorylation;
instead SciP likely prevents CtrA from interacting with RNA polymerase. CtrA activity is
thus tightly regulated by a novel protein-protein interaction which is critical to cell cycle
progression.
Introduction
All dividing cells must ensure the correct timing and order of DNA replication,
chromosome segregation, and cell division. These cell cycle events are carefully
orchestrated to ensure genome stability and cellular survival. Their execution and, hence,
proper cell cycle progression requires complex transcriptional programs, with batteries of
genes turned on and off according to their times of action during the cell cycle [1]. These
gene expression programs are controlled by master regulators whose activities are tightly
regulated. For example, in metazoans the transcription factor E2F regulates the
expression of hundreds of genes during late G1, many of which are critical to the ensuing
S phase [2]. To prevent the premature, inappropriate expression of E2F targets in early
G 1, the retinoblastoma protein Rb binds directly to E2F and prevents it from stimulating
gene expression, but without disrupting DNA binding [3]. The regulated inactivation of
Rb by phosphorylation in late G1 thus unleashes E2F activity and propels cells into S
phase. In the yeast S. cerevisiae, the transcription factors SBF and MBF drive the
expression of nearly 200 genes in late G1 [4]. Analogous to Rb, the yeast protein Whi5
binds directly to SBF and MBF to block transcription during early G1; the subsequent
hyperphosphorylation of Whi5 relieves this inhibition and permits SBF and MBF to drive
gene expression crucial to cell cycle progression [5].
The regulation of gene expression during the bacterial cell cycle remains poorly
understood. The Gram-negative bacterium Caulobacter crescentus is a tractable model
for elucidating the mechanisms underlying the prokaryotic cell cycle [6, 7]. The
Caulobacter cell cycle begins with a Gl-phased swarmer cell that is motile and cannot
initiate DNA replication until it differentiates into a stalked cell. Concomitant with the
swarmer-to-stalked cell transition, the cell enters S phase during which it replicates its
chromosome and partitions the new chromosomes to opposite poles of the predivisional
cell. Each cell division is asymmetric, producing two different daughter cells - a G1
swarmer cell and a stalked cell that can immediately enter S phase. DNA replication in
Caulobacteroccurs in a once-and-only-once manner, resulting in distinct GI, S, and G2
phases. Additionally, Caulobactercells can be easily synchronized, facilitating temporal
analysis of the cell cycle.
In Caulobacter,cell cycle progression requires the periodic activation and inactivation of
the master regulator CtrA, which functions both as a transcription factor for nearly 100
genes and as a direct repressor of DNA replication initiation [8-10]. CtrA is a response
regulator and hence part of a two-component signaling pathway, the dominant form of
signal transduction in bacteria [11, 12]. These pathways typically initiate with a sensor
histidine kinase that autophosphorylates and then transfers the phosphoryl group to a
cognate response regulator. Phosphorylation of the response regulator activates an output
domain that can drive changes in cellular behavior. Phosphorylation of CtrA enhances its
ability to bind DNA [13, 14] and, consequently, to modulate transcription and silence the
origin of replication.
CtrA activity is controlled by a combination of regulated phosphorylation, proteolysis,
and transcription [8, 15-19]. Phosphorylated CtrA is abundant in swarmer/G1 cells where
it silences the origin of replication [10], likely by occluding the replication initation factor
DnaA. At the G1-S transition, CtrA is dephosphorylated and degraded, freeing the origin
and enabling DNA replication to commence. As cells proceed through S phase, ctrA is
transcribed, and the newly synthesized CtrA is stabilized against proteolysis and activated
by phosphorylation. CtrA then activates the expression of more than 60 target genes,
many of which are important for late stages of the cell cycle and cell division [20].
Following septation, CtrA is maintained in the phosphorylated state in daughter swarmer
cells but proteolytically cleared from daughter stalked cells to permit another round of
DNA replication.
These mechanisms ensure that CtrA is abundant and phosphorylated in both swarmer and
predivisional cells. Consistent with this pattern of activity, CtrA-repressed genes are
expressed only in stalked cells after CtrA has been degraded. However, most of the genes
that are directly activated by CtrA are transcribed only in predivisional cells [20]. A
handful of CtrA-activated genes have mRNAs that are also abundant in swarmer cells,
but this pattern may result from transcription in predivisional cells followed by
stabilization of the mRNA in swarmer cells [21, 22]. These previous findings thus imply
the existence of either a predivisional-specific transcriptional cofactor for CtrA or a
swarmer cell-specific inhibitor of CtrA-dependent gene expression. Neither such factor
has been identified to date.
Here, we identify SciP, or small CtrA inhibitory protein, and demonstrate that it binds
directly to CtrA and inhibits its ability to activate transcription without disrupting DNA
binding or the repression of certain target genes. SciP is restricted to G1 swarmer cells
and thus plays a role in establishing the differential fates of daughter cells. SciP is critical
for proper cell cycle progression and we show that SciP levels must be tightly regulated,
as the ectopic production of SciP in predivisional cells is sufficient to block CtrAdependent gene expression and, consequently, proper cell cycling. SciP is highly
conserved and present in nearly every organism containing a CtrA ortholog emphasizing
the key role this molecule plays in cell cycle progression in a wide range of bacteria. SciP
represents a new class of molecules that modulate the output of two-component signaling
pathways without affecting phosphorylation, indicating that these critical signaling
pathways are subject to even more complex regulation than previously appreciated.
Results
Identification of sciP as a candidate regulator of CtrA and the cell cycle
To identify putative regulators of CtrA transcriptional activity we used a bioinformatic
screen for genes that are coexpressed, coinherited, or colocated in bacterial genomes with
ctrA [23].
We previously used a similar approach to identify the direct CtrA
phosphodonor, ChpT [15]. This screen identified CC0903, which we named sciP for
small CtrA inhibitory protein. The sciP gene is annotated to encode a 93 amino acid
protein of no known function. Orthologs of sciP are often found immediately adjacent to
or within a few genes of ctrA in a-proteobacteria such as Rhodobacter sphaeroides,
Sinorhizobium meliloti, Bartonellabacilliformis, Brucella melitensis, and Mesorhizobium
loti (Fig. 2. 1A). Orthologs of sciP were found in virtually all a-proteobacteria indicating
strona coconservation with ctrA with most orthologs nearly 70% identical and 85%
similar to the C. crescentus SciP (Fig. 2.2).
chpT
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Figure 2.1. Identification of sciP as a key regulator of cell cycle progression. (A) Genomic
organization of ctrA, chpT, and sciP orthologs in C. crescentus and other a-proteobacteria. (B)
Growth curve of AsciP and wild type grown in PYE. (C) Cellular morphology (top) and flow
cytometry analysis (bottom) of AsciP compared to wild type. Samples were treated with rifampin
and 50,000 cells from each population were analyzed. (D) Quantification of the distribution of cell
lengths in C. (E) Cell cycle abundance of SciP. Synchronized wild-type swarmer cells were
released into PYE and allowed to proceed through one cell cycle with samples collected every 10
min for Western blot analysis with CtrA or SciP anti-sera. After 100 min, the culture was again
synchronized to separate swarmer and stalked daughter cells (lanes labeled SW and ST). The
cell cycle diagram shown on top indicates relative stage of the cell cycle for each time point.
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of alpha-proteobacterial genomes were aligned and conservation visualized using BOXSHADE.
Residues shaded in black and grey match or are similar to, respectively, the consensus at that
position.
To test whether sciP is important for cell cycle progression, we deleted the chromosomal
copy of the gene from the wild type (CB15N) and replaced it with a tetracyclineresistance cassette to produce the strain AsciP. In the C. crescentus genome, sciP is
located between two operons that encode flagellar proteins, but is transcribed
independently [24], so the deletion of sciP is unlikely to affect nearby genes. Although
the AsciP strain is viable, it exhibited significant growth and cell cycle defects with a
doubling time of -120 minutes in rich PYE medium compared to -80 minutes for the
wild type (Fig. 2. 1B). Cells from a mid-log phase culture of AsciP were filamentous with
lengths ranging from slightly elongated to extremely filamentous (Fig. 2.lC). The
filamentous cells were either smooth and unpinched or had a single constriction which is
likely the site of active cell division. Flow cytometry analysis of the AsciP strain revealed
a modest, but reproducible accumulation of cells with 3N chromosomal content
compared to the wild type (Fig. 2.1C). These growth, morphological, and chromosomal
content defects were rescued by providing a wild-type copy of sciP in trans at the
chromosomal xylX promoter (see below). These results indicate that SciP is required for
proper cell cycle progression in Caulobacter.
SciP is present only in GJ swarmer cells
To determine when SciP functions during the cell cycle, we measured its abundance
during the cell cycle using a polyclonal antibody specific to SciP. Wild-type swarmer
cells were then harvested and allowed to proceed synchronously through the cell cycle
with samples isolated every 10 minutes for immunoblot analysis (Fig. 2.lD). We found
that SciP was abundant in swarmer cells but rapidly disappeared at the Gi-S transition
and did not accumulate again until very late in the cell cycle, coincident with the first
appearance of new swarmer cells (Fig. 2.3). This temporal pattern suggests that SciP may
be restricted primarily to the Gi/swarmer stage of the cell cycle. To test this conclusion,
we used density centrifugation to separate new swarmer cells from new stalked cells
immediately following cell division. Immunoblots of samples from each population
revealed that SciP was present only in the swarmer fraction (Fig. 2.1E, last two lanes).
Together, these data indicate that SciP is present primarily, and perhaps exclusively, in
Gi-phased swarmer cells. We note, however, that sciP mRNA was previously found to
be cell cycle-regulated, reaching peak levels in predivisional cells [25]. The delay in SciP
accumulation following its transcription suggests sciP expression may be posttranscriptionally regulated.
time (min.)
0
10
20
30
40
150
0
60
70
80
90
100
IN 2N
Figure 2.3.
Flow cytometry analysis of cells from Figure 2.1E. Synchronized wild-type
swarmer cells were released into PYE and allowed to proceed through one cell cycle with
samples collected every 10 min for FACS analysis.
To better characterize the cell cycle function of SciP, we created a depletion strain in
which the chromosomal copy of sciP was deleted and a wild-type copy of sciP was
placed at the chromosomal xylX locus under the control of the xylose-inducible, glucoserepressible promoter Pyz. When grown in PYE supplemented with xylose, the depletion
strain exhibited morphology, chromosome content, and SciP protein levels similar to wild
type (Fig. 2.4A-C). Upon shifting to medium supplemented with glucose, SciP was
rapidly depleted, with no protein detected after one hour (Fig. 2.4B). We then examined
the morphology and chromosomal content of cells following a shift from xylose to
glucose. After two hours, the culture showed a significant increase in cells with one
chromosome, suggesting that the loss of sciP may cause a transient arrest in GI (Fig.
2.4C). After 12 hours, the depletion strain had overcome this delay and reached a steadystate in which the chromosomal content profile was nearly identical to the sciP deletion
strain. By 12 hours, cells also exhibited an unpinched, filamentous morphology similar to
the AsciP strain (Fig. 2.4A).
To further examine whether sciP affects the Gi-S transition, we synchronized the
depletion strain grown in xylose and then released swarmer cells into medium containing
either xylose or glucose to maintain or inhibit SciP production, respectively. Cell cycle
progression for each population of cells was monitored by flow cytometry (Fig. 2.3) and
SciP levels examined by Western blotting (Fig. 2.4E). As expected, for cells released into
xylose, SciP was initially abundant but eliminated at the G1-to-S transition (30 min. time
point), as seen with wild-type cells. SciP accumulated again in these cells during the
latest stages of the cell cycle, at a time coincident with cell division and the generation of
new swarmer cells. Although sciP is constitutively expressed, SciP levels still drop after
GI (Fig. 2.4E), further demonstrating that sciP expression is post-transcriptionally
regulated. Following cell division, daughter cells from the culture grown in xylose
initiated a new round of DNA replication, manifest in the flow cytometry profiles by a
shift of the entire IN chromosomal peak toward 2N by the 100-minute time point.
PY:scIP
sciP depletion
xylose
glucose
synchronize, release Into
xylose or glucose
'I
time (min.)
0
20
B
glucose
hrs post-shift:
0
1
2
3
xylose
4
4
SciP
C
P xY,:Scip
Ir
wash, release into glucose
IN
2N
--- glucose
time post-synchronization (min.)
0
30
90
xylose
IN
2N
glucose
Figure 2.4 SciP depletion phenotype. (A) Cellular morphology of the sciP depletion strain
grown in PYE supplemented with xylose or shifted to PYE with glucose for 12 hrs to deplete SciP.
(B) Western blot analysis showing SciP levels post-shift to glucose or maintained in xylose at the
times indicated. (C) Flow cytometry analysis of chromosomal content in the sciP depletion strain
grown in PYE supplemented with xylose and then shifted to PYE with glucose. (D) Flow
cytometry analysis of synchronized swarmer cells from the depletion strain released into PYE
supplemented with glucose or xylose. Arrows indicate the delay in DNA replication initiation seen
after the first cell division for the strain depleted of sciP. (E) Western blot analysis of SciP levels in
cultures synchronized as in panel D,at the time points indicated.
When the swarmer cells of the depletion strain were released into glucose, SciP was also
initially abundant and then rapidly eliminated at the Gl-S transition (Fig. 2.4E).
However, in these cells SciP remained undetectable throughout the remainder of the first
cell cycle and after cell division, indicating a complete repression of sciP expression.
These cells divided at approximately the same time as the control cells grown in xylose,
demonstrating that SciP is not required for progression through the late stages of the cell
cycle or for cell division (Fig. 2.4D). However, following the first cell division (80 min
time point), approximately half of the daughter cells were delayed in initiating DNA
replication for up to 60 minutes, relative to the control cells, (Fig. 2.4D). This result
supports our conclusion that sciP is important for the Gl-S transition and proper cell
cycling.
We also resynchronized the culture of cells released into glucose after the first cell
division. However, this double synchrony produced a low yield of swarmer cells in
contrast to cells released into xylose. Based on flow cytometry (Fig. 2.4D) and time-lapse
microscopy (data not shown) we concluded that the lack of swarmer cells was not due to
a defect in cell division. Instead, cells lacking sciP likely do not undergo the change in
buoyant density that facilitates synchronization by centrifugation. The swarmer cells that
were harvested, though, showed a significant delay in initiating DNA replication.
Depletion of sciP affects CtrA-dependent gene expression
Next, we tested whether the lack of sciP in G1 swarmer cells affected gene expression,
particularly the CtrA regulon. Again, we synchronized the depletion strain and released
swarmer cells into media supplemented with either glucose or xylose to repress or
maintain SciP levels, respectively. After 100 minutes, once all cells in each culture had
divided, we isolated RNA from each culture and compared the samples by competitive
hybridization on DNA microarrays. These post-division samples include both swarmer
and stalked cells because, as noted, the yield of swarmer cells following a second
synchronization of the depletion strain is low. However, because SciP is only abundant in
swarmer cells (see above), differences in gene expression will only reflect differences in
swarmer cell gene expression.
Nearly all of the genes showing significant changes in expression level were CtrAdependent genes (GEO GSE22062). Figure 5 shows the changes in expression for genes
that are directly regulated by CtrA. Strikingly, nearly every CtrA-activated gene was
expressed at significantly higher levels in the cells depleted of sciP. In contrast, the
expression of CtrA-repressed genes were unchanged. These data indicate that SciP is
required to silence the expression of CtrA-activated genes in swarmer cells, thereby
restricting their expression to predivisional cells.
Overexpressing sciP disrupts the cell cycle and drives the down-regulation of CtrAdependent genes
The data thus far are consistent with SciP being a G1-specific inhibitor of CtrA. To test
whether SciP is sufficient to inhibit CtrA transcriptional activity, we examined the
consequence of forcing the synthesis of SciP in predivisional cells when it is normally
absent. For this experiment we used pHXM-sciP, a high-copy vector for expressing sciP
with an N-terminal M2-epitope tag. Although strains overexpressing either sciP or M2sciP gave similar phenotypes, only the latter could be synchronized. Inclusion of the tag
also enabled us to distinguish plasmid and chromosomally-encoded SciP, as needed
below.
The strain harboring pHXM-sciP exhibited normal cellular morphology and chromosome
content when grown in glucose to repress expression (Fig. 2.6A). To overproduce SciP,
cells were shifted to medium containing xylose. Immunoblots showed that SciP levels
increased approximately 7-fold after 4 hours of growth in xylose. After four hours postinduction, cells had become filamentous and accumulated up to four chromosomes per
cell, phenotypes consistent with a disruption of cell division (Fig. 2.6A). Immunoblot
analysis of synchronized cells demonstrated that SciP was now abundant throughout the
cell cycle, overcoming the regulatory mechanisms that usually limit it to swarmer cells
(Fig 2.6B).
To test whether the overproduction of SciP was sufficient to down-regulate CtrAactivated genes, we again used whole genome DNA microarrays. Cells harboring pHXMsciP were shifted to xylose for 2 hours and mRNA from these cells was compared to
mRNA from wild-type cells grown in identical conditions. Nearly every CtrA-activated
gene showed significant down-regulation in the cells overproducing SciP (Fig. 2.5). The
magnitudes of these drops in expression were comparable to, although slightly less than,
those seen in ctrAs and cckA"s strains [20, 26] grown at the restrictive temperature relative
to wild type (Fig. 2.5). As with the sciP depletion strain, genes repressed by CtrA were
not significantly affected by the overproduction of SciP.
*
/O
*
CtrA regulon
protein functlon
gene
2.monoygomeoe
Lno
CU270
hypoteticalprotein
ein
0027012
bypotbeilcolppro
C027114
hoerypotticalpro
0C1211 hpohiolpo,
CC3474
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CC3475
siliT ONA
polynravensign. factor. ECF family
CC3476pi
CC48 pm
e po09
CC14628
VT oill
Plou"
CC14993InA
re8F lopren"
CC03641 A
Coal=A .CCO4
ine
n
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CC#
CC13
potei
lrwlo.
1nA
CC2620 M
poteinri
modp
CCO2329
CC23 lne
flac n mod s protn
CC238 c A DNAm rylborom
CC2430 -cpA mobyioooplingobonotaxls protein
obomoteols proof
CC0831 regulator
CC0432 -h.3 obectaxlero
CC0433 -isA
obemotai
le
CC10434 cho mocaimeplorprotein
itblnsiams
n
bsemote
prosan
CC 80 -0.0
,bsmosudi liermi hte urt 0,l.0.es
000436Chas
rulo
1CO37
ch*pP obnts response
P bnmotae poin
CCO42 cpaV
CC319 hell bmotaxisProte
000440 0 otnloteei rospnoeresgu etur
COO41
protein
CC251m
ur chloEdhvotisn
COMIS W9
fsgefr 11t opqoWln
P. W
CC2312
606 lEG fiegellarmotorpoite protn
AM
o fleg rver
protein
CCOUUY
CO11OROD repose eguitor
0C953 M
fiuoellerblesel~body
rodprotein
CCOSO4 ft ngskrben-body rod proteio
CCOW51
fiageller hoobkbeea body coropis protein
o prot n
CC1457006 Opetenmodificabit
00140ODT flegeiler bloweniels rngolotor
0CC145 Wg flagoller
0020614 flo goilbosabbody rodproteio
C=204pn stbd-call
dlflimetol regolotor
dlsiC o dilafridlellonioll
C02402
h000
ODflg l pro, cyclereponseregulator
002541
0.A selydiison protein
niiblonp
it r
old
C002042
002001 murG pop Idoglwn Synthesis onoyrn
sell d~lnproino
002002
MlW
COU IMI&Aboldlboyntemlprotein
002620 ltB
W~ldtsyntesis protein
002060 noul8 no0,protein'putatire
C2M
hypothetical noel
hypo0nolical
protein
CC2950
CC2051
yb.leprtl
lntre eooO
factor
000000
mom
hypothe.tical protein
00022
CO04Sypoarelcal
protein
000713 f
lgnl
lP
promn
= asovdbypothetical
001101'
CC1102 bypot
Vi
protein
001000
GI 00060
b y protein
00106113 sloP ATPdspeod*Mprotbe. prolotic subuit
002S24
hybridbintdbkw
blenn
bypolhnoftsl
protein
C3
0054 Op pus
... rte protein
protein
CC2543opeE PMioassembly
0C2344 Spe pilesaesemblyprotein
Coano ope
pum- mmooeiblv poten
002544 pp0
usilo eeeoblyprotein
pilo asemnbly protein
002-dyopA
00214 pNA plus ebntrti
0021,.ds hypothrecl eprotlnh
CC0903scAP CtrAtruoelpionl Inhbitor
lografto)
-1
-0.
0
0.
I
Figure 2.5 Effects of sciP overexpression and depletion on CtrA-dependent gene
expression. Whole-genome DNA microarrays were used to examine gene expression changes
in the strains indicated. The cckAts and ctrAt s mutants were grown in PYE at 28*C, shifted to 370C
for two hours, and compared to wild-type grown under the same conditions (data are from [20,
26]). For sciP overexpression, strain ML1749, which harbors pHXM-sciP, was grown in PYE
supplemented with xylose for 2 hrs and compared to wild type grown under identical conditions.
For sciP depletion, strain ML1750 was grown in PYE supplemented with xylose, synchronized,
and released into PYE supplemented with glucose or xylose. Each population of cells were grown
for 100 min, and then compared to each other. Expression changes for each gene are colorcoded according to the legend shown at the bottom. The genes shown include those previously
reported to be direct CtrA targets based on ChIP-chip analysis and whose expression changed
significantly in both cckAt' and ctrAt ' strains after a shift to the restrictive temperature [20, 26].
Note, the divK/pIeD operon was not identified by ChIP-chip analysis, but was subsequently
confirmed as a direct CtrA target [17]. sciP is included at the bottom of the list.
61
pHXM-sciP
pHXM-scIP
- glu = 2.4 t 1.2 (n = 372)
xyl = 6.8 T3.0 (n = 319)
-0nm
s0
xyiose
aeMnIA&A
i
0
%
40
20
0
2
4
8
6
10 12
14 16
18
20
length (sm)
C
time(min.) 0 10 20 30 40 80 60 70 80 g0 100 110
78
34
12
1 2 3 4 5 6
--sses01en
-
7 8
Mi,.ausa-M2-SciP
# of chromosomes
D
35
30
25
15
10
E
A
pHXM4cIP(glu)
* pHXM-scIP(xyl)
5
0
20
40
60
80
*wt
*AsciP
100
time (min.)
30
0
60
90
time (min.)
glucose
xylose
CtrA-P
:0%
c f
CtrA
c o"
0.75
Gr
0.5
a.
AscIP
pHXM-sc/P
(gu)
0.25
pHXM-cIP
(xyl)
0
av.
0
10
20
30
40
s0
60
Reaction time (min)
Figure 2.6 Phenotypic consequences of sciP overexpression. (A) Cellular morphology (top)
and flow cytometry analysis (bottom) of cells overexpressing sciP from a high-copy plasmid
(strain ML1749). A culture was grown in PYE glucose and sciP expression induced by the
addition of xylose for 4 hrs. (B) Westem blot for SciP in samples taken from a synchronous
population of cells overexpressing sciP. (C) The overexpression strain was synchronized and
released into glucose or xylose to repress or overexpress, respectively, sciP. ccrM mRNA levels
as a function of cell cycle time were measured using qRT-PCR and expressed relative to a
control, rho. (D) Pulse-chase analysis of CtrA stability in the strains indicated. For the
overexpression strain harboring pHXM-sciP, cells were grown in the M2G or shifted to M2G
supplemented with xylose for 1 hour to induce sciP expression. Each experiment was performed
in triplicate except for wild type which was done in duplicate; error bars represent standard error
of the mean. (E) Phosphorylation levels of CtrA in vivo. Cells harboring pHXM-sciP were grown
in media supplemented with glucose and then shifted to media with xylose for two hours. In each
case, CtrA was immunoprecipitated from cells after labeling with [y P]-ATP and then separated
by SDS-PAGE followed by exposure to a phosphorimage screen (top panel), with CtrA levels
measured by Western blotting (bottom panel). (F) CtrA occupancy of the chromosomal origin,
Cori, and pilA and ccrM promoters was measured by chromatin immunoprecipitation of CtrA from
DsciP and cells overexpressing sciP. Each measurement was done in triplicate and divided by
the average triplicate percent input for wild type; error bars represent the standard error of the
mean. (5) Dephosphorylation of CtrA by posphorelay reversal in the absence or presence of 1
1iM SciP.
Notably, the expression of sciP itself was strongly decreased in ctrA'' and cckA's (Fig. 3)
and overproducing M2-SciP from a high-copy plasmid led to a significant drop in the
levels of native, chromosomally-encoded SciP (see Fig. 2.8B). Previous ChIP-chip
studies did not include the sciP promoter on the microarrays used, but the predicted
regulatory region has a near-consensus CtrA binding site, suggesting sciP is a direct CtrA
target.
To confirm that the down-regulation of CtrA-activated genes was due to the ectopic
production of SciP in predivisional cells, we synchronized the overexpression strain,
released cells into media with either glucose or xylose, and monitored cell cycledependent expression of ccrM, a CtrA-activated gene, using quantitative PCR. Cells
grown in glucose showed the expected peak in ccrM expression in predivisional cells,
while cells grown in xylose never induced ccrM above background levels (Fig. 2.6C). In
sum, our gene expression studies demonstrate that SciP is sufficient to inhibit CtrA from
activating its target genes.
SciP does not affect CtrA degradation orphosphorylation
How does SciP inhibit CtrA-dependent transcription? To begin addressing this question,
we first examined the steady-state levels of CtrA following both sciP depletion and
overexpression and found that they did not change significantly in either case (data not
shown). We also measured the stability of CtrA in the sciP deletion strain and in a strain
overproducing SciP. In AsciP, the half-life of CtrA was 62 minutes, similar to the value
measured in wild-type cells (Fig. 2.6D). With the overexpression strain, CtrA was more
stable in cells grown in xylose to induce SciP than in cells grown in glucose (50 versus
173 minutes) (Fig. 2.6D). These data indicate that SciP does not down-regulate CtrA
activity by stimulating degradation but may stabilize it. To test the effect of SciP on the
phosphorylation of CtrA, we first reconstituted CckA-ChpT-CtrA phosphorelay in vitro.
The addition of SciP had no significant effect on the rate or steady-state levels of CtrA
phosphorylation
(data not shown). SciP also had no substantial effect on the
dephosphorylation of CtrA-P by phosphorelay reversal (Fig. 2.6H). We also measured
the phosphorylation
level
of CtrA
in vivo in cells overexpressing
sciP by
immunoprecipating CtrA after growth in the presence of [y3 2 P]-ATP. CtrA-P levels did
not change significantly after overexpressing sciP for two hours (Fig. 2.6E).
Finally, we tested whether sciP affects DNA-binding by CtrA using chromatinimmunoprecipitation (ChIP) to examine the occupancy of CtrA at two target gene
promoters and at the origin of replication in vivo. Neither the overexpression nor
depletion of sciP had a significant effect on CtrA occupancy of the origin of replication,
or the pilA and ccrM promoters (Fig. 2.6F). These findings also corroborate our global
gene expression data showing that overexpressing and depleting sciP affects CtrAactivated, but not CtrA-repressed genes. Additionally, these data are consistent with the
observation that cells overproducing SciP exhibited relatively moderate chromosomal
accumulation. If SciP disrupted CtrA phosphorylation or DNA-binding, we would have
expected more significant chromosomal accumulation, as seen with ctrA' and cckA's
mutants [10, 18]. The mild accumulation of chromosomes following sciP overexpression
is likely due to the disruption of cell division that results from SciP's inhibition of CtrA
target genes, many of which are important for cell division. Consistent with this
interpretation, we found that cells depleted of the essential cell division geneftsZ showed
a similar mild chromosomal accumulation phenotype (Fig. 2.7).
PxyrftsZ
xylose
1 2 3 4 5 6 7 8
glucose
1 2
3 4
5 6 7 8
# of chromosomes
Figure 2.7. Morphology and chromosomal content of tsZ depletion strain. Cells were
grown in either xylose to maintain expression of ftsZ or in glucose to repress ftsZ. Cells were
examined by light microscopy (top) or flow cytometry (bottom).
SciP binds directly to CtrA
Our data suggest that SciP could bind to CtrA to prevent it from interacting productively
with RNA polymerase to drive transcription of target genes. To test whether SciP
regulates CtrA via a direct protein-protein interaction, we first tested whether CtrA and
SciP interact in a yeast two-hybrid system. CtrA was fused to the activating domain of
Gal4 and SciP to the DNA-binding domain of Gal4. Together, these constructs
reconstituted Gal4 activity and drove expression of GAL1-HIS3 at a level sufficient to
enable growth on minimal medium lacking histidine (Fig. 2.8A). Neither CtrA nor SciP
alone was sufficient to activate HIS3 expression. Also, SciP did not interact with the
response regulator PhoB, supporting the notion that SciP binds specifically to CtrA.
To identify amino acids important for the CtrA-SciP interaction, we constructed a series
of alanine substitutions in SciP at sites that are (i) perfectly conserved in sciP orthologs
and (ii) solvent-exposed in an NMR structure of an ortholog from R. sphaeroides solved
by a structural genomics project (PDB: 2JRT) (Fig 2.9). In total we made five mutants,
R35A, R40A, E57A, Y62A, and Y68A, and found that each mutation, except E57A,
completely disrupted the CtrA-SciP interaction in the yeast two-hyrbid assay (Fig. 2.8A).
We then tested the activity of these alanine mutants in vivo by overexpressing them in
Caulobacter cells using the high-copy vector pHXM. Each alanine mutation, with the
exception of E57A, completely eliminated the sciP overexpression phenotype. (Fig.
2.8C). Immunoblots of the overexpression strains indicated that only the R35A and R40A
mutants accumulated to levels equivalent to overexpressed wild-type sciP suggesting the
other mutations may affect SciP stability (Fig. 2.8B). These results suggest that the severe
phenotype of sciP overexpression requires a direct interaction between SciP and CtrA,
and that residues R35 and R40 are critical for this interaction.
B
UO
- His
AD
BD
dIvJ
dvK
ctrA
-
ctrA
SCIP
phoB
sciP
-.
sciP
%
ctrA
sciP(R35A)
ctrA
sclP(R40A)
sCIP
G6 X
+ His
scP(R35A)
G
X
sc/P(R40A)
G
X
XG
M2-ScIP
ScIP
C
sclP
sc/P(R35A)
sclP(R40A)
---glucose
----
# of chromosomes
xylose
# of chromosomes # of chromosomes
Figure 2.8. SciP and CtrA interact directly in a yeast two-hybrid assay. (A-B) Yeast strain
PJ69-4A was co-transformed with plasmids harboring fusions of the indicated genes to either the
activation or DNA-binding domain of Gal4, as indicated, and spotted onto histidine deplete or
replete solid media. Each row contains serial 10-fold dilutions of overnight cultures normalized to
OD 0.5. Growth after 72 hours is shown. Growth on media lacking histidine indicates
reconstitution of Gal4 activity by protein-protein interaction. (C)Cellular morphology (top) and flow
cytometry analysis (bottom) of Caulobacter strains overexpressing sciP or the indicated alanine
mutants of sciP. Cultures were induced with xylose for 4 hrs. (D) Western blot analysis of
plasmid-encoded M2-SciP and chromosomally-encoded SciP for the strains from panel C grown
in glucose or xylose to repress or overexpress, respectively, the plasmid-borne M2-sciP.
Figure 2.9 Model of SciP structure. (A) NMR structure (PDB: 2JRT) of a SciP ortholog from R.
sphaeroides. (B) Residues R35 and R40, which are important for inhibiting CtrA-dependent gene
expression are highlighted in magenta with spacefilling.
SciP binds CtrA without disrupting DNA-binding
Next, we used electrophoretic mobility shift assays to test whether SciP could interact
with CtrA when CtrA is bound to target promoters. We added purified CtrA~P to
radiolabeled, 50-base pair fragments of the ccrM, pilA, andfliFpromoters, which contain
CtrA-binding sites [9, 19, 27]. As expected, CtrA bound these DNA fragments, leading to
a decrease in unbound DNA and the appearance of lower mobility bands (Fig. 2. 1A-C).
The intensity of these latter bands increased with increasing CtrA concentration and were
correlated with decreases in unbound DNA. CtrA did not bind to a DNA fragment from
the vanA promoter which lacks a CtrA-binding site (data not shown). We then added SciP
to each reaction and found that it led to a dramatic enhancement of DNA-binding as
evidenced by the nearly complete disappearance of free, unbound DNA at the lowest
concentration of CtrA and the appearance of low mobility bands at lower concentrations
of CtrA than seen without SciP (Fig. 2.1OA-C). We found that SciP alone did not bind
any of the probes tested, even at concentrations up to 10 [M suggesting it does not bind
DNA alone or does so only in the presence of CtrA. We also found that SciP(R35A) and
SciP(R40A) did not supershift the fliF promoter as observed with wild-type SciP (Fig.
2.101D). In sum, these data support the conclusion that SciP interacts directly with CtrA,
but without disrupting CtrA's ability to bind DNA. SciP thus likely inhibits CtrAdependent gene activation by preventing CtrA from interacting productively with RNA
polymerase.
Pe,,'
PPUA
-ScIP
CtrA (pM)
0 0.2 0.4 0.6 0.8 1.0 0
0.2
0A 0.6 0.8
[pee
1.0
0 0.2 0.4 0.6 0.8 1.0 0
0.2 0.4 0.6 0.8 1.0
D
POF
CtrA (pM)
0 0.2 OA 0.6 0.8 1.0 0
CtrA (pM)
Complexes
- ScIP
+ ScIP
-ScIP
+ ScIP
pM
POW
+ ScIP
SCIP -4+-.
0.2 0.4 0.6 0.8 1.0
-
-
- +4+
+
-
-
+
-
+
-+
-
+-+
- +4+
OKAA
DNA
60 bp
60 bp
25 bp
26 bp
PmIF
ScIP
-
ScIP
CtrA
-
CtrA
RNAP
S-
2.0
-
0.4 0.4
-
-
-
+ .
0.2 2.0
-
-
-
-
1.0O
+
+
0.2 0.8
1A
OA OA
0.4
0.4
+ .
+
+
RNAP-DNA
-OR
ICtrA4CP.ONA
##
CtrA-ONA
AAA&
VWWWWWW I*
-
DNA
Figure 2.10. Electrophoretic mobility shift assays with CtrA and SciP. CtrA-P binding to a
fragment of the (A) ccrM, (B) pilA, or (C) fiiF promoter containing a CtrA-binding site. SciP, if
present, was added at a final concentration of 1.0 RM. The concentration of CtrA in each lane is
indicated. (D) Binding of CtrA-P to the fliF promoter in the presence of wild type SciP or the
mutant indicated. CtrA was present at 1 [LM and SciP, or mutant SciP, at 1 [tM.
Discussion
As with all organisms, cell cycle progression in Caulobacter requires the just-in-time
transcription of large batteries of genes [28]. At the heart of this transcriptional program
is CtrA, which directly controls the expression of nearly 100 genes [20] and indirectly
many more. CtrA both activates and represses genes; for activated genes, CtrA binds near
the -35 site and probably recruits or binds RNA polymerase, while for repressed genes,
CtrA binds near the +1 site and presumably occludes RNA polymerase [16, 29]. A longstanding conundrum is why CtrA represses genes and the origin of replication in both
swarmer and predivisional cells, but activates most of its target genes only in
predivisional cells. Here, we identified SciP and showed that it temporally restricts the
transcriptional activity of CtrA by inhibiting CtrA-activated transcription in G1 swarmer
cells.
SciP plays a crucial role in regulating the activity of CtrA and, consequently, in
regulating the cell cycle and the establishment of asymmetric daughter cells in
Caulobacter. CtrA and SciP collaborate to create three distinct states corresponding to
the three major Caulobactercell types (Fig. 2.11). In GI swarmer cells, CtrA-P and SciP
levels are both high such that CtrA can repress genes and silence the origin of replication
but cannot activate gene expression. At the G1-S transition stalked cells are cleared of
CtrA and SciP thereby permitting DNA replication and the expression of CtrA-repressed
genes. As cells progress through S-phase, CtrA-P again accumulates to high levels but
SciP levels remain low, enabling CtrA to activate the expression of target genes and
propel cells through the ensuing cell division.
CtrA-P
O0
O0
G2
Sc
CtrA-activated gene expression
iP
CtrA-P
CtrA-P
CtrA-activated
genes
=
OFF
CtrA-P
CtrA-repressed
genes
CtrA-P
a
a
Figure 2.11 Regulation of CtrA-dependent gene expression during cell cycle progression.
CtrA transcriptional activity is controlled by temporally-regulated proteolysis, phosphorylation, and
protein-protein interaction with SciP. The timing of each regulatory event is shown beneath a
schematic of the Caulobacter cell cycle. These layers of regulation restrict CtrA-activated gene
expression to the predivisional stage. A model for how SciP and CtrA combine to regulate gene
expression during cell cycle progression is shown at the bottom.
Why block transcriptional activition by CtrA in GI cells? The majority of CtrA target
genes are involved in polar morphogenesis, including flagellar biogenesis, chemotaxis,
and pili biogenesis. CtrA also activates the expression of key cell division genes and
ccrM, an essential DNA methyltransferase. These processes - polar morphogenesis, cell
division, and DNA methylation - occur exclusively, or at least primarily, in predivisional
cells. Constitutive expression of many of these genes, such as ftsA and ccrM, has
deleterious consequences to cell cycle progression [30, 31]. SciP thus helps fulfill the
important task of preventing the execution of these cell cycle processes at the wrong
time.
Some CtrA-activated genes produce transcripts that are abundant in swarmer cells [20]
and seemingly exceptions to our model for SciP. However, DNA microarrays measure
steady-state mRNA levels and reflect both transcription and mRNA decay. Thus, some
genes may be transcribed in predivisional cells but yield mRNAs that are unstable until
the swarmer phase, as is the case for some flagellin filament genes [21]. Alternatively,
some genes could have promoter architectures that prevent SciP from inhibiting CtrA.
Regulation of SciP
Given the importance of SciP to cell cycle progression and the consequences of misexpression, it comes as no surprise that SciP itself is tightly regulated and restricted to
G1. sciP mRNA is detectable in predivisional and swarmer cells. However, SciP protein
only accumulates in swarmer cells, suggesting either SciP is rapidly degraded in
predivisional cells or the translation of sciP is blocked in predivisional cells and/or
activated in swarmer cells. The rapid disappearance of SciP at the G1-S transition
suggests it could be subject to temporally-regulated degradation. Notably, CtrA is rapidly
degraded by the ClpXP protease at precisely this stage of the cell cycle [32]. Consistent
with post-transcriptional regulation of sciP, cells constitutively expressing sciP from a
xylose-inducible promoter on the chromosome still only accumulated protein during Gi
(Fig. 2.4E). However, the regulation could be overcome by constitutively expressing sciP
from a high-copy number plasmid (Fig. 2.6C), and importantly, this overexpression was
sufficient to disrupt CtrA-dependent gene activation and, consequently, the cell cycle.
Our results outline a new feedback loop important for cell cycle progression in
Caulobacter.sciP appears to be a direct transcriptional target of CtrA as sciP expression
drops significantly in ctrAt" strains and the regulatory region upstream of sciP has a nearconsensus CtrA binding site. In predivisional cells, active CtrA likely drives the
expression of sciP. Following cell division SciP accumulates in swarmer cells, where it
feeds back to inhibit CtrA as a transcriptional activator. This feedback is reminiscent of
two other feedback loops, involving the cell cycle regulators RcdA and DivK. Both rcdA
and divK are direct transcriptional targets of CtrA [20] and each feeds back to inhibit
CtrA. RcdA somehow contributes to the regulated degradation of CtrA at the Gl-S
transition [33], although its function is unknown, while DivK inhibits the phosphorelays
that drive the phosphorylation and proteolytic stabilization of CtrA [34, 35]. The
combination of multiple feedback loops is likely critical to maintaining robust, sustained
cell cycling in Caulobacter.
Mechanism of inhibiting CtrA
We found no evidence that SciP affected the steady-state
levels, stability, or
phosphorylation of CtrA. Instead, our data suggest that SciP directly binds to and inhibits
CtrA. SciP does not, however, prevent CtrA from binding to DNA, consistent with our
finding that sciP mutations do not affect CtrA-repressed genes. The simplest model to
account for our data is that SciP blocks CtrA from binding to RNA polymerase and
activating transcription of target genes. The interaction of SciP and CtrA in the presence
of DNA was particularly striking and suggestive of a highly cooperative interaction. At
concentrations of CtrA that yield only minimal DNA-binding, the addition of SciP
stimulated very strong binding. The shifted bands in the presence of CtrA and SciP were
also sharper than with CtrA alone suggesting the ternary complex is more stable than the
CtrA-DNA complex. SciP did not bind DNA on its own, although it may bind DNA nonspecifically in the presence of CtrA. In fact, sequence analysis and the NMR structure of
a SciP ortholog (PDB: 2JRT) indicate some similarity to helix-turn-helix proteins (Fig
2.11 A-B).
Regulating response regulators
SciP represents a novel mechanism for modulating the output of a two-component
signaling pathway. Response regulators are usually regulated
at the level of
phosphorylation by cognate kinases and phosphatases. There are also regulatory proteins
that can bind and stabilize the aspartyl-phosphate linkage. For example, in Salmonella
enterica the protein PmrD binds the response regulator PmrA to block dephosphorylation
[36]. There are only a few examples of proteins that regulate response regulators without
affecting phosphorylation, such as the B. subtilis proteins RapC and RapG which prevent
ComA and DegU, respectively, from binding DNA [37, 38]. The E. coli protein TorI was
reported to inhibit transcriptional activation by the regulator TorR in vitro, but torI null
mutants were not shown to affect TorR-dependent gene expression [39] and TorI is not
expressed during exponential phase growth (M. Ansaldi and V. Mejean, personal
communication). The role of TorI is also unclear as recent data indicate TorI's primary
function is as a prophage excionase [40].
Perhaps the protein most analogous to SciP is Spx in B. subtilis. Spx is a small protein
that binds to the alpha-subunit of RNA polymerase at the same site contacted by the
response regulators ComA and ResD [41]. Spx thereby prevents these regulators from
binding RNA polymerase to stimulate the transcription of target genes. This may not be a
common regulatory strategy though as binding to RNAP polymerase runs an inherent risk
of having pleitropic effects on transcription.
SciP's inhibition of CtrA-dependent expression thus represents a novel mechanism for
regulating gene expression in bacteria. However, this mechanism may be prevalent
throughout the bacterial kingdom. First, SciP exhibits strict coconservation with CtrA and
has an ortholog in nearly every a-proteobacterial genome. More generally, SciP shows
moderate homology to helix-turn-helix proteins; many bacterial genomes encode for
dozens of small, uncharacterized helix-turn-helix proteins and we speculate that some of
them could inhibit other response regulators by a similar mechanism to SciP.
Concluding remarks
SciP's function underscores the notion that bacterial gene expression is often not dictated
simply by the activation of a single transcription factor, but instead involves the
combinatorial action of multiple regulators. The identification of SciP also highlights the
importance of regulated protein-protein interactions to cell cycle control in organisms
from bacteria to metazoans. As noted earlier, the master transcription factor in
metazoans, E2F, is regulated by the binding of Rb, while in budding yeast the cell cycle
transcription factors SBF and MBF are regulated through the binding of Whi5. Other key
cell cycle regulators, such as cyclin-dependent kinases, are also regulated by protein-
protein interactions [42, 43]. For instance, SICI in S. cerevisiae accumulates to maximal
levels in GI cells where it inhibits S-phase-specific CDKs while Rumlp and Rux play
similar roles in S. pombe and D. melanogaster, respectively. In mammalian cells, two
major classes of inhibitors, exemplified by p27 and pl6mK, directly inhibit different
classes of CDKs. Regulated protein-protein interactions involving master regulators are
thus critical to cell cycle progression at many levels and in all organisms.
Experimental Procedures
Bacterial strains and media
E. coli and C. crescentus strains were grown as previously described [44]. PYE was
supplemented
with 0.3% xylose or 0.2%
glucose when indicated. M2G was
supplemented with 0.3% xylose when indicated. Xylose inductions were performed by
diluting cultures grown to mid-log phase into PYE supplemented with xylose or adding
0.3% xylose to M2G. For SciP depletion, a culture of ML1750 was grown to mid-log
phase, pelleted at 6,800 g for 1 minute, washed with PYE, and resuspended in PYE plus
glucose. Synchronizations were performed on mid-log phase cells using Percoll (GE
Healthcare) density gradient centrifugation.
Cloning and mutagenesis
All expression vectors were constructed using the Gateway cloning system (Invitrogen)
as previously described [44]. The CC0903 (sciP) open reading frame was amplified using
the primers listed in Table 2 and cloned into pENTR-D-TOPO to create pENTR-sciP.
Alanine mutations were made using the QuikChange kit (Stratagene) with pENTR-sciP
as template. The pENTR-sciP plasmids were recombined with pHXM-DEST, pHIS-
DEST, pAD-DEST and pBD-DEST to construct the expression plasmids pHXM-sciP,
pHIS-sciP, pAD-sciP and pBD-sciP, respectively. Entry vectors for divJ (cytoplasmic
domain), divK, ctrA, and phoB were recombined with pAD-DEST and pBD-DEST.
pAD-DEST was constructed by ligating the Rfa cassette (Invitrogen) into pGAD-c1
linearized with SmaI. pBD-DEST was made by ligating Rfa into pGBDU-c3 that had
been digested with Clal and blunted with the End-It DNA End-Repair kit (Epicentre).
The orientation of the Rfa cassettes was verified by EcoRI digestion. pNPTS-spec-DEST
was constructed by inserting the Rfa cassette into the SmaI site of pNPTS138 followed
by replacement of the kanamycin resistance cassette (digestion with XbaI and NscI
followed by end repair) with the spectinomycin resistance cassette from HP45Q (excised
with SmaI).
The MultiSite Gateway vectors pEl-CC0903_LFR, pE3-CC0903_RFR, and pE2-FRT-tet
were constructed by amplifying a 600 bp region upstream of CC0903 (including 15
nucleotides into the coding sequence), a 600 bp region downstream of CC0903 (including
the last 30 nucleotides of coding sequence), and the tetracycline resistance cassette from
pKOC3 using the primers specified in Table 2 and cloning the PCR products into
pDONR221 Pl-P4, pDONR221 P3-P2, or pDONR221 P4r-P3r, respectively, by BP
recombination (Invitrogen) in a reaction consisting of 40 ng PCR product, 0.5 ptL
pDONR vector, and 1 pL of BP clonase in a 5.5 pL reaction volume. The BP reactions
were incubated overnight at room temperature and then transformed into chemically
competent TOP1O cells and plated on LB with kanamycin. The pKO-CC0903 plasmid
was then constructed with MultiSite Gateway Pro three fragment recombination between
pEl-CC0903_LFR, pE3-CC0903_RFR, pE2-FRT-tet, and pNTPS-DEST in a reaction
containing 40 ng of each entry plasmid, 100 ng of the destination vector, and 2 pLL of
Clonase II enzyme in 10 tL total reaction volume. The recombination reaction was
incubated at room temperature overnight and transformed into chemically competent
DH5ca and plated on LB supplemented with spectinomycin and tetracycline.
The integrating plasmid pX-sciP was created by excising GFP from pXGFPN-2 with
NdeI and KpnI to replace the GFP cassette with the full sciP coding sequence derived by
digesting a PCR fragment amplified with primers harboring NdeI and KpnI restriction
sites as indicated in Table 2.
Deletion and depletion of sciP
SciP was deleted through a two-step homologous recombination method previously
described [44] using the pKO-CC0903 integrating plasmid. The sciP deletion was
verified by PCR using one primer outside the region used for homologous recombination
and one primer in the tetracycline resistance cassette (Table 2). The tetracycline-marked
deletion was transduced into a clean CB15N background using $CR30.
To generate a depletion strain, the integrating plasmid pX-sciP was transformed into
CB 15N by electroporation and integrants of sciP at the xylX locus were selected on PYE
plates containing kanamycin and glucose. The tetracycline-marked sciP deletion was then
transduced from ML1749. The depletion strain was selected on PYE plates containing
kanamycin, oxytetracycline, and xylose and verified by PCR to contain sciP only at the
xylX locus.
Protein purification and antibody production
His6-SciP was expressed from pHIS-sciP plasmid and purified, along with His 6-CtrA, as
described previously [44]. His6-RpoD was expressed from pHIS-rpoD and purified as
with His6-SciP and His6-CtrA but with the addition of 6 M GuHCl to the lysis buffer.
RpoD was bound to Ni-NTA resin under denaturing conditions and subsequently
renatured by washing in wash buffer with no GuHCl. Refolding was confirmed by CD
spectroscopy. TAP-tagged RNA polymerase was purified from ML1799 as previously
described [45] and dialyzed overnight at 4'C against 500 volumes of TGED buffer
(10mM Tris-HCL pH 7.9, 50% glycerol, 0.1 mM EDTA, 0.1 mM DTT, 500 mM NaCl).
Coomassie staining was used to verify that purified TAP-tagged RNA polymerase
contained alpha, beta, beta', and sigma subunits. Rabbit polyclonal antisera were
generated from purified His6-SciP (Covance). Rabbit polyclonal antisera were generated
from purified His6 -SciP (Covance).
Immunoblots
Cell pellets were collected at indicated time points. Cells were resuspended in 1x SDS
sample buffer and resolved on 15 %,Tris-HCl gels (Bio-Rad) run at 150 V for 1 hr at
room temperature, transferred to PVDF membrane and probed with a 1:2,000 dilution of
SciP antisera, 1:1,000 anti-M2 affinity-purified antibody (Sigma), or 1:10,000 of CtrA
antisera in lx TBS plus 0.05% Tween-20.
Microscopy
Cells were fixed with 0.5% paraformaldehyde, centrifuged at 6,800 g, washed with PBS
and resuspended in a small volume of PBS. Differential interference contrast images
were taken with a Zeiss Axiovert 200 microscope with a Zeiss aPlan-Fluar 100x/1.45 oil
objective and an Orca II camera (Hamatsu) controlled by MetaMorph 7.1.3.0 software.
Flow cytometry
Samples for flow cytometry were prepared and analyzed as described previously [35].
DNA microarray analysis
Cultures were grown in to mid-log phase, harvested by centrifugation at 20,000 g, and
frozen in liquid nitrogen. For the sciP depletion experiment, ML1750 was grown to midlog phase in PYE supplemented with xylose, synchronized and released into fresh PYE
containing either glucose or xylose. Cell pellets were collected after 100 minutes,
immediately after cell division in each culture. RNA was isolated using the RNeasy kit
(Qiagen). 30 [tg of total RNA was directly labeled using the Superscript-II kit
(Invitrogen) and either Cy3- or Cy5-dCTP. RNA template was degraded with NaOH.
Cy3- and Cy5-labeled cDNA samples for comparison were mixed and unincorporated
nucleotides removed using the QIAquick PCR purification kit (Qiagen). Labeled cDNAs
were hybridized to Agilent arrays, arrays scanned using an Agilent scanner, and data
extracted using the Agilent feature extraction scripts within the Matlab Bioinformatics
Toolbox. Analysis of the depletion and overexpression strains were done in duplicate and
triplicate, respectively, and results averaged in each case.
Yeast two-hybrid assays
Protein-protein interactions were assayed in the yeast two-hybrid system described
previously [46] except using the Gateway modified pAD-DEST and pBD-DEST vectors.
S. cerevisiae PJ69-4A was co-transformed with combinations of pAD and pBD vectors
using LiAc. Double transformants were selected on solid CSM-Leu-Ura. Yeast strains
were grown at 30 0C in liquid SD minimal media supplemented with 2% glucose and
CSM-Leu-Ura or CSM-His-Leu-Ura plus 2% bactoagar for solid media.
Electrophoretic mobility shift assays
EMSAs were performed using His 6 -SciP and His 6-CtrA purified protein. 50 base pair
PAGE purified primers (Sigma-Genosys) were annealed in TEN buffer (10 mM Tris-HCl
(pH 8.0), 1 mM EDTA, and 50 mM NaCl) at 95*C for 10 minutes and slowly cooled to
room temperature. The double-stranded DNA was labeled with [y32P]-ATP using T4
polynucleotide kinase (NEB). 10 liM CtrA was phosphorylated using 1jxM MBP-EnvZ in
HKEG buffer (10 mM HEPES-KOH [pH 8.0], 50 mM KCl, 10% glycerol, 0.1 mM
EDTA, 1 mM DTT) plus 5 mM MgCl2 and 1 mM ATP for 20 min at 37 C. The CtrA-P
was diluted into HKEG plus MgCl 2 to
lOX before
being added to the gel shift reactions.
Proteins were pre-incubated at room temperature for 30 min in gel shift buffer (50 mM
KCl, 5mM MgCl 2 , 20 mM Tris HCl (pH 8.0), 100 [M EDTA, 1 mM DTT, 1 mg/ml
BSA, 10% glycerol, 100 ng/ml competitor DNA (Poly (I)-Poly(C))) and then incubated
reaction volume. 2
with 3.5 nM labeled DNA at room temperature for 30 min in a 18 1A
tl of 0.2% bromophenol buffer was added to each reaction immediately before the
samples were loaded onto a Novex 6% DNA retardation gel (Invitrogen) and run at 300
V in chilled 0.5x TBE buffer for approximately 17 min or until the dye front migrated
sufficiently far through the gel. The gel was then transferred to Whatman paper, covered
on top with Saran wrap, dried on a gel drier for 1 hr, and exposed to a phosphor screen
for 1 hour. Screens were scanned with a Typhoon scanner.
Chromatin immunoprecipitation
Bacterial ChIP was performed as described previously using 2.5 ptL of polyclonal c-CtrA
antibody for each pull-down [47]. The % input values for the ChIP samples were
calculated using the average Ct values from triplicate measurements and a standard curve
generated from five two-fold serial dilutions of the input samples. Each ChIP experiment
was performed on three independent colonies to calculate standard error bars. The %
input values for the ChIP samples were calculated using the average Ct values from
triplicate measurements and a standard curve generated from five two-fold serial
dilutions of the input samples. Each ChIP experiment was performed on three
independent colonies to calculate standard error bars. The relative expression of genes
was calculated using the following formula: ratio = ((Etarget)^ACPtarget(control-sample))/
((Erho)AACPrho(control-sample)). For expression analysis, 1 [tg of total RNA was reverse
transcribed into cDNA using Superscript-II, RNA template degraded was with RNAse H,
and nucleotides were removed using the QIAquick PCR purification kit (Qiagen).
Quantitative real-time PCR
Quantitative real-time PCR for ChIP and expression analyses was performed using the
DNA Engine Opticon 2 system (MJ Research) and the QuantiFast SYBR Green PCR Kit
(Qiagen). Each reaction contained 2.5% of total ChIP sample or 2% total cDNA for
expression analysis, 10 pmol of forward and reverse primer, lx Quantifast SYBR Green
PCR master mix (Qiagen) in a 25 pl reaction. Samples were amplified by incubating at
504C for 2 min, 95'C for 15 min followed by 45 cycles of 95'C for 15 sec, and 60'C for
30 sec followed by fluorescence data collection. Cycle threshold (Ct) values were
calculated using Opticon Monitor software (MJ Research). The relative expression of
genes was calculated using the following formula: ratio = ((Etarget)AACPtarget(controlsample))/ ((Erho)AACPrho(control-sample)). For expression analysis, 1 pg of total RNA
was reverse transcribed into cDNA using Superscript-II, RNA template degraded was
with RNAse H, and nucleotides were removed using the QlAquick PCR purification kit
(Qiagen).
Quantitative real-time PCR was performed using the DNA Engine Opticon 2 system (MJ
Research) and the QuantiFast SYBR Green PCR Kit (Qiagen). Each reaction contained
2.5% of total ChIP sample or 2% total cDNA for expression analysis, 10 pmol of forward
and reverse primer, lx Quantifast SYBR Green PCR master mix (Qiagen) in a 25 I
reaction. Samples were amplified by incubating at 50'C for 2 min, 95'C for 15 min
followed by 45 cycles of 95*C for 15 sec, and 60*C for 30 sec followed by fluorescence
data collection. Cycle threshold (Ct) values were calculated using Opticon Monitor
software (MJ Research).
Pulse-chase analyses
CtrA stability was assayed as described previously [8]. Cultures grown in M2G were and
for ML1748, pre-induced with xylose for one hour and labeled with [ 35S]-methionine and
chased with 1 mM methionine and 0.3% casmino acids. I mL aliquots were removed at
indicated time points, cells were pelleted by centrifugation at 20,000 x g, and frozen in
liquid nitrogen. Cell pellets where resuspended in 50 pL of lysis buffer (10 mM Tris pH
8, 1% SDS, 1mM EDTA), boiled for 2 minutes to lyse, and diluted with 800 pL chilled
IP buffer (50 mM Tris pH 8, 150 mM NaCl, 0.5% Triton X-100). Samples were precleared with 20 pL Pansorbin Cells (Calbiochem) immunoprecipitated with 1.5 pL CtrA
antibody at 4 C over night, and immune complexes were collected with 30 pL protein A
beads (Invitrogen). The samples were washed 3x with chilled IP buffer and eluted in 30
pL of 2x SDS sample buffer by boiling for 5 min. The samples were loaded onto 10%
Tris-HCl gels (Bio-Rad) and run at 150 V for 40 min at room temperature.
Gels
transferred to Whatman paper, dried for 1 hr and exposed to a phosphor screen for 72
hours.
In vitro phosphorelay dynamics
In vitro phosphorylation and phosphorelay reversal assays were done as previously
reported [48].
In vivo phosphorylation
Analysis of CtrA phosphorylation in vivo was performed essentially as described
previously [8]. Briefly, one colony was inoculated into M5G medium containing 0.05
mM phosphate and grown overnight at 30'C. The cultures were grown at 30'C until the
optical density of the culture measured at 660 nm was in the range 0.1 to 0.2. Cultures
were then split with one maintained in glucose and the other grown in xylose for 2 hours.
Equal numbers of cells (based on OD) were labeled for 5 minutes with 30 pCi/mL [y32 P]ATP
before immunoprecipitating CtrA.
Table S1 - Strains and Plasmids
Strain or Organism or Plasmid
Plasmild
Category
Strain
Plasmid
Strain or Plasmid Name
C. crescentus
CB15N
ML1748
ML1749
ML1750
E. coli
DH5a
BL21-Tuner
One Shot ccdBSurvival Ti
TOP10
PJ69-4A
S. cerevisiee
pHXM-DEST
destination vectors
pHIS-DEST
pNPTS-spec-DEST
pAD-DEST
pBD-DEST
pHXM-sciP
expression vectors
pHXM-sciP(R35A)
pHXM-sdP(R40A)
pHXM-sciP(E57A)
pHXM-sdP
pHXM-sciP(R35A)
pHXM-sciP(R40A)
pHXM-sdP(E57A)
pENTR/D-TOPO
vectors
general purpose
pKOC3
pNPTS138
HP45Q
P1-P4
pDONR221
P3-P2
pDONR221
P4r-P3r
pDONR221
pXGFPN-2
pGBDU-c3
pGAD-cl
pEl-CC0903_LFR
donor vectors
pE3-CC0903_RFR
pE2-FRT-tet
pENTR-sciP
entry vectors
pENTR-sciP(R35A)
pENTR-sciP(R40A)
pENTR-sciP(E57A)
pENTR-phoB
pENTR-divK
pENTR-divJ
pENTR-ctrA
pKO-CC0903
plasmids
Integrating
pX-sclP
vectors pAD-phoB
yeast two hybrid
pBD-phoS
pAD-divK
pBD-divJ
pAD-ctrA
pBD-crA
pBD-sciP
pBD-sciP(R35A)
pBD-sciP(R40A)
pBD-sciP(E57A)
Table 2.1. Strains list.
Descprtion
Source or Reference
Evinger and Agabian, 1977
this study
this study
this study
Invitrogen
Novagen
Invitrogen
Invitrogen
James at a, 1996
Skerker at al, 2005
Biondi et al, 2005
this study
this study
this study
this study
pJS71X-M2-sciP (specp)
this study
pJS71-M2-sciP(R35A)(specR)
this study
pJS71-M2-sc/P(R40A)(spec")
this study
pJS71-M2-sc/P(E57A)(spec")
this study
pET-His.-sdP (amp")
this study
pET-His-sciP(R35A) (ampR)
this study
pET-Hise-sciP(R40A)
(amp")
this
study
pET-His-srdP(E57A) (ampR)
Invitrogen
ENTRYvector forGateway cloning system (kan")
integrating vector with FRTflanked tetracycline resistance cassette (chlorR) Hamilton at al, 1989
lab collection
integration vector with sacB (kan")
lab collection
source of omega cassette (spec")
Invitrogen
DONR vector forGateway multisite cloning system
invitrogen
DONR vector forGateway multisite cloning system
invitrogen
DONR vector forGateway multisitecloning system
Thanbichler at al, 2007
integrating plasmld(kan")
James at al, 1996
for fusions to C-terminaldomainof GAL4binding domain (LEU2,amp')
James at al, 1996
for fusions to C-terminaldomainof GAL4activating domain (URA3,amp")
this study
600 base pairs upstreamof CC0903in pDONR221
P1-P4 (kanR)
this study
600 base pairs downstream
of CC0903in pDONR221
P3-P2 (kan")
this
study
pDONR221
P4r-P3rwithFRT flanked tetracycline resistance cassette (kanR)
this study
CC0903 in pENTR/D-TOPO
(kane)
this study
(kan")
CC0903(R35A) InpENTR/D-TOPO
this study
CC0903(R40A) InpENTR/D-TOPO
(kanR)
this study
CC0903(E57A) in pENTR/D-TOPO
(kan")
Skerker et al, 2005
CC0294 in pENTR/D-TOPO
(kan")
0
Skerker et al. 2005
(kan )
CC2463 in pENTR/D-TOPO
Skerker et al, 2005
(kan")
CC1067 (cytoplasmic domain) in pENTR/D-TOPO
0
Skerker at al. 2005
CC3035 in pENTRID-TOPO
(kan )
this study
In-frame deletion construct for sciP (tet")
Thanbichler et al, 2007
integrating plasmidfor sciP at Pxyl (kan")
this study
(carbR,LE2)
pGAD-phoB
this study
pGSDU-phoB (carb", URA3)
this study
pGAD-divK (carbR, LEU2)
this study
pGBDU-divJ (carb",URA3)
this study
pGAD-crA (carb, LEU2)
this study
pGBDU-ctrA (carbR,URA3)
this study
pGBDU-sciP (carb", URA3)
this study
pGBDU-sciP(R35A) (carb", URA3)
this study
pGBDU-sciP (R40A)(carb", URA3I
this study
pGBDU-sciP (E57A)(carbR,URA3)
Synchronizable derivative of wild-type CB15
CB15N+ pHXM-sciP (spec")
CB15NAsciP (tet")
0
CB15N AscP xyl::sciP(te", kan )
General cloning strain
Strain for protein expression andpurification
of destination vectors
Strain for propagation
clones
General cloning strain for pENTR/D-TOPO
GAL7-IacZ
strain for yeast two-hybrid with GALI-HIS3, GAL2-ADE2,
pJS71X-M2;
high-copy, P. M2tag (spece,chlore)
pET-His.(amp')
pNPTS1
38 derivativemodifiedwith Rfacassette (spec")
pGAD-CIGa4 AD fusion plasmid (carb", LEU2)
Gal4 BD fusion plasmid (carb", URA3)
pGBDU-C3
Table S2 -
Primem
R-
FO-Nd
ENM pft.1ft
pENM-F
$ft OW.Md
Mutw
CACCTTGTTGCAGCACCAGCOCAC
OGG"GTCACATCAACTCA
pd-
l.
GCGGAGACCCAGQCrrGGGrrAT=T
ACG"TAAC4CX
GCCTGWTCT=GC
PENM-.ORR40A)
TGC4"ATCCGTGCGMaGCCGAWTC
GACCTMGCCTrCOCACGOATAACCCA
PENM-F(EVA)
CTMOCTCQACGCG=TGCGATMT
pENM--FYRW)
ACOATCGCACG=CG=AGCGAW
canobvft
DoUtion
PEI-CCOW3
R
pE3-CCODOM
_LHR
OC0903
KO cw*
PE2.FRT4@1
.ftgr.u.d=p..YDr
bon
TTACTGACOCAGTACAACA=
GOGGACAAGTMTACAAAAAAGCAGr
CATGr.C
r4C.ACAACrTTGTATAGAAAAGTTCG.TGCTGCTGCTGC
GOGOACAACTrrOTATAATAAAGTrOCTGGAOCACCCGGATCCAOC
G<VX2ACCACTrn3TACAAGAAAGCTGGGTACCGArGGAGCCOGAOCYCACOAG
GACOCCCTGOOCATCOAC
GAArGrrQAAGM00GAr.A
GOMACMCMA7TATA'TTGTGAAGTTCCTATC
CQCGACAACTTTTCTATACAAA(MGCTAAGAAG"CCTATACTTTC'rAGAGAATAGGAACTTCTCMTOOTCAM-.MCTOGGTCC
TCCAGG-
WTCCAG
CAGCGATCGGCTOGTTGCO
GGGCGGTCAC;ATCMCTCA
PCM
P
(hWW.
K -Q
GOTACCOMAArOGTACTOCTOGA _C
AG=GGOGAGACOACCATATGiTTOCAOCAOCAGCOCAC
up
Ar-W
ACTCGCACAGCCCGTCCCGG
TCCCCAAGCCTGCGCCGCAT
MIPpXA
GCCAGCATCACM)C:-MGG
CGACTGCACTrAATGGCCAG
3'r--
GTCAACGGCGAGCTAGACCT
CAGTCrCCGTGGATGATGGT
q RT4CR
MW
PAP
CCGATCCGCCCrATAOLTCTG
CCTACCGGCAAGCTGATra
GITGTGATAGCTGCCGATCA
AACrGGACCAGGMCGCCAC
w shift
.w
CTCAAAArCOCCTGAMQGCCrTGGTTAACGGOCCGCTAACCACGT=T
AGACs
OGTGGTrAGCGGOCCGTTAACCACrGC4TrTrAGGCOCTtTTrAG
pAA
'rTOWAGCGATCGC4"OTGCATOGTTAAGAACAAATAACOGTAAATACA
TOTATTTACZGTTATTTOTTCT'rAACCATOCACTCCGCGATCOCTOCCAA
W
GGAAAOCCrGACOATCGGCAGATATAAACOCCIrGTTTACCTrGTACTOG
CCAGTACAAGGTAAACOAC-3rGMATATCTGWGATCGTCAWrMCC
MM
AGGTCOCOGAACCACGATGCGAGOAAAOrACGATGGGTAOCGAAACCGTGCACGGTrTrGGTACCCATOOTOGMOCTCGrAYCGTOGTTCGMaACCT
Table 2.2. Primers.
Acknowledgements
We thank J. Modell, K. Milaninia, and B. Pando for help with experimental techniques.
We thank F. Solomon, M. Neiditch, A. Fan, A. Grossman, and members of the Laub lab
for comments on the manuscript. M.T.L. is an Early Career Scientist at the Howard
Hughes Medical Institute. This work was supported by an NIH grant (5R01GM082899)
to M.T.L.
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Chapter 3
The Caulobacter cell cycle regulator SciP is proteolyzed at the
G1-to-S transition by Lon
This work is a manuscript in progress by Kasia G Gora, Matt Wohlever, Barrett S
Perchuk, and Michael T Laub.
K.G.G and M.T.L designed the experiments, analyzed data, and wrote the manuscript.
K.G. performed all the experiments with the exception of in vitro degradtion performed
by M.W and microarrays perfomed by B.S.P.
Introduction
In the previous chapter I described the identification and characterization of SciP, a new
cell cycle regulator in Caulobacter crescentus. I demonstrated that SciP prevents the
activation of CtrA-dependent genes through a direct interaction with CtrA.
SciP
accumulates to maximal levels in G1-phased swarmer cells, is rapidly eliminated during
the Gl-S transition, and then begins accumulating again shortly before cell division. The
rapid elimination of SciP during the G1-S transition is reminiscent of the rapid
degradation of CtrA by the protease ClpXP during the same cell cycle [1] In this chapter
I examine the regulation of SciP and demonstrate that (1) cell cycle changes in SciP
abundance involve regulated proteolysis by Lon and (2) an inability to degrade SciP
disrupts normal cell cycle progression.
Lon in an AAA+ protease whose homologs are widely distributed in both eukaryotic and
prokaryotic organisms [2]. Like other bacterial AAA+ proteases, Lon is functional as a
hexamer but unlike ClpXP, Lon consists of an ATPase regulatory domain fused to a
proteolytic domain [3]. In bacteria, Lon important for the proteolysis of unfolded proteins
through the recognition exposed hydrophobic residues in unfolded structures in addition
to other native key regulators [4]. In Caulobacter Lon is a non-essential protein and has
been shown to degrade the cell cycle methyltransferase CcrM in late predivisional cells
[5]. Here we demonstrate that Lon also plays a key role in the degradation of SciP
specifically at the G 1-to-S transition.
Results
SciP is regulated primarily at a post-transcriptional level
Consistent with its proposed role as a swarmer-specific inhibitor of CtrA-activated gene
expression, SciP abundance is maximal in swarmer cells and then drops rapidly during
the G1-to-S transition [6]. This pattern suggests that cells may tightly regulate SciP
levels.
The transcription of sciP depends on CtrA and is consequently strongly cell cycleregulated, peaking in predivisional cells [6, 7]. To test whether regulated transcription
contributes to the cell cycle changes in SciP, we measured SciP abundance in a strain
harboring a deletion of the native sciP and expressing sciP from the chromosomal xylX
locus, i.e. the sciP depletion strain [6]. Growth in the presence of xylose leads to
constitutive expression from the PsY promoter [8]. Synchronized swarmer cells from this
strain were isolated and released into rich media containing xylose or glucose. Consistent
with previous results,
SciP levels were high in swarmer cells, but decreased
approximately 30 minutes post-synchronization, despite the constitutive expression of
sciP (Fig. 3.1) [6]. Immunoblotting for CtrA in the same samples indicated that the
disappearance of SciP was coincident with the proteolysis of CtrA (Fig. 3.1). These
results suggested that changes in SciP abundance may be driven by proteolysis during the
G1-to-S transition in addition to changes in transcription.
minutes post
0
15
30
45
60
76
90 105
synchrony
y
4
CtrA
SciP
PxyrsciP
OAW
CtrA
SSciP
Figure 3.1 Post-transcriptional regulation of SciP. Cell cycle abundance of SciP under
constitutive expression. The sciP depletion strain was grown in PYE plus xylose, synchronized,
and released into PYE plus xylose or glucose as indicated. Samples were taken at 15-minute
intervals for Western blot analysis with CtrA and SciP antisera.
SciP is an unstable protein, but is not degraded by CIpXP, CIpAP, or HslUV
To measure the stability of SciP we used pulse-chase analysis on wild-type cells
expressing sciP from a low-copy plasmid and grown in a minimal medium, M2G. Under
these conditions in which the cell cycle is -140 minutes, SciP had a half-life of only -23
minutes indicating it is a relatively unstable protein (Fig. 3.2A). We therefore sought to
identify the protease(s) responsible for SciP degradation. Because SciP normally
disappears during the Gi -to-S transition, concurrent with the proteolysis of CtrA, we first
investigated ClpXP, the primary CtrA protease [1]. As both ClpP and ClpX are essential
proteins, we assayed SciP levels in cells depleted of ClpP [1]. Strain UJ199 harbors a
deletion of the native, chromosomal clpP gene with a single copy of clpP under control
of the P,,/ promoter at the xylX locus [1]. This strain was grown in rich medium
containing glucose for four generations (-6 hours) to deplete ClpP. We then isolated
swarmer cells, released them into glucose or xylose to repress or induce, respectively,
c/pP expression, and sampled cells every 30 minutes for immunoblot analysis (Fig.
3.2B). As expected, cells released into glucose failed to proteolyze CtrA, whereas cells
released into xylose eliminated CtrA after -60 minutes. In contrast to CtrA, the same
samples, from both xylose and glucose, showed a nearly complete elimination of SciP by
the 60-minute time point. These data indicate that cells lacking ClpP can still degrade
SciP during the Gl-to-S transition. Because ClpP is the peptidase subunit for both ClpXP
and the protease ClpAP, these data argue against the involvement of either protease in
SciP proteolysis.
To confirm these results, we also monitored SciP abundance in synchronized Ac/pA and
AcpdR strains [9, 10] ClpA is the ATPase subunit of ClpAP and is not essential for
viability [10]. CpdR is a non-essential protein that is required for the ClpXP-dependent
degradation of CtrA at the G1-to-S transition [9]. In AclpA cells, SciP was eliminated
during the GI-to-S transition, as in wild-type cells (Fig 3.2C). In AcpdR cells, SciP levels
dropped significantly in the middle of the cell cycle though not with the same timing as in
wild-type cells (Fig 3.2C). This delay may result from the fact that AcpdR has a longer
cell cycle at least in part because these cells fail to clear CtrA at the Gl-to-S transition
[9]. Alternatively, ClpXP may make a minor contribution to SciP proteolysis that was not
detected in our analysis of the ClpP depletion strain.
We also tested the role of HslUV, another ATP-dependent protease [11]. We
synchronized a strain harboring a transposon within the hslV coding region, hslV::Tn5,
and monitored SciP levels by Western Blot. However, SciP was still eliminated during
the Gi -to-S transition, indicating that HslUV does not degrade SciP (Fig 3.2D).
0
100%1
5 ScIP In Alon
minutes post
synchrony
0
30
60
90
120
*ScP-M2
1
10%
CtrA
* M2-SciP
ASCIP
Pxyr.CIpP
4
CtrA
0%
0
30
60
Time Post Pulse
90
SciP
-
C.
D.
minutes post
synchrony
0
30
60
90
120
minutes post
synchrony
SclP
AcpdR
0
30
90
120
SclP
hs+n
*%gyp
4
AcipA[egas
60
4
CtrA
SciP
E.
minutes post
synchrony
0
30
60
1.-14
90
120
SciP
Alanr11
1 CtrA
Figure 3.2 SciP stability in various protease mutant strains. (A) Pulse-chase analysis of SciP
stability in the strains indicated. Wild type or Dion cells harboring the indicated allele of SciP on a
low copy plasmid grown in M2G and pre-induced with xylose for 30 minutes before the pulse
chase. Each experiment was performed in duplicate or triplicate and error bars indicated standard
error of the mean. (B) The c/pP depletion strain was grown in PYE plus xylose and switched to
PYE plus glucose to deplete SciP for 6 hrs. Cells were synchronized and released into PYE plus
glucose or xylose as indicated and samples were taken at 15-minute intervals for Western blot
analysis with CtrA and SciP antisera. The Ac/pA, AcpdR (C), hsIV::tn5 (D), Alon strains (E) were
grown in PYE, synchronized, and analyzed as in (B).
SciP is degraded by Lon
We also tested SciP proteolysis in cells harboring a disruption of the gene encoding the
Lon protease [5]. We synchronized Alon cells and monitored both SciP and CtrA levels
during cell cycle progression. Whereas CtrA disappeared, as expected, at the 30 minute
time point, SciP was still present after 30 minutes and was maintained at significant
levels throughout the cell cycle (Fig. 3.2E) We also measured SciP stability in a mixed
population of Alon cells using pulse-chase analysis. We found that wild-type SciP protein
had a half-life of -230 minutes in the Alon mutant compared to just 23 minutes in the
wild type (Fig. 3.2A) Taken together, these data strongly suggest that SciP is degraded by
Lon.
Next we tested the ability of purified E. coli Lon to proteolyze SciP in vitro. We
incubated 10 [M SciP with 500 nM E. coli Lon hexamer and observed rapid and almost
complete degradation within 30 minutes at 30 C (Fig. 3.3). Given the high degree of
sequence similarity between E. coli and Caulobacter Lon (62% identical), these data
further support the conclusion that SciP is a direct substrate for Lon in Caulobacter.
reaction
time (min) 0
10
20
30
all-e
+Lon
*-Pyruvate kinase
4-SciP
Degradation
Products
Figure 3.3 In vitro proteolysis of SciP by E. col Lon. 10 [tM SciP was incubated with 500 nM
Lon hexamers from E. coli and degraded for 30 min at 30C. Samples were taken every 10 min for
SDS-PAGE gel analysis. Pyruvate kinase was added to generate ATP from phosphoenolpyruvate
present in the reaction.
Addition of a C-terminal tag interferes with SciPproteolysis
To test the importance of regulated SciP degradation to cell cycle progression, we sought
to create a proteolytically stable version of SciP. Because proteases often rely on N- or Cterminal sequences for substrate recognition, we tested the effects of placing an M2
epitope at the N- or C-terminus of SciP. We transformed wild-type cells with low copy
plasmids expressing sciP, M2-sciP, or sciP-M2 under the control of Pxyl.
Mixed
populations of cells were grown in rich media supplemented with glucose to repress the
plasmid-borne copy of sciP, synchronized, and released into medium supplemented with
xylose to induce sciP, M2-sciP, or sciP-M2 expression. Samples were collected every 15
minutes for immunoblot analysis (Fig. 3.4). Note that we did not induce expression of
the plasmid-borne copies of sciP prior to synchronization as such pre-induction
significantly decreased the yield of swarmer cells for the sciP-M2 strain during
synchronization.
0
minutes post
synchrony
PYrsciP
15
30
45
60
75
90 105 120
j
SciP
4
PxyrM2 -sciP
_________________________4
PxyrsciP-M2_4
M2-SciP
SciP
ScIP-M2
Figure 3.4 Stabilization of SciP through an C-terminal M2 tag. Wild-type cells harboring a low
copy plasmid with the indicated sciP allele were grown in PYE plus glucose, synchronized, and
released into PYE plus xylose to induce sciP expression at time 0. Samples were taken every 15
minutes for Western blot analysis with SciP antisera.
For cells synthesizing untagged SciP, we found that SciP was high in swarmer cells,
almost completely eliminated after ~30-45 minutes, and then accumulated again during
later stages of the cell cycle (Fig. 3.4), a pattern similar to wild-type and cells in which
the only copy of sciP is expressed from the chromosomal xylX locus (see Fig. 3.1). By
contrast, for cells expressing M2-sciP, we found that M2-SciP protein accumulated
during the early stages of the cell cycle as significantly higher levels of M2-SciP were
seen at the 30 and 45 minute time points compared to untagged SciP (Fig. 3.4). An even
more pronounced effect was seen in the cells expressing sciP-M2, with high levels of
SciP-M2 detectable throughout the time course examined (Fig. 3.4). Consistently, the
SciP-M2 that accumulated blocked expression of the chromosomally-encoded sciP, a
CtrA-activated gene, during late stages of the cell cycle; as the native SciP is slightly
smaller than SciP-M2 the two forms were distinguishable on immunoblots using SciPspecific antibodies (Fig. 3.4). These results suggest that the C-terminus, and to a lesser
extent the N-terminus, of SciP is important for the degradation of SciP. The presence of a
C-terminal M2 epitope likely stabilizes SciP by interfering with its proteolysis.
We also assayed the stability of M2-SciP and SciP-M2 using pulse-chase analyses of
mixed populations of cells grown in minimal medium. In cells producing M2-SciP or
SciP-M2, we measured a protein half-life of 36 and 58 minutes, respectively, compared
to 23 minutes for the untagged protein (fig. 3.2A). These results further indicate that
adding a C-terminal tag to SciP interferes with its degradation.
Proteolysis of SciP is necessary for proper cell cycle progression
To investigate the phenotypic consequences of producing stable SciP, we expressed sciPM2 from a low-copy plasmid under the control of a xylose-inducible promoter in
otherwise wild-type cells. This strain was grown in rich medium supplemented with
glucose and then shifted to medium containing xylose, leading to a significant growth
defect (Fig. 3.5A). After four hours, cells were also moderately filamentous and flow
cytometry analysis indicated that cells had accumulated up to four chromosomes per cell,
consistent with a defect in cell division (Fig. 3.5B).
In contrast to sciP-M2, cells
expressing untagged sciP or M2-sciP from the same low-copy plasmid exhibited no
significant defects in growth, morphology, or chromosome content (data not shown). The
phenotypic consequences of producing SciP-M2 from a low-copy plasmid are
reminiscent of those previously reported for cells producing M2-SciP from the same
xylose-inducible promoter on a high-copy plasmid.
100
PxytsciP-M2
1pxyj scIP-M2
.8
glucose
0
-*-glucose
-O-xylose
1.4-
1.2
0
0
90
180
270
360
45(
lime (minutes)
1
2 3 4 5 6
70
15
45
1 2 3 4 5 6 7 8
1J
A ReF
ICtri
mprud
goe*
minutes post
synchrony
0
F
30
60
a
I
75
90
105 120
4 ScIP- M2
*~
~
+CtrA
PxytsciP-M2
CtrA-sctvh1
gal-
-1
0.5
0
0.5
4 SciP
L
ctrA
1
Figure 3.5 Phenotypic consequences of stable SciP. (A) Growth curve of wildtype harboring
PxyrsciP-M2 on a low copy plasmid grown in PYE supplemented with glucose or xylose. (B)
Morphology (top) and FACS analysis (bottom) of PxyrsciP-M2 induction for 4 hrs. (C) Microarray
analysis of CtrA regulon in cells expressing SciP-M2 for 4 hrs vs. wildtype grown in similar
conditions. Changes in gene expression are compared to previously published data sets in
reference [6]. (D) CtrA dynamics in synchronized wild type cells harboring PxyrsciP-M2 grown in
PYE plus glucose, synchronized, and released into PYE plus glucose or xylose as indicated.
Samples were taken every 15 minutes for Western blot analysis with SciP or CtrA antisera.
To further compare the phenotypes that result from synthesizing the non-degradable
SciP-M2 from a low-copy plasmid or M2-SciP from a high-copy plasmid, we examined
changes in gene expression in these strains using DNA microarrays. Wild-type cells
harboring the low-copy sciP-M2 expression plasmid were induced with xylose for 4
hours; mRNA was isolated from these cells and compared to mRNA from identically
101
treated wild-type cells using whole genome microarrays (Fig. 3.5C). We then directly
compared the results to those obtained for cells expressing sciP from a high-copy plasmid
and to data for ctrA's and cckA" strains grown at a restrictive temperature [6]. The lowcopy expression of sciP-M2 gave rise to an -3-fold increase in sciP mRNA relative to
wild-type cells while high-copy expression of sciP resulted in an -18-fold increase.
Despite the lower induction of sciP-M2, the consequent down-regulation of CtrA target
genes was very similar to that seen in cells expressing M2-sciP from a high-copy plasmid
(Fig. 3.5C). In both cases, CtrA-repressed genes were generally unaffected.
To determine whether the down regulation of CtrA-activated genes was due simply to
changes in CtrA levels, we measured CtrA abundance in synchronized cells expressing
sciP-M2 from a low copy plasmid. Cells were grown initially in the presence of glucose,
synchronized, and then released into media containing xylose or glucose, and sampled
every 15 minutes for immunoblot analysis. In both conditions CtrA levels fluctuated as in
wild type cells, indicating that the accumulation of SciP-M2 likely down regulates CtrAtarget genes through a direct interaction with CtrA at target promoters (Fig 3.5D). Taken
all together, our results suggest that SciP is normally proteolyzed at the Gl-S transition
and that the failure to properly degrade SciP prevents the activation of CtrA-dependent
genes in predivisional cells, thereby disrupting cell cycle progression.
102
Discussion
Post-translational Regulation ofSciP
In our initial report identifying SciP, we observed the importance of its tight regulation
for proper cell-cycle progression. In wild-type cells SciP protein is present predominantly
in swarmer cells and either its ectopic expression or depletion results in a dysregulation
of the cell cycle. Consistent with the tight temporal regulation of SciP, previous reports
have found that the sciP mRNA is strongly cell-cycle regulated [7, 12]. In this study, I
demonstrated that SciP is also subject to post-transcriptional regulation. Specifically,
SciP protein is proteolyzed at the G 1-S transition by the Lon protease.
Why proteolyze SciP at the G 1-S transition? A priori, one might expect that fluctuations
in sciP mRNA levels are sufficient to restrict SciP to swarmer cells. Several studies,
including ours, have shown that sciP mRNA is present in swarmer cells, disappears
during the Gl-S transition, and is transcribed again in predivisional cells following the
synthesis of CtrA [6, 7, 12]. However, we observed that fluctuations in SciP levels still
occur when sciP is constitutively expressed from a chromosomal locus. As with CtrA,
proteolysis may allow cells to quickly and irreversibly clear SciP from cells ensuring a
robust transition from the GI/swarmer state to the S/stalked state.
Regulated Proteolysis by Lon
The Lon protease is an AAA+ protein that is widely conserved throughout the bacterial
kingdom. CaulobacterLon is highly similar to the E. coli Lon protein that has been well
characterized experimentally. In E. coli, Lon is responsible for degrading unfolded
103
proteins as well as several key regulators like the transcription factor SoxS and the
division inhibitor SulA [2, 13, 14]. The rules for Lon substrate specificity are variable but
Lon is thought to recognize hydrophobic patches in unfolded proteins and the N- or Ctermini of the substrates SoxS and SulA, respectively [4, 15, 16].
In Caulobacter,Lon is known to degrade the cell-cycle methyltransferase CcrM [5]. The
Lon deletion strain exhibits moderate filamentation and accumulation of up to six
chromosomes per cell which has been attributed to a failure of degradation of CcrM [5].
However the Alon phenotype is also similar to the phenotypic consequences of
expressing non-degradable sciP-M2 in wild type cells suggesting that failure to clear SciP
at the G 1-to-S transition may also contribute to the pleotropic effects of a Ion mutant.
In this study, we observed that SciP is proteolyzed specifically at the Gl-to-S transition
and is only allowed to accumulate in early predivisional cells when constitutively
expressed. Previous reports suggested that Lon protease is preferentially synthesized at
the stalked pole of predivisional cells and that stalked cells have approximately twice as
much Lon as swarmer cells immediately after cell division [5, 17]. Perhaps newly
differentiated stalked cells are also slightly enriched in Lon to provide a boost to the
proteolysis of SciP during the Gl -S transition. Alternatively some as of yet unidentified
adapter protein may mediate SciP degradation at the Gl-to-S. In addition, it is possible
that differential localization of Lon during the cell cycle regulates its activity similar to
what has been reported for ClpXP [9].
While SciP has no obvious Lon degrons, we found that SciP is stabilized by the addition
of a C-terminal M2 epitope (DYKDDDDK). One possibility is that some of the many
104
positively charged aspartic acid residues of the M2 epitope may form very stable
interactions with several negatively charged arginine residues at the C-terminus of wildtype SciP (RTTRIQQYR). Such a secondary structure could obstruct normal substrate
recognition by Lon by forming a secondary structure at a normally floppy C-terminal end
or by preventing the C-terminal from interacting with Lon.
Concluding Remarks
Our analysis of the post-transcriptional regulation of SciP provides another key example
of the importance of regulated proteolysis in bacterial cell cycle control. As with other
important regulatory proteins such as CtrA, and CcrM, Caulobacter has evolved
redundant mechanisms for ensuring their correct expression during the cell cycle [1, 5].
Bacterial cells exert transcriptional control in addition to other post-transcriptional
regulation such as phosphorylation, sub-cellular localization, and proteolysis to ensure
robust, stable progression through the cell cycle.
105
Experimental procedures
Bacterial strains and media
E. coli and Caulobacter strains were grown as previously described [18]. PYE was
supplemented with 0.2% glucose (PYEG) and 0.3% xylose (PYEX) as indicated. MG2
was supplement with 0.3% xylose as indicated. ML1750 was grown in PYEX,
synchronized, and released into PYEG or PYEX. KG91, KG313, KG314 were grown in
PYEG supplemented with oxytetracyline, synchronized, and released into PYEG
supplemented with oxytetracyline or PYEX with oxytetracyline. UJ838, UJ199, LS2382,
AcpdR, and hslV: :tn5 were grown in PYE. UJ199 was depleted and synchronized as
previously reported [1]. All synchronies were preformed in a 50% Percoll gradient in 50
mL tubes (Falcon).
Cloning and mutagenesis
The plasmid pENTR-Pxyl-sciP was constructed by TOPO cloning a translational fusion
of sciP to the Pxyl promoter. The translational fusion was made by S.O.E. PCR as
previously described [18]. The Pxyi promoter was amplified from pHXM-DEST using the
primers Pxyl F and Pxyl R. SciP was amplified from genomic DNA using the primers
sciP F and sciP R. The plasmid pENTR-Pxyl-sciPM2 was constructed by TOPO cloning
a PCR fragment amplified from pLX-sciP using the primer Pxyl F and sciP M2 R.
The low copy plasmids with PxI-sciP (pLX-sciP) and Pxyi-sciP-M2 (pLX-sciPM2) were
constructed by LR recombination of pENTR-Pxyl-sciP and pENTR-Pxyl-sciPM2 into the
destination vector pMR20-DEST respectively as previously described [18]. The low copy
106
plasmid with Pxy-M2-SciP (pLXM-sciP) was constructed by LR recombination of
pENTR-sciP into pLXM-DEST. The expression plasmids were electroporated into
CB 15N and Alon electrocompetent cells as indicated.
The pBAD-lon plasmid was constructed by digesting pBAD33-lon with NdeI and Sbft
and ligating in a PCR product amplified from CB15N genomic DNA using the primers
lon F and R and digested with NdeI and SbfL.
Immunoblots
Cell pellets were resuspended in the equivalent of 50 pL 1 X SDS sample buffer per 1
mL of OD 0.15 culture. 10 ptL of lysate was resolved on a 12% gel (Bio-Rad) run at 130
V for 1 hr at room temperature. Samples were transferred to PVDF membrane and
probed as previously described [6].
Flow cytometry
Samples were analyzed as previously described [19].
Pulse-chase
SciP stability was assayed as previously described with the following modifications [6].
Expression of various sciP alleles in KG91, KG313, KG314, and KG336 expression was
pre-induced for 30 minutes immediately before the pulse-chase. Samples
were
immunoprecipitated with 2 RL of SciP anti-sera for 2 hrs at 4 C and immune complexes
were collected for 1 hr at 4 C.
107
DNA microarrayanalysis
Microarrays were conducted as previously reported on Agilent arrays [20]. Experiments
were run in duplicate and the average results are reported.
Protein purifications
CtrA, SciP, and Lon were purified as previously described [4, 6].
In vitro degradation
In vitro degradation assays were run in 25 mM Tris pH 8.0, 10 mM MgC12, 100 mM KCl
buffer with 5 mM DTT, 20 mM phosphoenolpyruvate, pyruvate kinase (0.01 units/pL),
10 iM SciP, and 500 nM E. coli Lon (hexamer) at 30 C. 10 RL aliquots were collected at
indicated time points and quenched in 4x SDS PAGE loading buffer. Reaction products
were separated on 4-20% gradient gel (Biorad).
108
Table S1 - Strains and Plasmids
Strain or Organism or Plasmid
Plasmid
Category
Strain
C.crescentus
E coli
Plasmid
destination vectors
entry vectors
expression vectors
Strain or PlasmidName
Descprition
Synchronizable derivativeof wild-type CB15
CB15N AsciPxylX::sciP (tetRkanR)
CB15N pLX-scP (tet", chlor")
(tot", chlor")
CB15N pLXM-scIP
CB15N pLX-sciPM2 (tet". chlor*)
CB15N Alon pLX-sc/P (tetR,chor,spec")
CB15N AcipA(spec")
CB15N AcIpP xylX::cfpP (spec", tet")
CB15N &cpdR (tet")
CB15N hs/1Vtn5 (kan")
5
CB15N Alon (spec )
LS2382
General cloning strain
DH5a
of destination vectors
One Shot ccdB Survival TV Strain for propagation
clones
General cloning strainfor pENTRID-TOPO
TOP1O
of transgenic Lon
Alonstrain for purification
ER2566
0
(tete,chlor )
pMR20, tow-copy
pMR20-DEST
pMR20-M2; low-copy, Pxyl,N' M2tag (tetRchiorR)
pLXM-DEST
Pxyl,N' M2 tag (spec", chlor")
pJS71X-M2;high-copy,
pHXM-DEST
ENTRY vector forGateway cloning system (kan")
pENTR/D-TOPO
(kanR)
CC0903in pENTR/D-TOPO
pENTR-sciP
(kan")
In pENTRID-TOPO
Pxyl fused to CCO903
pENTR-Pxyi-sc/P
(kanR)
Pxyl fused to CC0903with M2 epitope a C' in pENTR/D-TOPO
pENTR-Pxyl-sciPM2
pMR20-X-sc/P (tet", chlor")
pLX-scIP
pMR20-X-M2-sciP (tet", chlor")
pLXM-sciP
pMR20-X-sc/P-M2 (tet", chior")
pLX-sclPM2
wild-type ctrA promoterincludingP1and P2 fused to lacZ
pctrA290
pctrA290 with muatedP1 (tot")
pctrA-P1
pctrA290 with mutatedP2 (tet")
pctrA-P2
pctrA290 with muated-39 to -48 putative SciPbinding site (tet")
pctrA-bs1
pctrA290 with musted-70 to -79 putatitve SciPbinding site (let")
pctrA-bs2
pctrA290 with both-39 to -48 and -70 to -79sites mutated (tet")
pCtrA-bsl +2
pBAD33-ion with Caulobacter tonreplacing E. colt on (chlor")
pBAD-lon
Ionexpression plasmid(chlor")
pBDA33-lon
CB15N
ML1750
KG314
KG91
KG313
KG336
UJ838
UJ199
Source or Reference
Evinger and Agabian, 1977
Gora et al, 2010
this study
this study
this study
this study
Grunenfelder et al, 2004
Jenal et al, 1998
Skerker et al, 2005
Josh Modell, unpublished
Wright et al, 1996
invitrogen
Novagen
invitrogen
New England Biolabs
Chen et al, 2009
Skerker st al, 2005
Skerker et al, 2005
Invitrogen
Gora et a, 2010
this study
this study
this study
this study
this study
Domian et aL.1999
Domian et al. 1999
Domian et al. 1999
this study
this study
this study
this study
Christensen st al. 2004
Table 3.1. Strains.
TableS2 - Primers
PusionPCR
Pxyl
sciP
sciPM2
Gel Shift
Ion
Forward primer
CACCGGTACCTCGAACAGGGCCGTCAGGTCGCG
AGTTTGGGGAGACGACCATATGTTGCAGCAGCAGCGCAC
TTTCATATGTCCGAACTACGTACGCTTCCTG
Reverseprimer
ATGGTCGTCTCCCCAAAACT
AGTTTTGGGGAGACGACCATATGTTGCAGCAGCAGCGCAC
TTACTTGTCGTCGTCGTCCTTGTAGTCACGGTACTGCTGGATCCGGG
TTCCTGCAGGTrAGTGCGTCAGCAT
Table 3.2. Primers.
109
Acknowledgements
We thank J. Modell for his generous gift of the hslV::tn5 strain and U. Jenal for providing
published strains. M.T.L. is an Early Career Scientist at the Howard Hughes Medical
Institute. This work was supported by an NIH grant (5R01GM082899) to M.T.L.
110
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Sauer, R.T. and T.A. Baker, AAA + Proteases:A TP-fueled Machines of Protein
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Gur, E. and R.T. Sauer, Recognition of misfoldedproteins by Lon, a AAA(+)
protease. Genes & Development, 2008. 22(16): p. 2267-2277.
Wright, R., et al., CaulobacterLon protease has a critical role in cell-cycle
control ofDNA methylation. Genes & Development, 1996. 10(12): p. 1532-42.
Gora, K.G., et al., A cell-type-specific protein-proteininteractionmodulates
transcriptionalactivity of a master regulatorin Caulobactercrescentus.
Molecular Cell, 2010. 39(3): p. 455-67.
Tan, M.H., et al., An essential transcriptionfactor, SciP, enhances robustness of
Caulobactercell cycle regulation.Proceedings of the National Academy of
Sciences of the United States of America, 2010. 107(44): p. 18985-90.
Meisenzahl, A.C., L. Shapiro, and U. Jenal, Isolation and characterizationof a
xylose-dependentpromoterfrom Caulobactercrescentus. Journal of Bacteriology,
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Iniesta, A.A., et al., A phospho-signalingpathway controls the localizationand
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Grunenfelder, B., et al., Identification of the Protease and the Turnover Signal
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Rohrwild, M., et al., HslV-HslU: A novel A TP-dependentproteasecomplex in
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Chen, L.S., D. Mullin, and A. Newton, Identification,nucleotide sequence, and
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Mizusawa, S. and S. Gottesman, Proteindegradationin Escherichiacoli- The
Lon gene controls the stability of SulA protein. Proceedings of the National
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Griffith, K.L., I.M. Shah, and R.E. Wolf, Proteolytic degradationofEscherichia
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Shah, I.M. and R.E. Wolf, Sequence requirementsfor Lon-dependent degradation
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112
Chaper 4
Conclusions and Future Work
112
In this thesis I have presented work that furthers our understanding of the complex
regulation of CtrA, a master regulator of the Caulobacter cell cycle. In Chapter 2, I
described the identification of SciP, a swarmer-specific inhibitor of CtrA transcriptional
activity and showed that SciP binds CtrA at CtrA-dependent promoters and likely
prevents CtrA from recruiting RNA polymerase to these promoters. I showed that SciP,
although not essential, is critical for cell cycle progression and either the absence or
overproduction of SciP caused severe cell cycle defects through misregulation of the
CtrA regulon. In Chapter 3, I investigated the post-transcriptional regulation of SciP and
showed that SciP is specifically proteolyzed at the Gl-S transition by Lon protease. I
created a stable allele of sciP by epitope-tagging the C-terminus and demonstrated that a
failure to clear SciP during the G 1-S transition disrupts cell cycle progression.
The regulation of CtrA by SciP constitutes a novel mechanism for regulating the activity
of a response regulator. It is the first protein shown to block transcriptional activation by
a response regulator without affecting phosphorylation or DNA-binding. Like CtrA, SciP
homologs are present throughout alpha-proteobacteria and are highly conserved, with
orthologs typically exhibiting more than 75% amino acid identity. Given the coinheritance of SciP and CtrA, it seems likely that SciP is an important regulator of CtrA
throughout the alpha-proteobacteria. However it remains to be seen whether these SciP
orthologs do in fact regulate CtrA orthologs or if even small changes in the amino acid
sequence allow them to perform different functions.
113
Furthercharacterization of the interaction between CtrA and SciP
Although the biochemical experiments in chapter 2 support the idea that SciP and CtrA
form a complex at CtrA activated promoters, several questions remain about the
molecular nature of this interaction.
Does SciP contact DNA and if so, how?
In chapter 2, I used electrophoretic mobility assays (EMSA) to demonstrate that SciP
alone cannot bind DNA, but that it does form a complex with DNA-bound CtrA. Our
analyses further showed that complex formation requires free DNA adjacent to that
bound by CtrA; 25 base pair probes that are likely completely covered by CtrA did not
experience a super-shift upon addition of SciP, in contrast to 50 basepair probes. Our
analyses did not uncover any sequence requirements for this putative SciP-DNA
interaction suggesting that any interaction between SciP and DNA is non-specific. In
contrast, a recent study suggested that SciP can bind DNA on its own and that it
specifically binds the motif GTCGC [1]. However, the binding of SciP reported was
weak and not concentration-dependent. Moreover, the motif GTCGC is unlikely to
provide binding specificity within a genome that has nearly 70% GC content.
Nevertheless, I tested the importance of this putative motif for SciP/CtrA/DNA complex
formation by mutating the core nucleotides of the motif in a pilA promoter probe used in
Chapter 2. We mutated TGTCGCG to TGTTTTG and tested the ability of CtrA and SciP
to bind the mutant probe (Fig. 4.1A). As expected, purified CtrA~P alone bound both the
wild-type and mutant probes, producing nearly identical shifts that correspond to a CtrADNA complex (Fig. 4. 1B). We then added purified SciP along with CtrA-P and observed
114
a similar pattern of super-shifting with both probes (Fig. 4.1 B). These results suggest the
TGTCGG motif is not required for a SciP-CtrA-DNA complex to form on this probe.
This conclusion is also consistent with our previous report that SciP and CtrA form a
complex on a 50 base-pair probe taken from the fliF promoter that does not include the
TGTCGG motif.
To corroborate these results in vivo, we examined a Pet,.A-lacZ reporter; this construct
contains both the P1 and P2 promoters of ctrA fused to lacZ and two instances of the
TGTCGG motif (Fig. 4.1 C) [1, 2] We mutated each motif independently and in a double
mutant to create PctrAbs1, PetrAbs2, and Pct,.Absl+2 respectively. The PetrAbs2 reporter
exhibited activity comparable to the wild-type reporter suggesting that the distal
TGTCGG motif is not important for the regulation of the ctrA promoter (Fig 4.1D). The
PetrAbs] and PetrAbs1 +2 reporters both exhibited a 25% increase in activity relative to the
wild-type reporter. As a control, we compared the differences in activity to two additional
PetrA mutants, PctrA-P1 and PctrAP 2 that preserve only the P1 and P2 promoters of ctrA,
respectively. In our hands, eliminating P1 (PctrA-P2) resulted in a -25% decrease in
activity and eliminating P2 (PctA-PI) reduced activity by ~60% in general agreement
with previously published data (Fig. 4, 1D) [2].
To test if mutating the TGTCGG motifs affects SciP-dependent regulation of ctrA we
examined expression from PctrAbsl, PctrAbs2, and PctrAbsl+2 in cells overexpressing sciP.
As shown in chapter 2, overproducing SciP downregulates ctrA expression. We induced
expression of sciP from a high copy plasmid for 2 and 4 hrs and then measured promoter
activity. We found that sciP overexpression reduced the activity of all three reporters to
similar extents (Fig. 4. 1E). Therefore although one of the TGTCGG motifs has a modest
115
effect on ctrA expression, that effect does not depend on SciP raising the possibility that
that motif may be important for the binding of a yet unknown regulator.
A.
be
5
0
ppp.A
aca
C. s' ACGCTGAAAA
e-2L
CACCCGCCTG
CGACGCCTGC
GACCAATGTG
PpILAbs
bel
x XXX
A TAT
TTCAGGGCT CCGACGGC TTGCACCCGA TTCGCAAATC
consensus A/C
TGTCGCA
AGMKC3
CTCClAC GGG 3'
crA bei
CGC
crA b2
A/C
consensus
B.
PpILA
ScIP
(1IpM)
CtrA (pM)
+
0 0.5 1
D.
PpIlAbs
+
0 0.5 1
+ +
0 0.5 1
1.75
+
1.50
0 0.5 1
1.25
1
TGTCGCA
1.00 - - - - - 0.75
complexes
0.50
0.2
0.25 -P
kesDNA
E.
P2 bsl bs2 bs1+2
Reporter Construct
1.00-
o
S0.75
21
Fiue0.50
wt
bel
bs2
bs1+2
Reporter Construct
Figure 4.1. Dispensability of the putative SciP motif in vitro and in vivo. (A) Mutation of the
putative SciP motif (arrow) in a PpiA probe. CtrA binding sites are boxed. The motif is compared
to the reported consensus sequence below. (B) Gel shift analysis with wt PpiiA vs PpilAbs.
CtrA-P was added at the indicated final concentration and SciP was at luM when present. (C)
Mutation of two putative SciP motifs (arrows) in the PctrA-lacZ reporter with annotations as in A.
Only a subset of the promoter fragment is shown. (D) Beta-galactosidase activity of PctrAbsl
(bsl), PctrAbs2 (bs2), PctrAbsl+2 (bsl+1) relative to wt PctrA in CB15N shown in grey. The
PctrAP1 and PctrAP2 mutants are shown for reference. (E) Activity of PctrA (wt), PctrAbsl (bsl),
PctrAbs2 (bs2), PctrAbsl+2 (bsl+2) in CB15N harboring sciP on a high copy xylose inducible
plasmid, induced for 2 hrs (dark grey) or 4 hrs (light grey) vs. uninduced wt.
116
Why does SciP require free DNA for complex formation?
Recent homology searches suggest that SciP has a helix-turn-helix motif and an NMR
structure of the SciP homolog from Rhodobacter sphaeroides allows us to model the
three-dimensional structure of SciP [3]. Based on the R. sphaeroides structure, SciP
consists of two beta sheets followed by four distinct alpha helices with unstructured N
and C-terminal tails (Fig 4.2A, B). A recent study has speculated that SciP interacts with
the major groove of DNA through the third alpha helix [1]. In this model the residues
R35 and R40 are adjacent to the DNA backbone but are not within the canonical DNA
recognition helix. This model is consistent with our mutational analysis in Chapter 2 in
which the mutations SciP(R35A) and SciP(R40A) disrupt the interaction between SciP
and CtrA in both the yeast two-hybrid and EMSA assays suggesting these residues
mediate a direct interaction between SciP and CtrA. If helix three is the main determinant
of the SciP DNA interaction, then R35 and R40, which are located at the beginning of
helix one and are exposed to adjacent DNA, might be free to interact with CtrA.
Interestingly, our attempts to mutate residues in helix one, two, and three destabilized the
SciP protein based on the failure of these mutants to accumulate when overexpressed in
vivo as well as poor purification yields (unpublished data).
117
ED ED
MLQQQRTNSRGEKYVIGPTGAPLTLADLPPAETQRWVIRRKAEVVAAVRGGLLSLDEACDRYKLTNDEFLSWQQSIDRHGLAGLRTTRIQQYR
tft
B.
C.
D60A
T65M
R40A
R40S
R40
R35
W36A
k
Figure 4.2. Modeling the three-dimensional structure of SciP. (A) Secondary structure
domains superimposed on the SciP primary amino acid structure. R35 and R40 are indicated by
arrows. SciP was modeled onto the R. sphaeroides SciP homolog 2jRT (B). (C) Proposed
interaction between SciP helix three and DNA. Image taken from [1].
What is the molecular surface that mediates the interaction between CtrA and SciP?
In Chapter 2, I used alanine scanning to probe SciP for residues necessary for interacting
with CtrA. We are also interested in identifying the residues of CtrA that mediate its
interaction with SciP. To find candidates, I performed a structural alignment of CtrA with
a structure of E. coli OmpR to identify solvent exposed residues. Because SciP interacts
with CtrA but not Caulobacter PhoB, I compared the solvent exposed residues of CtrA
with PhoB to identify significant differences. This analysis yielded approximately 20
candidate resides for mutational analysis, 11 of which I tested in the yeast two-hybrid
assay for interaction with CtrA. Alanine mutations at 4 of these 11 sites led to a
disruption of the interaction between SciP and CtrA in the yeast two-hybrid assay. One
mutant, CtrA(Y179A), was able to weakly bind DNA in EMSAs but failed to interact
with SciP (Fig. 4.3). Residue Y179 is located within the DNA-binding domain, is solvent
118
exposed, and predicted based on a comparison to the OmpR to not contact DNA. This
work is still preliminary but it may provide a clue to the CtrA/SciP interface. Ultimately a
co-crystal structure of the SciP/CtrA/DNA complex would be most informative.
wt
CtrA(uM)
ScIPiuM
1
+
1
+
H177A
1 1 2 2
+
+
Y179A
1 1 2 2
+
+
Figure 4.3. Electrophoretic mobility assays with mutant CtrA and wild-type SciP. Wild-type
CtrA-P and indicated CtrA mutants binding to a fragment of the fliF promoter containing a CtrAbinding site. SciP, if present, was added at a final concentration of 1.0 RM. The concentration of
CtrA in each lane is indicated.
Regulated proteolysis and the Caulobacter cell cycle
In Chapter 3, I showed that the Lon protease degrades SciP in vivo and in vitro. Our
experiments also indicate that SciP is specifically proteolyzed at the Gl-S transition.
However, the Lon protease is present throughout the cell cycle and degrades another
substrate, CcrM at a different, later stage of the cell cycle [4]. These observations beg the
question of how Lon proteolyzes SciP only during the GI-to-S transition.
One possible answer is that SciP is targeted for proteolysis during the G 1-S transition by
an adaptor protein that is only produced or active during this window of time. There are
many examples of adapter proteins that regulate proteolysis of key regulatory proteins in
119
other bacteria. For instance, in E. coli, the sigma factor RpoS is present at low levels
during exponential growth because it is proteolyzed by ClpXP in an RssB dependent
manner [5]. RssB is a response regulator that acts as an adapter protein binding to a
recognition sequence on RpoS [6]. RpoS interacts with the phosphorylated form of RssB
leading to the speculation that dephosphorylation of RssB under stress conditions
stabilizes RpoS [6]. One strategy for identifying a SciP adaptor(s) would be to perform a
yeast two-hybrid screen. One could use a bait plasmid containing SciP and co-transform
with a random genomic library of prey plasmids and select for interactors using the
appropriate nutritional markers. Because SciP is highly soluble and readily interacts with
CtrA in the yeast two-hybrid, such a selection might quickly yield potential adaptor
proteins.
Another possibility is that the activity of Lon is regulated by some other mechanism such
as subcellular localization. As previously discussed, Caulobacter ClpXP proteolyzes
CtrA specifically at the Gl-to-S transition despite the fact that ClpP and ClpX subunits
are present throughout the cell cycle [7]. ClpXP activity is thought to be controlled by its
localization to the stalked pole during the Gl-S transition [8]. An additional adapter
protein, RcdA, may be required to bring CtrA to the stalked pole for proteolysis [9]. The
subcellular localization of Lon has not been examined so it is possible a similar
mechanism might regulate the activity of Lon and the degradation of SciP during the G 1S transition.
Finally, it is possible that some form of post-translational modification could play a role
in regulating SciP proteolysis at the Gl-to-S transition. Recent whole-proteome mass
spec studies have only begun to catalog the full diversity and prevalence of post120
translational modifications in bacteria including proteolytic cleavages, methylation, and
acetylation [10, 11]. In contrast to ubiquitin-targeted proteolysis in eukaryotic systems,
post-translational modifications have only rarely been implicated in proteolytic control in
bacteria. One key example is stabilization through proteolytic cleavage of UmuD, an
alternate RNA polymerase subunit induced during the SOS response in E. coli [12]. Upon
SOS induction, RecA cleaves the 24 N-terminal amino acids of UmuD which protects the
protein against proteolysis by Lon by ostensibly eliminating a degradation signal in that
N-terminal domain [13]. Although we have not observed any conspicuous shifts in SciP
migration in Western blots, such a proteolytic cleavage might result in a small shift and
therefore difficult to detect without mass spec analysis of the Caulobacter proteome.
Interestingly, mass spec analysis is not well suited to identifying post-translational
modification by phosphorylation in two-component systems because of the lability of
phosphoramidate bonds characteristic of His~P and Asp~P modifications [14]. However
in our analysis of SciP we were unable to find any conspicuous phosphorylation site
motifs suggesting that SciP is not likely subject to regulation by phosphorylation.
Outstanding questions in cell cycle regulated transcription in Caulobacter
Although the work presented here contributes to our understanding of the fine-tuning of
CtrA-activated transcription, questions remain about CtrA and cell cycle regulated
transcription in general. For instance, some CtrA-activated genes have abundant
transcripts in swarmer cells [15]. It is unclear whether these genes are actively transcribed
in swarmer cells and somehow immune to the effects of SciP or if these genes are
expressed in predivisional cells with transcripts that are unusually stable and hence still
present in swarmer cells, as is known to be the case with some flagellin genes [16]. If the
121
late genes are actively transcribed in swarmer cells, how do they avoid inhibition by
SciP? Perhaps the late genes share a distinct promoter architecture that is not inhibited by
SciP. Alternately or in addition, these genes may require another co-factor to fine tune
their expression pattern.
Although CtrA is a key global regulator of cell cycle transcription in Caulobacter, it
directly regulates only ~100 out of more than 500 cell cycle-regulated genes [15].
Another transcriptional regulator, GcrA has been implicated in the regulation of an
additional 125 genes [17]. However, most of these genes showed very limited, i.e. less
than two-fold, changes in expression upon depletion of GcrA [17]. Further, it remains
unknown if GcrA directly binds DNA. Regardless, the expression of at most 30% of cell
cycle regulated genes can be explained by CtrA and GcrA suggesting that additional
factors remain to be discovered.
What regulates the remaining 70% of cell-cycle-regulated genes? One possibility is that
one or more uncharacterized transcription factors play key regulatory roles. The
Caulobacter genome contains approximately 200 predicted transcription factors, 44 of
which are response regulators [18]. A previous study from our lab deleted each of the
response regulator genes and screened for cell cycle defects [19]. Five were found to be
essential, including ctrA and divK [19]. Two of the remaining three, (petR and ntrY) are
homologs of transcription factors that regulate metabolic genes, and the third, cenR,
regulates genes important for cell envelope metabolism [19]. Two other response
regulators exhibited significant cell cycle defects: flbD and tacA [19]. FlbD is a
122
transcriptional regulator of class III and V flagellar genes while TacA controls stalk
biogenesis by activating the transcription factor StaR [20, 21]. These two regulators thus
regulate some, but not all, of the remaining cell cycle genes. Our lab is currently deleting
and characterizing the other remaining predicted transcription factors in the Caulobacter
genome. However, preliminary results suggest that few have severe cell cycle phenotypes
suggesting there may be no other global regulators of transcriptional activity or that any
remaining global regulators are at least partially redundant in function.
It is also possible that sigma factors play a significant role in cell cycle transcription. In
eubacteria, the RNA polymerase core enzyme consists of five subunits (a2ppa) that must
associate with a sigma factor to form the holoenzyme that can recognize promoters and
activate transcription [22]. In E. coli, there are seven different sigma factors that can
associate with the RNAP core enzyme to direct global transcription patterns of different
classes of genes [22]. C70, encoded by the rpoD, directs the expression of genes in
exponential growth while as (rpoS) drives expression of genes during the general stress
response [22] . Y54 (rpoN) regulates genes involved in nitrogen assimilation and other
metabolic functions [23].
F
(fliL) directs the expression of late flagellar genes while -2
(rpoH), dE (rpoE), and Fec respond to specific stress conditions [22]. The Caulobacter
genome encodes 16 predicted sigma factors, almost three times as many as E. coil [18].
As in E. coli, rpoD encodes the dominant sigma factor, 073, and rpoH encodes
&2
[24-
26]. In addition the gene rpoN encodes Y54, which regulates the expression of class II and
IV flagellar genes as well as genes involved in stalk morphogenesis analogous with the
role of J in E. coli [20, 27]. Of the remaining 13 predicted sigma factors, only three have
been described. oJ (sigF) is thought to protect Caulobacter from oxidative stress during
123
stationary phase, while ad (sigT) and d
(sigU) may have a more general role in
responding to oxidative stress and heavy metals [28-30]. Given the precedence of sigma
factors regulating specific stress responses, it seems unlikely that the remaining 10
uncharacterized sigma factors in Caulobacter contribute significantly to cell cycleregulated transcription [22].
One final possibility is that chromosome structure may play an important role in cell
cycle-regulated transcription in Caulobacter.In bacteria, the chromosome forms a highly
condensed structure of supercoiled DNA called the nucleoid. In E. coli, it has been shown
that DNA topology changes in response to different growth conditions and environmental
stresses through changes in the activity of topoisomerase, DNA gyrase, and various
nucleoid associated proteins [31]. Several studies have shown that changes in DNA
supercoiling influences global gene expression [32]. For instance quickly reducing
supercoiling changes the expression of about 300 genes in E. coil [33]. In this study,
chromosome relaxation was accomplished by inhibiting gyrase and topoisomerase with
antibiotics or through temperature sensitive alleles [33]. Although nucleoid structure and
dynamics in Caulobacterhas not been as extensively studied as in E. coli, recent work
has shown that the physical location of various chromosomal loci with respect the long
axis of the cell depends linearly on their distance from the origin suggesting that the
Caulobactergenome is also highly ordered and structured [34]. It would be interesting to
define nucleoid structure as a function of the cell cycle using new methods such as Hi-C
and to then look for correlations between changes in local nucleoid structure and the
expression patterns of cell cycle genes [35].
124
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