Regulatory pathways controlling cell division
after DNA damage in Caulobacter crescentus
by
Joshua Wexler Modell
B.S. Biology
Duke University, Durham, North Carolina, 2005
SUBMITTED TO THE DEPARTMENT OF BIOLOGY IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY IN BIOLOGY
AT THE
MASSACHUSETTS INSTITUTE OF TECHNOLOGY
ARCHVE$
H
IT}
-
SEPTEMBER 2013
0 2013 Joshua Wexler Modell. All rights reserved.
The author hereby grants MIT permission to reproduce and distribute publicly
paper and electronic copies of this thesis document in whole or in part in any medium now
known or hereafter created.
Signature of Author:
Joshua Wexler Modell
Department of Biology
August 13, 2013
Certified by:
Michael T. Laub
Assosciate Professor of Biology
Thesis supervisor
Accepted by:
Stephen P. Bell
Professor of Biology
Graduate committee chair
1
Regulatory pathways controlling cell division
after DNA damage in Caulobacter crescentus
by
Joshua Wexler Modell
Submitted to the Department of Biology
on April 29, 2013 in partial fulfillment of the requirements for the degree of
Doctor of Philosophy in Biology at the Massachusetts Institute of Technology
ABSTRACT
All cells must coordinate DNA replication with cell division in order to faithfully propagate
whole chromosomes to daughter cells. During episodes of DNA damage, cells often delay
division until the lesions have been repaired and replication has completed. The paradigm for the
bacterial response to DNA damage is the transcriptional induction of "SOS" genes, and many
organisms encode an SOS-induced cell division inhibitor. However, the mechanistic details of
division inhibition are understood only in the y-proteobacterium E. coli, and it is unclear whether
there are SOS-independent modes of division inhibition.
I have studied the DNA damage response in the u-proteobacterium Caulobactercrescentus and
identified two damage-induced cell division inhibitors. sidA is an SOS-induced division inhibitor
whereas didA is induced in an SOS-independent fashion. Unlike most division inhibitors, SidA
and DidA do not disrupt the localization of the cell division scaffold FtsZ or any other
component of the cell division machinery or "divisome". Instead, SidA and DidA target the lateacting division proteins FtsW, FtsI, and FtsN to prevent divisome constriction, demonstrating
that divisome components other than FtsZ can serve as regulatory targets. I have characterized
mutations infts W andftsI which suppress the activities of both inhibitors, likely by causing cells
to divide hyperactively. These results suggest that the FtsW/FtsI/FtsN subcomplex serves as an
important regulatory node and may play an unexpected role in triggering divisome constriction
in Caulobacter. I show that cells require at least one inhibitor to properly delay division
following DNA damage, as cells lacking both inhibitors divide prematurely and suffer a viability
defect in the presence of the DNA damaging agent mitomycin C (MMC). This finding suggests
that some degree of redundancy exists within the Caulobacter response to MMC. Finally, I
describe ongoing experiments which explore the origins of the SOS-independent induction of
didA.
Thesis supervisor: Michael T. Laub
Title: Associate Professor of Biology
2
ACKNOWLEDGEMENTS
I am grateful to many people for helping me to grow as a person and scientist and for helping me to grow
Caulobacter.
Dr. Michael Laub, you have been supportive, creative, diligent and humorous in all the right amounts,
and I can't imagine working for a better boss. I also enjoyed extorting Red Sox tickets from you under the
threat of coining ironic names for new genes.
Tracy and Alex, one of you was the best UROP I ever had. None of this work would have been possible
without you guys, and you never once complained about my incoherent babbling. Both of you gave me your
hearts and souls, and I returned them covered in bacteria. Alex, we really should have redesigned the food
wheel after the BioCafe closed. Tracy, I wish you had put more trust in the food wheel.
Barrett, you taught me everything I currently know about Caulobacterhusbandry and the state of Rhode
Island. Thank you for your patience and steady guidance, and for preventing the lab from falling into a postapocalyptic Thunderdome scenario.
Anna, thank you for your nonsensical banter, catchy ringtones and backup vocals. Your propensity for
abominable internet videos and news reports of questionable authenticity have somehow kept me sane through
four difficult years. I ghaze we'll sieve what happens next.
Christos and Emma, it was a true pleasure watching you two fall in love every day. Thank you for your
generosity in friendship and reagents, and especially for your willingness to constantly lose to me in Nerf
basketball, presumably to make me feel better about my science. Erin, Phil, Kasia, Emily and Darwin, you
guys were so furry and cute, I loved watching you run around and sleep under the table outside Mike's office.
Chris, thanks for being the only other lab member to maintain him/herself in peak physical condition. Andy, it
was a joy fostering a Norwegian conference-child with you. Diane, thanks for letting me borrow xylose during
that xylose emergency. Michael S., you will make a worthy gossip chair. Thank you for never spoiling and
occasionally unspoiling Game of Thrones for me. Katie and Kyle, you guys made room three into a real family
dining experience. To everyone else in the lab, thanks for the good times and for not making me smile during
my mouth surgery.
To my grandparents, thank you for being fruitful and multiplying. To my parents and sisters, thank you
for feeding and loving me and for occasionally feigning interest in Caulobacter.Your support and perspective
was often all that stood between me and madness, and I am blessed to have been raised and nurtured by each
of you. Thank you for never questioning my decision to attend graduate school, and please never allow me to
return.
To my friends in Boston, New York and beyond, thanks for celebrating and commiserating with me and
for making way more money than me and not rubbing it in too much.
To Charles, you were a tiny gentleman in a cat suit, and I know you're licking an adult male's head of
hair and eating sushi in cat heaven.
To Cleo, thanks for giving me a purpose and a constant reason to smile. You have reawoken my love for
chocolatey, peanut-buttery sundaes, and you have kept me balanced, healthy and happy during these trying
times. You are a sweet, sweet angel that God sent to me to fulfill a contractual obligation with the Burren, and
I am especially grateful for you.
3
TABLE OF CONTENTS
Abstract.................................................................................................
.. 2
A cknow ledgem ents......................................................................................
3
Chapter O ne: Introduction...............................................................................
7
Introduction
Bacterial cell division
Identification of cell division genes
FtsZ positioning
Divisome assembly
The execution of cytokinesis
What drives cytokinesis?
The temporal regulation of cell division
Control of cell division during the cell cycle
Conditional control of cell division
The DNA damage response
The SOS regulators RecA and LexA
The SOS regulon
SOS-induced division inhibitors
SOS-independent regulation
Caulobacter crescentus as a model system
The regulation of cell division in Caulobacter
Research summary
References
Chapter Two: A DNA damage checkpoint in Caulobactercrescentus inhibits cell division
through a direct interaction with FtsW ................................................................
51
Abstract
Introduction
Results
DNA damage induces cell cycle arrest and a change in global transcription patterns
Identification of sidA, a novel SOS-induced cell division inhibitor
SidA is the principal SOS-induced cell division inhibitor
SidA is sufficient, when overproduced, to inhibit cell division
SidA does not disrupt divisome assembly or localization
Mutations infts W andftsl suppress the effects of overproducing SidA
SidA resides in the inner membrane and interacts directly with FtsW and FtsN
SidA prevents constriction of the Z-ring without disrupting synthesis of nascent septal peptidoglycan
Discussion
Is SulA an outlier?
Redundancies in cell division regulation
Final perspectives
Materials and Methods
Acknowledgements
References
Chapter Three: A DNA damage-induced, SOS-independent checkpoint regulates cell division in
Caulobactercrescentus..................................................................................93
Abstract
Introduction
Results
Identification of didA, a DNA damage-induced, SOS-independent cell division inhibitor
DidA interacts with the late-arriving divisome component FtsN
Mutations inftsW can suppress the division inhibition caused by either SidA or DidA
SidA and DidA suppressor mutations drive hyperactive cell division
4
SidA and DidA redundantly regulate division during MMC treatment
Discussion
SOS-independent regulation of the DNA damage response
The execution and regulation of cell division
Final Perspectives
Materials and Methods
Acknowledgements
References
Chapter Four: Conclusions and future directions......................................................
126
Conclusions
New Roles for FtsW/FtsI/FtsN in cytokinesis
Characterizing the FtsW/FtsI/FtsN activity pathway
Why target FtsW/FtsI/FtsN?
The SOS-independent regulation of didA
DriD is required for DidA induction
The identification of a gene transfer agent (GTA)
References
5
LIST OF FIGURES
Chapter One: Introduction
Figure
Figure
Figure
Figure
Figure
1.1. Mitotic checkpoints in eukaryotes........................................................................10
12
1.2. An overview of bacterial cell division......................................................................
1.3. Temporal and spatial regulators of division converge on FtsZ.......................................25
30
1.4. SOS induction of the division inhibitor sulA............................................................
34
1.5. The Caulobacter cell cycle..................................................................................
Chapter Two: A DNA damage checkpoint in Caulobactercrescentus inhibits cell division
through a direct interaction with FtsW
Figure
Figure
Figure
Figure
Figure
Figure
Figure
Figure
Figure
Figure
Figure
Figure
2.1. DNA damage induces global changes in gene expression and inhibits cell division...............
2.2. Annotated gene expression profiles................................................................................
2.3. sidA is the primary SOS-induced cell division inhibitor..............................................
2.4. Overproducing SidA is sufficient to inhibit cell division..............................................
2.5. SidA does not prevent assembly of the cell division machinery........................................
2.6. Mutations infts W andftsI suppress the SidA overproduction phenotype. ..........................
2.7.fts W andftsI suppressor mutations display wild-type morphology during growth in rich medium
2.8. SidA interacts with the late-arriving divisome components FtsW and FtsN...........................
2.9. SidA and FtsW localize to constriction sites..............................................................
2.10. Overproducing SidA does not inhibit the translocation of septal peptidoglycan precursors......
2.11. Quantification of Van-FL foci...........................................................................
2.12. A DNA damage checkpoint regulating cell division in C. crescentus.............................
58
60
62
65
67
71
72
74
75
77
78
81
Chapter Three: A DNA damage-induced, SOS-independent checkpoint regulates cell division in
Caulobactercrescentus
Figure 3.1. Filamentation in sidA and recA mutants.................................................................
Figure 3.2. didA is induced by DNA damage and is not SOS regulated.............................................
Figure 3.3. Annotated gene expression profiles........................................................................
Figure 3.4. DidA is sufficient to inhibit cell division..................................................................
Figure 3.5. DidA is a small, inner-membrane protein that interacts directly with FtsN...........................
Figure 3.6. DidA interacts directly with FtsN...........................................................................
Figure 3.7. Mutations infts W suppress SidA and DidA overproduction phenotypes. ...........................
Figure 3.8. SidA interacts directly with FtsW............................................................................
Figure 3.9. Mutations that suppress sidA and didA overexpression likely hyperactivate cell division..........
Figure 3.10. Suppressor mutant growth properties.....................................................................
Figure 3.11. Cells lacking sidA and didA cannot properly regulate cell division following DNA damage....
Figure 3.12. Parallel pathways regulate cell division in Caulobacter following DNA damage..................
98
100
101
102
104
105
108
109
110
110
112
117
Chapter Four: Conclusions and future directions
Figure
Figure
Figure
Figure
Figure
Figure
Figure
Figure
4.1.
4.2.
4.3.
4.4.
4.5.
4.6.
4.7.
4.8.
Models of FtsW/FtsI/FtsN and SidA/DidA activity.......................................................133
didA is induced specifically by DNA damaging agents...................................................137
138
sidA and didA show are differentially regulated...........................................................
139
Screening for transcriptional regulators of didA...........................................................
139
DriD is activated post-translationally to induce didA.....................................................
SprD represses didA during stationary phase...............................................................141
Gene transfer agents (GTAs).................................................................................141
GTA production in stationary phase induces didA.........................................................142
6
Chapter 1
Introduction
7
Introduction
My grandfather always told me to "be fruitful and multiply." Whether this suggestion was
meant for the cells in my seminiferous tubules or the bacteria in my gut, the task is easier said
than done. In order to multiply, cells must grow and divide, endowing each daughter cell with all
the components necessary for survival and future growth. Specifically, each daughter must
receive an intact cellular envelope, a full genome and the macromolecular machinery needed to
perform repair, regulatory and biosynthetic functions. As division events are energetically costly
and irreversible, nearly all cells have mechanisms to ensure that division is tightly regulated,
occurring only after the accumulation of enough of these essential goods. However, even in the
most well-studied model organisms, the genetic networks responsible for spatially and
temporally controlling cell division remain incompletely understood.
One of the most crucial tasks for a dividing cell is the faithful partitioning of a complete set
of chromosomes to each daughter cell. A failure in genome partitioning that causes certain
chromosomal regions to be absent or over-represented could result in lethality or, in higher
organisms, oncogenesis (reviewed in (Holland and Cleveland, 2012)). Cells must therefore
divide only after the DNA has been fully replicated and each copy spatially segregated to create
a DNA-free division plane. This task is challenging owing to the difficulties inherent in
packaging and manipulating a genome which, when unwound, is orders of magnitude longer
than the cell in which it resides. Furthermore, cells frequently encounter endogenous and
environmental DNA damaging agents which cause lesions that can necessitate lengthy periods of
repair. As a result, the timing of cell division must be somewhat flexible, and the decision to
divide must be intimately linked to the status of DNA replication and repair.
How do cells ensure that division occurs only after a successful round of DNA replication?
Hartwell and Weinert described two general strategies for establishing a dependent relationship,
whereby the initiation of event B requires the completion of a prior event A (Hartwell and
8
Weinert, 1989). In a "substrate-product" relationship, event B is triggered by an intrinsic
property of event A revealed only upon its completion. For instance, the successful completion
of replication could expose a chromosomal structure that serves as a substrate or physical
component of the division machinery. Alternatively, some dependencies are enforced by
dedicated control mechanisms or "checkpoints", in which extrinsic factors that do not participate
in the completion of either step nonetheless provide regulatory control. In this case, incomplete
or stalled DNA replication could activate an extrinsic surveillance system that prevents cell
division; or, the completion of DNA replication could inactivate a system that otherwise
constitutively prevents division.
Examples of both substrate-product and checkpoint models are abundant, but in eukaryotes,
DNA replication and segregation are coordinated with division mainly through a series of
checkpoints (reviewed in (Tyson and Novak, 2008)). Eukaryotic cells divide by a process known
as "mitosis", during which replicated chromosomes are spatially segregated by spindle
microtubules, and a contractile ring of actin and myosin filaments drives constriction of the
envelope and splitting of the cell (reviewed in (Fededa and Gerlich, 2012; Glotzer, 2005)). Two
closely related checkpoints ensure that cells initiate mitosis only once the DNA has been fully
replicated and all lesions have been repaired. While stalled replication forks and damaged DNA
structures are likely recognized by distinct classes of sensor proteins, both ultimately activate
signal transducers that inhibit the activity of a cyclin-dependent kinase (CDK) involved in
mitotic entry (reviewed in (Hurley and Bunz, 2007)). Cyclin-dependent kinases belong to a
family of regulatory proteins whose substrates perform essential cell cycle steps in a
phosphorylation-dependent manner. The mitotic CDK complex, mCDK, phosphorylates targets
involved in chromosome condensation and spindle formation which initiate the mitotic program
(Andersen, 1999; Kimura et al., 1998). Activation of the DNA replication or DNA damage
checkpoints stabilizes inhibitory phosphorylations on mCDK and thus prevents mitosis
(reviewed in (Harper and Elledge, 2007)). A third checkpoint system, known as the mitotic
9
checkpoint complex (MCC), surveys the appropriate attachment of chromosomes to the
segregation machinery (reviewed in (Musacchio and Salmon, 2007)). Only after every
chromosome pair is securely attached does the MCC activate the anaphase-promoting complex
(APC), a ubiquitin ligase whose targets enable further progression through mitosis (Hwang et al.,
1998; Kim et al., 1998). The mCDK and MCC thus stand as gate-keepers of two crucial mitotic
decision points (Figure 1.1). Furthermore, each complex is extrinsic to the process it regulates
and not a component of the machinery that actually executes the biochemistry of chromosome
segregation and division. Perhaps for these reasons, mCDK and MCC are widely conserved
mediators of eukaryotic checkpoint control.
interphase
telophsae/
cytokinesis
cell membrane
nucleus
stalled
replication
DNA
d image
(Q~
SDNA
prophase
lob.
anaphase
methaphase
mimicl
spindle
tensroT
I0
Figure 1.1. Mitotic checkpoints in eukaryotes. The entry into prophase, the first phase of mitosis, is enabled by the
mitotic cyclin-dependent kinase (mCDK) in complex with a cyclin. DNA damage or stalled replication forks prevent
mitotic entry by activating a signaling pathway, which includes the kinase chkl, that introduces inhibitory
phosphorylations on mCDK. A second checkpoint system prevents progression through mitosis in response to
improper chromosome attachment to the segregation machinery. Once the chromosomes are firmly attached, a
tensive force inactivates the mitotic checkpoint complex (MCC) which normally represses activity of the anaphase
promoting complex (APC). Once activated, APC causes the degradation of securin which releases separase to free
the sister chromatids and allow chromosome segregation during anaphase. Figure adapted from (Harper and Elledge,
2007; Musacchio and Salmon, 2007).
10
The relationship between DNA replication and cell division in bacteria is much less well
understood. The bacterial DNA damage response has been studied for decades, and there are
now examples in several species of DNA damage checkpoints that prevent cell division.
However, these checkpoint systems are poorly conserved and are mechanistically understood
only in the y-proteobacterium E. coli. Thus, it remains unclear (1) how bacteria outside the yproteobacteria regulate division in response to DNA damage and (2) whether the general
strategies and principles of regulatory control are conserved even though the regulators
themselves are not. I have pursued these questions in the a-proteobacterium Caulobacter
crescentus, identifying and subsequently characterizing two damage-induced cell division
inhibitors which are functionally distinct from those found in E. coli and other organisms. Before
describing these new regulators, I will first introduce bacterial cell division, known examples of
how bacterial division is temporally regulated, the canonical DNA damage response systems in
bacteria, and finally the model organism employed in my work, Caulobacter.
Bacterial cell division
Bacteria display tremendous morphological diversity, taking the form of spheres, rods and
spirals and varying in size by over three orders of magnitude. The bacterial envelope is similarly
diverse, as evolutionary pressures to survive and interact with different environments have
yielded variations in the thickness and composition of the cell wall, the presence or absence of an
outer membrane and the type and number of accessory proteins and lipids. Despite these
differences, most bacteria divide in generally the same fashion: A division site at mid-cell is
chosen, a multi-protein cell division complex is constructed, and constriction of one or both cell
membranes is accompanied by the synthesis of a new peptidoglycan cell wall, known as the
"septum", which divides the cytoplasm into two compartments and serves as the new poles for
daughter cells following cytokinesis (Figure 1.2; reviewed in (Egan and Vollmer, 2013;
Lutkenhaus et al., 2012)). Here, I will describe the constituents, assembly and function of the cell
11
division machine which will provide a foundation for understanding known and prospective
modes of division regulation.
FtsZ ring
assembly
"divisome"
assembly
Z-ring constriction,
septum construction
OM
0 000
Q
*
L Jl
IM
Figure 1.2. An overview of bacterial cell division. The first step in bacterial cell division involves polymerization of
the divisome scaffold FtsZ into a "Z-ring" at mid-cell on the cytoplasmic surface of the inner membrane (IM). Next,
divisome components are recruited to the Z-ring forming a multi-protein cell division machine. Each divisome
component is labeled with the last letter in its gene name such that "Z" denotes FtsZ, "A" denotes FtsA, etc...
Following divisome assembly in Gram-negative bacteria, the IM, peptidoglycan (PG) cell wall and outer membrane
(OM) constrict sharply at mid-cell, and cytokinesis ultimately produces two daughter cells. Adapted from (Vicente
and Lwe, 2003).
Identification of cell division genes
Members of the division machinery, known as the "divisome", were originally identified in
E. coli through screens for thermosensitive mutations that prevented division and caused
filamentation without disrupting DNA synthesis or chromosome segregation at the restrictive
temperature. Genes harboring these mutations were called filamentous temperature-sensitive or
"fis"genes. The first division genes to be identified in this fashion wereftsA (Van de Putte et al.,
1964) andftsZ (Hirota et al., 1968), which were later mapped to neighboring locations near the 2
minute position on the E. coli linkage map (Wijsman, 1972). Subsequently,ftsI (Suzuki et al.,
12
1978),ftsQ (Begg et al., 1980),ftsW(Ishino et al., 1989), andftsL (Guzman et al., 1992) were
mapped to the same 2 minute cluster, whileftsB (Ricard and Hirota, 1973),ftsN (Dai et al., 1993)
andftsK (Begg et al., 1995) were found elsewhere in the genome. Remarkably, despite vast
differences in envelope properties across bacterial species, these core division genes are
conserved in most bacteria that have not undergone significant periods of genome reduction.
Many bacterial genomes contain additional division proteins whose essentiality and/or
conservation differ, but the bulk of the divisome is derived from a common origin.
Several early lines of evidence suggested that among the many division genes, FtsZ was a
critical player. Genetic experiments in E. coli demonstrated that FtsZ participated in an early cell
division stage asftsZ mutants were unable to form membrane invaginations at mid-cell unlike
mutants in other division genes (Begg and Donachie, 1985). Subsequently, FtsZ levels were
determined to be rate-limiting for division (Bi and Lutkenhaus, 1990), and apparent FtsZ rings,
or "Z-rings", were found localized to mid-cell by immuno-electron microscopy (Bi and
Lutkenhaus, 1991). It is now appreciated that Z-ring formation on the cytoplasmic surface of the
inner membrane is the earliest known event in bacterial cell division. At mid-cell, the Z-ring
determines the future division site by serving as a scaffold upon which the remainder of the
divisome is constructed (reviewed in (Goehring and Beckwith, 2005)). Below, I will discuss the
mechanisms by which cells position the Z-ring and the general stages of divisome assembly.
FtsZ Positioning
FtsZ is a homolog of the eukaryotic microtubule-forming monomer, tubulin (Nogales et al.,
1998). Although they share little sequence identity, both proteins polymerize into long head-totail filaments in the presence of GTP (Mukherjee and Lutkenhaus, 1994). Despite its initial
appearance as a continuous closed loop, superresolution microscopy in E. coli, B. subtilis, and
Caulobacterhas revealed a Z-ring that is formed by discontinuous bundles of proto-filaments
with extensive lateral associations (Fu et al., 2010; Li et al., 2007; Strauss et al., 2012). In order
13
to correctly position Z-rings, many bacteria employ a strategy whereby concentrations of FtsZ
inhibitors are highest at the cell poles and lowest at mid-cell. In E. coli and B. subtilis, MinC
prevents polymerization of FtsZ at the poles, and its localization is mediated by the membrane
tether MinD which associates with a third topological factor (de Boer et al., 1989; Lee and Price,
1993). In E. coli, this factor is MinE, which shuttles from pole-to-pole and stimulates the
membrane release and inactivation of MinD (Fu et al., 2001; Hale et al., 2001; Raskin and de
Boer, 1999). These rapid oscillations result in time-averaged MinC/MinD concentrations that are
lowest at mid-cell. In B. subtilis, MinD is stably recruited to both cell poles by DivIVA, which
accumulates in regions of high membrane curvature (Edwards and Errington, 1997; Lenarcic et
al., 2009). Caulobacterdo not encode MinC/MinD homologs, but a functional analogue exists in
the Z-ring inhibitor MipZ which is directed to the poles by an association with the origins of
replication, which are themselves polarly localized (Thanbichler and Shapiro, 2006). Each of
these systems is essential for directing division events away from the poles, and mutants lacking
components of the MinC/MipZ complexes display an increased frequency of polar division
events that result in nucleoid-free "mini-cells".
A second, complementary system for correctly positioning the Z-ring is known as 'nucleoid
occlusion' whereby Z-ring assembly is prevented on top of nucleoid-dense regions (reviewed in
(Wu and Errington, 2012)). In E. coli and B. subtilis, the nucleoid occlusion factors SlmA and
Noc, respectively, each contain DNA-binding domains that mediate an association with the
chromosome and effector domains that prevent proper Z-ring assembly (Cho et al., 2011; Wu et
al., 2009). Together, mini-cell prevention and nucleoid occlusion reinforce the mid-cell assembly
of Z-rings such that cells lacking both systems display lethal division defects (Wu, Errington,
2004; Bernhardt, de Boer, 2005). However, in addition to its role in spatial organization,
nucleoid occlusion may also partially mediate the temporal coordination of cell division with
DNA replication. As DNA replication and segregation occur concomitantly in bacteria, a
nucleoid-free, and thus Noc/SlmA-free, region at mid-cell is only revealed during late stages of
14
replication and segregation. These nucleoid occlusion effectors could therefore serve as a type of
DNA replication checkpoint, although it could be argued that they are part of a hybrid substrateproduct system given their close association with the nucleoid.
There are no obvious homologs of noc or slmA in Caulobacter,and a nucleoid occlusion
mechanism may be wholly absent given that Z-rings form during a cell cycle stage when DNA
still occupies the mid-cell region. However, it is formally possible that a novel nucleoid
occlusion factor regulates a step in division downstream from FtsZ polymerization, as will be
discussed further below. Interestingly, noc and slmA were both identified in screens for genes
that are synthetically lethal with mini-cell mutants, but a similar screening strategy is not
possible in Caulobactergiven that mipZ is essential. It thus remains unclear whether nucleoid
occlusion or yet other unidentified systems affect the positioning and temporal control of
division in Caulobacter.
Divisome Assembly
The construction of fluorescently-labeled translational reporters has allowed the subcellular
localization of divisome members to be visualized in real-time in live bacterial cells, revealing
unique sets of assembly times and recruitment dependencies for each component. In F. coli, the
divisome assembles in two general stages (reviewed in (Goehring and Beckwith, 2005)). First, a
set of FtsZ-interacting proteins are recruited to the mid-cell where they promote the formation of
the Z-ring. The essential FtsZ interactors FtsA and ZipA tether FtsZ proto-filaments to the inner
membrane while the non-essential FtsZ association proteins (ZapA-D) may promote Z-ring
bundling and integrity (Dajkovic et al., 2010; Durand-Heredia et al., 2012; Ebersbach et al.,
2008; Galli and Gerdes, 2010; Gueiros-Filho and Losick, 2002; Hale and de Boer, 1997; Low et
al., 2004; Pichoff and Lutkenhaus, 2005). The Caulobactergenome harbors no ZipA homolog,
but does contain FzlA, a protein conserved in a-proteobacteria with a role in promoting the
curvature of FtsZ filaments (Goley et al., 201 Ob). FtsE is another early-arriving FtsZ interactor
15
that, together with FtsX, may coordinate the activity of peptidoglycan-remodeling amidases with
FtsZ dynamics (Meisner et al., 2013; Yang et al., 2011). The FtsEX complex is essential in
Caulobacter(Christen et al., 2011) but in E. coli is required only under low-salt conditions (de
Leeuw et al., 1999).
Following the establishment and stabilization of the Z-ring, the second stage of divisome
assembly involves the recruitment of the remaining divisome components, most of which are
single or multi-pass transmembrane proteins (reviewed in (Errington et al., 2003)). Studies in E.
coli that assessed the ability of individual components to localize in the presence or absence of
other components, suggested a linear recruitment hierarchy in the order FtsZ > ZipA/FtsA >
FtsK > FtsQ > FtsB/FtsL > FtsW > FtsI > FtsN (Goehring et al., 2006; Goehring et al., 2005;
Vicente and Rico, 2006). In this hierarchy, the recruitment of each divisome member requires the
proper assembly of all upstream factors. In B. subtilis, assembly may follow a more cooperative
rather than sequential model (Errington et al., 2003), and even in E. coli, there is evidence that
sub-complex formation may in some cases precede recruitment to mid-cell (Buddelmeijer and
Beckwith, 2004; Fraipont et al., 2011). Furthermore, in all organisms surveyed, the pair-wise
interaction map of divisome components, as revealed by bacterial two-hybrid experiments,
appears not as a linear sequence but as a complex web (Daniel et al., 2006; Di Lallo et al., 2003;
Karimova et al., 2005; Maggi et al., 2008). While some of these interactions may be required for
downstream steps in divisome constriction and septum-synthesis, others may reflect a more
nuanced reality of divisome assembly. In Caulobacter,divisome components assemble in
roughly the same order as E. coli, although FtsA is recruited late in the cell cycle, at a time when
the Z-ring has already been well established (Goley et al., 2011). Localization dependencies in
Caulobacterwere also observed to be much less strict, as very few proteins other than FtsZ and
FtsL were individually required for the recruitment of any downstream component. Nonetheless,
the assembly of the bacterial divisome likely requires a combination of sequential and
cooperative protein-protein interactions.
16
The execution of cytokinesis
Following site-selection and assembly, the divisome must somehow orchestrate the final
stage of division which involves constriction of the membrane(s), synthesis of the septal cell wall
and the separation of daughter cells. While most, if not all, division proteins play some role in
divisome assembly by recruiting downstream components, it is less clear which proteins possess
the additional activities that are specifically required for cytokinesis. In contrast to the steady
accumulation of data describing the dynamics of divisome assembly, assigning functions to
individual divisome components has progressed more slowly, owing in part to the difficulty in
designing and performing biochemical assays on a multi-component and membrane-bound
machine. Nonetheless, many enzymatic and other functional domains have been characterized
and domains of unknown but essential functions have been isolated. Here, I will summarize the
known and putative functions for individual divisome components and speculate on how they
might cooperate to perform cytokinesis.
FtsZ and Z-interactors: As mentioned, FtsZ is the first recruit to the divisome and is thus
involved in division site selection and the recruitment of all subsequent division proteins. ZipA
(in E. coli) and FtsA tether FtsZ to the inner membrane, and ZapA-D, FzlA, FzlC have been
implicated in regulating Z-ring properties such as stability, lateral interactions and filament
curvature.
FtsK: FtsK is a four-pass transmembrane protein with a C-terminal DNA translocase domain that
facilitates the segregation of replicated chromosomal termini (Aussel et al., 2002). In E. coli, this
translocation function is non-essential (Wang and Lutkenhaus, 1998), and may only be required
to resolve the formation of chromosome dimers or during certain stressful conditions (Britton
and Grossman, 1999; Sivanathan et al., 2009; Yu et al., 1998). The essential function of FtsK
resides within its transmembrane domains (Draper et al., 1998; Dubarry et al., 2010) and can be
bypassed by activating mutations in FtsA or overexpression of theftsQAZ operon (Geissler and
17
Margolin, 2005). In Caulobacter,the C-terminal translocase domain is essential, possibly
because it is involved in recruiting the topoisomerase IV subunit ParC to the replisome and thus
coordinating DNA segregation and replication (Wang et al., 2006).
FtsQLB: In E. coli, FtsQLB is thought to form an essential subcomplex prior to its recruitment to
the mid-cell (Buddelmeijer and Beckwith, 2004). Each component is a bitopic membrane protein
with cytoplasmic and periplasmic domains, and together, they have been shown by bacterialtwo-hybrid experiments to form multiple interactions with most other divisome components (Di
Lallo et al., 2003; Karimova et al., 2005). None of these proteins has a known enzymatic or other
function, but in B. subtilis, FtsL is highly unstable and rate-limiting for division, and it has been
implicated in mediating division regulation during replication stress (Bramkamp et al., 2006;
Breier and Grossman, 2009; Goranov et al., 2005). Due to its placement within the recruitment
hierarchy and its ability to serve as a protein interaction hub, it is thought that FtsQLB could
somehow coordinate constriction of the Z-ring with the synthesis of septal cell wall.
FtsWIN: FtsWIN are also thought to form an essential subcomplex within the divisome, and they
are generally among the latest-arriving divisome components (Fraipont et al., 2011; Mercer and
Weiss, 2002; Wissel and Weiss, 2004). Each of these proteins possesses a known or putative
function in constructing or modifying the septal peptidoglycan cell wall. FtsW is a predicted 10pass transmembrane protein that translocates the peptidoglycan precursor Lipid II from the
cytoplasm into the periplasm at mid-cell (Mohammadi et al., 2011). Once in the periplasm, septal
Lipid II is joined into chains by an unknown transglycosylase and then cross-linked by the
transpeptidase activity of FtsI into a rigid, 3D structure (Adam et al., 1997). The function of FtsN
is less clear, although it is thought to both stabilize the divisome and recruit peptidoglycanmodifying enzymes to mid-cell (Goehring et al., 2007; M611 et al., 2010; Peters et al., 2011).
Peptidoglycan-modifiers: Just as new cell wall must be synthesized at the septum, likely by FtsW
and FtsI among other proteins, it must also be simultaneously remodeled and cleaved to allow
18
new pole formation and daughter cell separation. In E. coli, the amidases AmiA, AmiB and
AmiC are thought to selectively remove peptidoglycan cross-links and facilitate cell wall
remodeling and cleavage (Heidrich et al., 2001). Although these amidases are individually
dispensible, cells lacking all three are defective for separation and exhibit a chaining phenotype.
In Caulobacter,homologs of AmiABC are absent, but a putative endopeptidase DipM is
involved in remodeling septal peptidoglycan (Goley et al., 2010a; M611 et al., 2010; Poggio et al.,
2010).
Tol-Pal: In Gram-negative bacteria, the outer membrane must invaginate in tandem with the cell
wall and inner membrane. In E. coli, the Tol-Pal system is a five protein complex that localizes
to mid-cell and forms contacts with all three envelope layers (Gerding et al., 2007). Although the
Tol-Pal system is not essential in E. coli, it is essential in Caulobacterand likely facilitates
proper invagination of the outer membrane in both organisms (Yeh et al., 2010).
What triggers cytokinesis?
After, or perhaps shortly before, the division machine is fully constructed, it must shift
from a state of assembly into one of septum-synthesizing activity and envelope constriction. In
tandem with these cytokinetic activities, the divisome must remodel itself in order to account for
its own shrinking circumference and the dynamic nature of its substrate. The molecular
mechanism that triggers this change from static assembly to dynamic activity is largely
unknown. In E. coli, the onset of constriction temporally coincides with the arrival of the latest
recruited divisome component FtsN (Gerding et al., 2009), and logically, the arrival of a latearriving component would serve as a useful trigger, indicating that the assembly stage is near
completion. However, it remains unclear what set of protein-protein interactions and
conformational changes within the divisome constitute this triggering pathway.
19
Once the decision to divide has been made, the divisome is faced with the task of cutting
the cell in half, an energy-intensive process that must be tightly controlled to avoid envelope
rupturing. As mentioned, some divisome components likely play roles in septum synthesis,
membrane constriction and DNA segregation, while others may be required only for structural
support and divisome assembly. An overlapping and perhaps broader question is which of these
proteins, subcomplexes and/or activities actually provides the force and directionality for
constriction? Theories as to which divisome component(s) is in the driver's seat can be generally
grouped into two categories (Errington et al., 2003):
FtsZ-centric: FtsZ forms proto-filaments in the presence of GTP that bend following GTP
hydrolysis in vitro (Lu et al., 2000). It has therefore been proposed that FtsZ could
simultaneously translate the force supplied by GTP hydrolysis into the directionality of inward
invagination. In Caulobacter,FzlA may enhance the constrictive force by increasing FtsZ
bending during successive rounds of FtsZ assembly, GTP hydrolysis, filament breakage and
reassembly (Goley et al., 201 Ob). In support of these models, a membrane targeted FtsZ fusion
can itself form Z-rings and mediate constrictions in lipid vesicles (Osawa et al., 2008, 2009).
Furthermore, Mollicutes and artificially derived L-form bacteria retain functional FtsZ but
possess no cell walls (Margolin, 2005; Onoda et al., 2000; Siddiqui et al., 2006), perhaps
together suggesting that FtsZ itself provides the cytokinetic force. In a related model, FtsZ protofilaments could slide inwards against the ATPase FtsA which would serve as a motor (Feucht et
al., 2001; Paradis-Bleau et al., 2005). Intriguingly, gain of function mutations inftsA have been
identified which cause a small-cell phenotype, indicative of a propensity to hyperactively divide
(Geissler et al., 2007). These results suggest that FtsZ, perhaps in association with FtsA, could
drive constriction and effectively drag the rest of the divisome along with it.
Cell wall-centric: It is alternatively possible that division is advanced by the inward growth of
the septal cell wall. Accordingly, FtsI and the peptidoglycan remodeling proteins could provide
20
the motive force for division while the Z-ring would be required mainly for correct positioning
of the divisome. GTP hydrolysis-defectiveftsZ mutants are still capable of division suggesting
that other components may indeed provide the constrictive force (Bi and Lutkenhaus, 1992;
Dajkovic and Lutkenhaus, 2006; Mukherjee et al., 2001). Interestingly, the chlamydiae lack
FtsZ, and while they also lack a cell wall, some peptidoglycan synthesis proteins are retained,
and division can still be inhibited by the FtsI transpeptidase inhibitor penicillin (Ghuysen and
Goffin, 1999; McCoy and Maurelli, 2006). It is unclear whether the existence of Z-less bacteria
indicates a less integral role for FtsZ in divisome constriction in other clades.
The temporal regulation of cell division
Cell division must be temporally regulated such that it follows DNA replication and
chromosome segregation during each cell cycle. Additionally, division must be transiently
delayed during stressful conditions that perturb the natural replication cycle or otherwise disfavor
division. In order to achieve such temporal control, regulation could theoretically occur at any
step in divisome assembly, such that the recruitment of essential downstream components is
prevented until division is desired. Alternatively, any of the many essential enzymatic,
biosynthetic or other divisome activities could serve as targets for temporal regulation, either by
inhibition until, or activation at, the appropriate time. Many temporal regulators of division have
now been found in several model systems, and indeed, assembly and activity steps can both
serve as regulatory nodes. Here, I will summarize our understanding of temporal division
regulators and discuss the common regulatory strategies that have emerged.
Control of cell division during the cell cycle
The means by which bacterial cells coordinate cell division with DNA replication and the
broader cell cycle are poorly understood. One potential mechanism could be the cell-cycle
controlled expression of division genes such that divisome components are only made available
21
when needed. In E. coli,ftsZ is transcribed by at least six promoters (Flardh et al., 1997), and its
mRNA levels oscillate with the cell cycle, peaking during the initiation of DNA replication when
Z-rings are formed (Garrido et al., 1993). Because FtsZ polymerization is concentrationdependent in vitro (Romberg and Levin, 2003), titrating FtsZ levels during the cell cycle could
be an effective mode of control. However, FtsZ protein levels are largely unchanged during
steady-state growth in E. coli and B. subtilis (Weart and Levin, 2003), and artificially increasing
or decreasing FtsZ levels does not significantly affect the timing of division (Palacios et al.,
1996; Ward and Lutkenhaus, 1985; Weart and Levin, 2003). Furthermore, a dependency of FtsZ
levels on DNA replication has not been observed in B. subtilis (Rowland et al., 1997), and while
a slight dependency may exist in E. coli, its relevance and mechanism are unclear (Liu et al.,
2001). In B. subtilis, a detailed analysis of transcriptional fusions to nine divisome components
revealed minimal cell cycle-dependent oscillations (Trip et al., 2013), and similar results have
been observed at the protein level for several components in E. coli (Rueda et al., 2003). These
results together suggest that the division cycle is unlikely to be enforced by oscillations in
divisome component availability.
A more likely model is that during each cell cycle, division proteins are regulated at the
level of activity, localization, or by post-translational modification. The nucleoid occlusion
proteins SlmA and Noc prevent Z-ring formation until late stages of DNA segregation and could
in theory serve as cell cycle checkpoints that regulate FtsZ post-translationally (Bernhardt and de
Boer, 2005; Wu and Errington, 2004). However, cells lacking either factor are unaffected during
vegetative growth, suggesting that other factors can compensate to correctly coordinate Z-ring
formation with the cell cycle. Intriguingly, nucleoid occlusion may prevent division during
replication fork arrest in B. subtilis cells in a Noc-independent manner suggesting the existence
of additional nucleoid occlusion proteins (Bernard et al., 2010). Another attractive candidate for
enforcing the dependency of division on replication is the division protein FtsK whose Cterminal cytoplasmic domain participates in late stages of chromosome segregation. FtsK could
22
thus coordinate the status of DNA segregation with divisome assembly given that it is required
for the localization of several downstream components (Chen and Beckwith, 2001). Despite
these features, there is no evidence of an actual replication-division dependency enforced by
FtsK, and the dispensability of its C-terminal domain suggests that a role in checkpoint control is
unlikely. It thus remains unclear whether bacteria employ substrate-product relationships,
checkpoints or some other form of control to ensure the proper timing of division within the cell
cycle.
Conditionalcontrol of cell division
One remarkable feature of bacterial physiology is the ability to rapidly adjust intracellular
activities in response to a changing environment. Bacterial cells are equipped with sensory
proteins that can detect a wide variety of internal and external cues and mount the appropriate
regulatory response through mediators and effectors. Many of these regulatory systems modulate
the bacterial cell cycle, and some impinge directly on the cell division machinery. For instance,
many cells regulate their size in response to nutrient availability. In B. subtilis and Caulobacter,
nutrient-rich conditions activate cell division inhibitors resulting in longer division cycles and
larger cells. In B. subtilis, the metabolite uridine-5'-diphosphoglucose (UDP-Glc) binds and
activates UgtP which in turn prevents FtsZ assembly (Weart et al., 2007). Similarly, Caulobacter
cells detect the metabolite NAD(P)H by a sensor unrelated to UgtP called KidO, which somehow
targets FtsZ to delay division (Radhakrishnan et al., 2010). In each case, a single protein surveys
the levels of an internal metabolite and appropriately regulates cell division through an
interaction with FtsZ. Nutrient-influenced size control has also been documented in E. coli,
although the metabolite sensed and mode of regulation are unknown (Donachie and Begg, 1989;
Reeve et al., 1984).
Many Gram-positive bacteria can produce dormant but resilient cells known as "spores"
in response to starvation. The developmental program resulting in sporulation applies several
23
unique constraints on cell division, the first of which involves an asymmetric division event that
creates a small forespore and a large mother cell (reviewed in (Errington, 1993)). In B. subtilis,
FtsZ is redistributed from its normal position at the mid-cell to the cell poles by an association
with the multi-functional membrane protein SpoIIE (Beall and Lutkenhaus, 1991; Khvorova et
al., 1998; Levin et al., 1997; Lucet et al., 2000). Following the asymmetric division, the mother
cell engulfs and nurtures the spore until its ultimate lysis when the spore is released. Importantly,
the mother cell is terminally differentiated and its activities are dedicated solely to spore
development. To help ensure this exit from the cell cycle, B. subtilis encodes a division inhibitor
mciZ, which specifically inhibits FtsZ polymerization in the mother cell (Handler et al., 2008). S.
coelicolor is another sporulating bacterium which lives normally as a multi-nucleate filament
that does not require FtsZ for normal growth and viability (McCormick et al., 1994). However,
following nutrient limitation, S. coelicolor filaments assemble Z-rings at multiple division sites
in order to create chains of spores (Schwedock et al., 1997). Intriguingly, the membraneassociated factor SsgB recruits FtsZ to these division sites and is the only known example of a
divisome component whose localization precedes that of FtsZ (Willemse et al., 2011).
Many types of stress that do not result in a new developmental program like sporulation
can nonetheless require cell division to be regulated. The oxidative stress response in M
tuberculosis causes FtsZ-interacting protein A, FipA, to bind FtsZ, allowing division to occur
and sustaining growth during macrophage engulfment (Sureka et al., 2010). In E. coli, the nonessential divisome component FtsP may stabilize the divisome during oxidative stress (Samaluru
et al., 2007). AlthoughftsP mutants can be suppressed by mutations that stabilize Z-ring
assembly, its direct binding partners are not known. FipA and FtsP are examples of proteins that
promote division during oxidative stress, a condition where protein damage poses a barrier to
division. A more common stress response involves the prevention of division until the stress has
been resolved. During the stringent response to amino acid starvation, the alarmone (p)ppGpp
may somehow prevent FtsZ transcription in E. coli (Powell and Court, 1998). During stationary
24
phase in E. coli, a condition when cells are likely starved and subjected to changing pH, the
RNA-binding protein Hfq prevents minicell formation by down-regulating FtsZ expression
(Takada et al., 1999). Similarly, during stationary phase in Caulobacter,FtsZ levels are kept low
although the mechanism is unknown (Wortinger et al., 1998). Division is likely regulated by the
heat shock response machinery (Tsuchido et al., 1986), and there are many other types of stress
during which cells arrest the division cycle, although the regulators and their targets have not
been identified.
positioning factors
starvation/
stationary phase
regulators
MP
sporulation
regulators
eaoi
reltaors
rgltr
FtsP
oxidative
stress
stabilizers
divisome stabilizers/
shapers
Figure 1.3. Temporal and spatial regulators of division converge on FtsZ. Each of the Z-ring regulators are grouped
based on their proposed function and/or regulatory category. Z-ring inhibitors are shown above FtsZ and positive
regulators are shown below. Figure adapted from (Kirkpatrick and Viollier, 2011).
Within single bacterial cells, multiple regulators of cell division together ensure the
proper timing of divisions both during the normal cell cycle and in response to a wide variety of
changing environmental conditions. Although we have likely found only a small fraction of these
regulators, it is clear that the most common regulatory strategy involves targeting FtsZ at the
transcriptional, translational, or post-translational levels (Figure 1.3). Many division inhibitors
25
prevent Z-ring formation and thus disrupt all subsequent steps in divisome assembly. This is
perhaps the surest way to completely abolish any chance of division. However, many stresses are
ephemeral and cells requiring a transient delay in cell division might not want to invest in the
complete disassembly and re-assembly of the division machine. DNA damage is one such stress
given that cells frequently encounter many types of DNA lesions, some of which can be quickly
repaired. The bacterial DNA damage response has been studied extensively, and indeed, several
division inhibitors have unique mechanisms compared to those already discussed. In the next
section, I will introduce the DNA damage response in bacteria and examine its associated modes
of division regulation.
The DNA damage response
DNA damage poses a direct challenge to the replication-division dependency, because it
causes replication to slow or stall during periods of repair. It is formally possible that cells could
employ a single regulatory system to coordinate division with replication during both normal cell
cycles and episodes of DNA damage. For instance, nucleoid occlusion might suffice to
accommodate DNA damage and any stress that similarly prolongs replication and leaves
unreplicated DNA at mid-cell. However, the canonical bacterial response to DNA damage,
known as the 'SOS response' (reviewed in (Erill et al., 2007)), typically includes the induction of
a specialized cell division inhibitor (Figure 1.4). These inhibitors are not required for normal
viability suggesting that their roles are specifically reserved for encounters with DNA damage.
There is also a growing appreciation that some cells have additional, SOS-independent
mechanisms for damage-induced division control, although these are poorly understood. Here, I
will introduce the bacterial response to DNA damage and discuss the known modes of division
regulation.
26
The SOS regulatorsRecA and LexA
The first hints of a multifunctional bacterial DNA damage response came nearly fifty
years ago in a series of experiments in E. coli. Cells subjected to DNA damage by UV irradiation
formed long filaments (Green et al., 1969), underwent prophage induction (Hertman and Luria,
1967) and exhibited increased rates of mutagenesis (Weigle, 1953). Mutants that mapped to the
lexA and recA loci were sensitive to UV irradiation and defective in each of these UV-induced
activities (Gudas and Pardee, 1975). It was soon proposed that these genes orchestrate an
inducible DNA damage response that requires de novo protein synthesis for full function
(Radman, 1975).
Years of rigorous experimentation have solidified the roles of RecA and LexA as the
gatekeepers of SOS induction. LexA is a transcriptional repressor whose N-terminal DNAbinding domain silences the promoters of SOS genes in undamaged cells (Brent and Ptashne,
1980; Little and Harper, 1979; Little et al., 1981). Following DNA damage, LexA undergoes a
RecA-dependent autocatalytic cleavage which de-represses the regulon and allows the induction
of SOS genes (Lin and Little, 1989; Little et al., 1980). RecA is a DNA-dependent ATPase and
homolog of the eukaryotic repair gene Rad51 (Ogawa et al., 1979; Roberts et al., 1979).
Following DNA damage, RecA is activated by the presence of single-stranded DNA (ssDNA)
which can arise in several ways (reviewed in (Indiani and O'Donnell, 2013)). When DNA
polymerase is paused at lesion sites, continued activity of the replicative helicase could unwind
DNA revealing stretches of ssDNA. Alternatively, the replisome could skip lesion sites and
resume replication downstream, leaving behind ssDNA gaps. DNA repair can also generate
ssDNA ends following the enzymatic processing of double-strand break ends or ssDNA gaps
during excision repair. Once bound to ssDNA, RecA polymerizes into filaments along the DNA
whose assembly requires ATP binding (Lee and Cox, 1990). The ATP-bound RecA filament
represents its active form (RecA*) which performs two essential functions in DNA repair. As a
27
recombinase, RecA* catalyzes strand exchange and mediates homology searching during the
recombination-based repair of double-strand breaks and damaged single-strand gaps (McEntee et
al., 1979). Its second function, likely located at a distinct site within RecA* (Adikesavan et al.,
2011), is to bind LexA and promote its cleavage.
The SOS regulon
Microarray experiments indicate that hundreds of genes are regulated by DNA damage,
but only a subset are bonafide SOS members (Au et al., 2005; Fernindez De Henestrosa et al.,
2000). The E. coli SOS regulon is comprised of 40 genes that are directly repressed by LexA
(reviewed in (Kelley, 2006)). Among these, RecA and LexA themselves are SOS-induced. As
mentioned, RecA mediates recombination repair and, with Ssb, stabilizes stalled replication forks
(Buss et al., 2008), while newly synthesized, uncleaved LexA is required to shut off the SOS
system when the damage has been fully repaired (Little, 1983). Many SOS genes are involved in
DNA damage repair, as UvrABCD and RuvCAB participate in nucleotide excision and
recombination-based repair pathways respectively (Sancar and Rupp, 1983; Seigneur et al.,
1998). PolB, DinB, and UmuDC constitute another class of SOS genes involved in translesion
DNA synthesis (Yeiser et al., 2002). These polymerases catalyze error-prone base additions that
sacrifice fidelity for an ability to proceed through DNA lesions. The final class of SOS genes are
the checkpoint genes which prevent cell cycle progression until the damage has been removed.
In E. coli, SulA prevents cell division and will be discussed further below, while UmuDC delays
DNA replication before it undergoes a RecA-mediated autocleavage that activates its function in
translesion synthesis (Opperman et al., 1999).
Most sequenced bacterial genomes contain homologs of LexA, although LexA binding
motifs are widely divergent between clades (Erill et al., 2007). The motif in most Gram-positive
bacteria, GAAC-N4-GTTC, and that for the P- and y-proteobacteria, CTGT-N8-ACAG, are both
palindromic sequences that bind LexA dimers with varying hinge angles to accommodate the
28
different spacer lengths (Cheo et al., 1991; Erill et al., 2003). Unexpectedly, the motif in aproteobacteria, GTTC-N7-GTTC, is a direct repeat instead of a palindrome, indicating a
completely novel dimer interface and binding mechanism (Fernindez de Henestrosa et al., 1998).
Other oddities have emerged as more LexA binding motifs have been sequenced, including some
that are asymmetrical and imperfectly palindromic (Campoy et al., 2003; Campoy et al., 2002;
Yang et al., 2002). Despite these marked differences in binding sites, a similar set of core SOS
genes are commonly found throughout the bacterial kingdom, including lexA, recA, uvrA,
ruvCAB and ssb (Erill et al., 2007). Interestingly, very few bacteria contain all of these core
genes, and more recently, the in silico analysis of a highly conserved SOS-inducible operon
encoding the mutagenesis genes imuAB and dnaE2 has led to questions regarding which genes
are the oldest SOS members (Abella et al., 2004). Although imuAB-dnaE2 are not found in B.
subtilis or E. coli, their tight genetic and physical linkage with lexA in most other phyla suggests
that the SOS response may have originally been an inducible translesion synthesis system (Erill
et al., 2006).
Regardless of what the ancestral SOS regulon looked like, it is clear that many bacteria
respond to DNA damage by inducing a similar set of regulatory and effector genes. The SOSinduced division inhibitors are a notable exception as those characterized to date show very little
sequence homology. Given that these genes are likely unrelated functional analogs, have they
nevertheless converged to inhibit division in similar ways, by targeting the same components
and/or activities? And, do they, like most of the known positional and conditional regulators of
division, also target FtsZ and prevent formation of the Z-ring?
SOS-induced division inhibitors
29
The E. coli SOS member sulA has long been the lone example of a bacterial DNA damageinduced cell division inhibitor. sulA mutations were originally identified as suppressors of lossof-function mutants in the protease lon, which normally exhibit irreversible filamentation
following exposure to UV light (George et al., 1975). The same screen yielded suppressor
mutations inftsZ, which encodes the target of SulA. During SOS-induction, SulA accumulates
and prevents division by sequestering the free ends of FtsZ monomers or proto-filaments and
blocking the polymerization of additional subunits (Bi and Lutkenhaus, 1993; Cordell et al.,
2003; Mukherjee et al., 1998). De novo Z-ring formation is thus prevented, and extant Z-rings
are dissolved due to the highly dynamic nature of FtsZ filament assembly and disassembly
(Stricker et al., 2002). Following DNA repair, Lon degrades SulA and allows division cycles to
resume (Figure 1.4; (Mizusawa and Gottesman, 1983)).
DNA damage
FtsZ
0.0D
RecA
FtsZAA
------ +
ng
Figure 1.4. SOS induction of the division inhibitor sulA. In undamaged cells, the repressor LexA silences
transcription at the promoters of SOS genes. Following DNA damage, RecA forms active filaments on singlestranded DNA and causes the autocatalytic cleavage of LexA. SOS genes are de-repressed, and SulA, an SOSinduced division inhibitor in E. coli, prevents polymerization of FtsZ. After the damage is repaired, the Lon protease
degrades SulA allowing Z-ring formation.
30
The existence of SulA initially supported the notion that most, if not all regulators of
bacterial cell division target FtsZ. However, in the last decade, several SOS-induced division
inhibitors with seemingly novel mechanisms have been identified in Gram-positive bacteria.
YneA is a strongly SOS-induced cell division inhibitor in B. subtilis that shares few other
characteristics with its E. coli counterpart (Kawai et al., 2003). While SulA is a cytoplasmic
protein like its target FtsZ, YneA is a single-pass transmembrane protein with an extracellular
LysM peptidoglycan-binding domain. There is no evidence that YneA interacts with FtsZ in
vitro, and Z-ring formation is reduced but not abolished during YneA overproduction. While its
precise target and mechanism are unknown, the transmembrane and extracellular domains are
required for its function suggesting that it may interact with the many divisome components
downstream from FtsZ that have transmembrane domains or the septal cell wall itself (Mo and
Burkholder, 2010). The SOS-regulated division inhibitors ChiZ and DivS have been identified in
the Gram-positive Actinobacteria M tuberculosis and C. glutamicum respectively, and both of
these are single-pass transmembrane proteins like YneA, although they share limited sequence
similarity (Chauhan et al., 2006; Ogino et al., 2008). ChiZ contains a LysM domain, interacts
with FtsI and FtsQ by bacterial-two-hybrid analysis and exhibits cell wall hydrolase activity;
similar mechanistic insights are lacking for DivS (Vadrevu et al., 2011). Both inhibitors may
have mild effects on Z-ring formation, but as with YneA, evidence for a direct interaction with
FtsZ is lacking.
Although detailed mechanisms are lacking for the Gram-postive division inhibitors, it is
clear that they do not completely disrupt Z-ring assembly like SulA. This distinction could be a
testament to the inherent differences between the envelopes of Gram-negative and Gram-positive
bacteria. The Gram-positive cell wall is two- to eight-fold thicker than that of Gram-negatives,
and it is enriched in teichoic acids which are polysaccharides that provide the cell wall with
increased rigidity (Wicken and Knox, 1975). Division in Gram-positive bacteria may thus follow
a more cell wall-centric mode of constriction, and inhibitors of division could target divisome
31
components and activities more intimately linked to peptidoglycan synthesis and remodeling.
Given that so few division inhibitors have been identified, it is difficult to say whether
distinctions among the current group reflect fundamental differences in bacterial physiology. As
described in Chapter 2, I have sought to identify and characterize the SOS-induced division
inhibitor in the c-proteobacterium Caulobactercrescentus, in part to determine whether SulA
and its target FtsZ truly serve as a paradigm for regulatory control in Gram-negative bacteria.
SOS-independent regulation
Several lines of evidence have suggested that in E. coli, SulA may not be the sole means
of damage-induced division inhibition. Cells lacking sulA and those unable to induce SOS genes
still exhibit filamentation following replication arrest or DNA damage (Burton and Holland,
1983; Hill et al., 1997; Howe and Mount, 1975; Huisman et al., 1980; Ishioka et al., 1997; Jaffe
et al., 1986; Liu et al., 2001). Similar results have been observed in B. subtilis (Love and Yasbin,
1984), and in M tuberculosis, ChiZ induction is not completely eliminated in cells lacking recA
(Rand et al., 2003). These results suggest that SOS-independent modes of division inhibition
likely exist, but the associated regulators and inhibitors have not been identified.
How do cells respond to DNA damage if not by the RecA-ssDNA-mediated induction of
SOS genes? Exposed stretches of ssDNA are one of many abnormal DNA structures that could
serve as DNA damage signals for regulatory systems. In B. subtilis, the di-adenylate cyclase
DisA monitors genome integrity and likely binds to branched DNA structures that arise during
recombination-based repair (Bejerano-Sagie et al., 2006). When paused at such sites, the diadenylate cyclase activity of DisA is inhibited causing decreases in cellular levels of cyclic-diAMP (c-di-AMP). c-di-AMP is a secondary messenger that in undamaged cells activates SpoOA,
a master regulator involved in triggering sporulation (Oppenheimer-Shaanan et al., 2011; Witte
et al., 2008). DisA is thus an example of a checkpoint system whose activity is regulated by a
non-canonical DNA damage-associated structure.
32
A second possibility is that a separate regulatory system is tasked with responding directly
to replication stress which is intimately but not exclusively linked to DNA damage. For instance,
the universally conserved replication initiation factor DnaA has been proposed to function as
both a sensor of replication stress and a mediator of response elements. During replication
initiation, DnaA-ATP multimers promote double-stranded DNA melting at the origin and recruit
replisome components to the nascent replication forks (Bramhill and Kornberg, 1988). DnaAATP also functions as a transcription factor for genes involved in replication, transcription,
translation, division and, in B. subtilis, sporulation inhibition (Burkholder et al., 2001; Goranov
et al., 2005; Hottes et al., 2005; Messer and Weigel, 1997). Following replication initiation,
DnaA activity is inhibited by members of the elongation complex in order to prevent
overinitiation (Camara et al., 2005; Katayama and Sekimizu, 1999). When replication is stalled,
transcriptional activities of DnaA are reinstated. Intriguingly, it has been proposed that DnaA
mediates a replication checkpoint on cell division by preventing ftsL transcription in B. subtilis
(Goranov et al., 2005).
In summary, there are likely damage-induced, SOS-independent regulatory modules and
modes of division inhibition in many bacteria. In Chapter 3, I have used Caulobacterto identify
and characterize an SOS-independent inhibitor of cell division.
Caulobacter crescentus as a model system
Several unique features of the Caulobacterlife cycle make it an ideal system for studying
the coordination between DNA replication and cell division (reviewed in (Skerker and Laub,
2004)). Unlike many bacteria, which in rich media can initiate multiple rounds of replication
between consecutive divisions, Caulobacterreplicates once and only once per cell division
resulting in distinct G 1, S and G2 cell cycle stages (Marczynski, 1999). Additionally, pure
populations of GI cells are readily obtained by density centrifugation, facilitating the analysis of
individual cell cycle stages and their associated dependencies (Evinger and Agabian, 1977).
33
Finally, as a genetically tractable a-proteobacterium, Caulobacterallows principles gleaned
from the y-proteobacterium E. coli to be placed in a broader evolutionary context.
swarmer
stalked
predivisional
0 CtrA~P
(2D MipZ
----
--
Z-ring
c!*vo divisome
0
Figure 1.5. The Caulobactercell cycle. Phosphorylated CtrA (CtrA~P) is indicated by blue shading. The Z-ring is
indicated in green, and the fully assembled divisome is indicated in purple. The Z-ring inhibitor MipZ is localized to
the cell poles through an association with chromosomal origins of replication.
Caulobacterhas long been studied as a model system for understanding the molecular
events that drive a bacterial cell cycle (Fig. 1.5). Its mandatory progression through distinct cell
cycle stages with respect to replication is reminiscent of the eukaryotic cell cycle. It is therefore
perhaps not surprising that the Caulobactercell cycle is orchestrated by the oscillating activity of
a phosphorylated regulator, CtrA, much as the eukaryotic cell cycle is orchestrated by cyclindependent kinases (Quon et al., 1996). CtrA is a transcription factor that, when activated by
phosphorylation, binds to DNA and regulates the expression of nearly 100 genes (Domian et al.,
1997; Laub et al., 2002). In GI 'swarmer' cells, CtrA represses replication initiation by binding
to and silencing the origin, which is located at the flagellated cell pole (Quon et al., 1998). At the
end of G 1, CtrA is dephosphorylated and degraded allowing replication to initiate.
Concomittantly, the flagellum is shed and in its place a 'stalk' is built, which facilitates surface
adsorption and nutrient uptake (Stoveoindexter and Cohen-Bazire, 1964). The newly replicated
34
origin is immediately shuttled to the opposite cell pole which establishes the bipolar localization
of the origin-associated Z-ring inhibitor MipZ (Thanbichler and Shapiro, 2006). As S phase
proceeds and the stalked cell elongates, the Z-ring is established at mid-cell where concentrations
of MipZ are lowest (Kelly et al., 1998). Mid-way through S phase, CtrA again accumulates in its
phosphorylated form, allowing the transcription of genes necessary for cell division and other
late cell cycle events. As replication nears completion, the fully assembled divisome causes cells
to constrict at mid-cell during the 'pre-divisional' cell cycle stage (Goley et al., 2011). Finally,
an asymmetric cell division produces a flagellated swarmer cell which resides in G 1 and a
stalked cell which immediately begins another round of replication.
The regulation of cell division in Caulobacter
Microarray experiments on synchronized Caulobactercultures revealed that nearly 500
genes varied during the course of the cell cycle (Laub et al., 2000). Furthermore, many of these
genes were expressed during the cell cycle stage when their activity was required, a phenomenon
termed 'just-in-time' transcription. In early predivisional cells, CtrA transcribes the essential
division genes,ftsQ,ftsA andfts Wat a time when divisome assembly is just underway.
Furthermore, the ctrA loss-of-function phenotype includes extensive filamentation, suggesting
that the transcriptional activation of these divisome components in predivisional cells could be a
prerequisite for division (Quon et al., 1996). CtrA is thus well situated to serve as a cell division
checkpoint, and indeed, its activity appears to be regulated by genome integrity and the status of
DNA replication. Replication initiation is required to somehow activate the hybrid histidine
kinase CckA (Iniesta et al., 2010) which in turn phosphorylates and stabilizes CtrA through the
histidine phosphotransferase ChpT (Biondi et al., 2006). Furthermore, cells treated with DNA
damaging agents or hydroxyurea, which depletes nucleotides pools and stalls replication,
demonstrate reduced CtrA activity (Wortinger et al., 2000).
35
Taken together, these results suggest that in Caulobacter,DNA damage and replication
stress could prevent division by decreasing the activity of the cell cycle regulator CtrA.
However, it is unknown whether the inhibition of CtrA is necessary or sufficient to prevent
division during these conditions.
An alternative and perhaps complementary mode of division control could be provided by
direct division inhibitors activated during the CaulobacterDNA damage response. Following
damage, Caulobacterinduces an SOS regulon similar to that of E. coli in its size and
composition (da Rocha et al., 2008). However, no homologs of sulA or any other division
inhibitor are present, and SOS-induced division inhibitors have not been identified in any uproteobacteria. It thus remains unclear whether u-proteobacterial genomes encode a novel class
of damage-induced division inhibitors, or whether CtrA or other cell cycle genes instead provide
the necessary control.
Research Summary
The subsequent chapters describe a series of experiments that explore the mechanisms of
division inhibition during the CaulobacterDNA damage response. Chapter 2 details the
identification and characterization of sidA, an SOS-induced cell division inhibitor that is
conserved in many a-proteobacteria. sidA encodes a short, 29 amino acid protein that is localized
to the membrane at mid-cell following DNA damage. I provide evidence that SidA does not
prevent Z-ring formation or divisome assembly, but instead inhibits division through a direct
interaction with the late-arriving divisome component FtsW. I show that SidA does not disrupt
the cell wall translocase activity of FtsW suggesting that FtsW likely performs an additional role
in executing division. My work demonstrates that divisome components other than FtsZ can be
targeted by division inhibitors and identifies FtsW as a critical regulatory node in Caulobacter.
36
During our work with sidA, I noticed that cells incapable of mounting an SOS response can
nonetheless delay cell division during DNA damage. In chapter 3, I identify and characterize
didA, an SOS-independent division inhibitor. Like sidA, didA encodes a short transmembrane
protein that does not prevent any step in divisome assembly. DidA interacts with the late-arriving
divisome component FtsN, and intriguingly, mutations infts W andftsI can suppress the activities
of both SidA and DidA. My work suggests that these suppressor mutations do not reduce the
affinities of SidA and DidA for their divisome targets, but instead hyperactively promote cell
division. Finally, I show that cells require at least one of these inhibitors to properly regulate
division during DNA damage illustrating a degree of redundancy within the CaulobacterDNA
damage response. My work uncovers an SOS-independent DNA damage response in
Caulobacterand provides further support for the importance of the FtsW/FtsI/FtsN complex in
triggering division.
In chapter 4, I discuss the implications of our work for understanding bacterial cell division
and its regulation. I also present ongoing work which explores the molecular mechanisms
underlying the SOS-independent regulation of didA.
37
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Yang, M. K., Y. C. Yang, and C. H. Hsu, 2002, Characterization of Xanthomonas axonopodis pv. citri
LexA: recognition of the LexA binding site: Mol Genet Genomics, v. 268, p. 477-87.
Yeh, Y. C., L. R. Comolli, K. H. Downing, L. Shapiro, and H. H. McAdams, 2010, The caulobacter TolPal complex is essential for outer membrane integrity and the positioning of a polar localization
factor: J Bacteriol, v. 192, p. 4847-58.
Yeiser, B., E. D. Pepper, M. F. Goodman, and S. E. Finkel, 2002, SOS-induced DNA polymerases
enhance long-term survival and evolutionary fitness: Proc Natl Acad Sci U S A, v. 99, p. 873741.
Yu, X. C., E. K. Weihe, and W. Margolin, 1998, Role of the C terminus of FtsK in Escherichia coli
chromosome segregation: J Bacteriol, v. 180, p. 6424-8.
50
Chapter 2
A DNA damage checkpoint in Caulobacter crescentus inhibits cell
division through a direct interaction with FtsW
This work was published as Modell, J. W., Hopkins, A. C., and Laub, M. T. 2011, A DNA
damage checkpoint in Caulobacter crescentus inhibits cell division through a direct interaction
with FtsW: Genes Dev, v. 25, p. 1328-43.
JWM, ACH and MTL conceived and designed the experiments. JWM and ACH performed the
experiments.
51
Abstract
Following DNA damage, cells typically delay cell cycle progression and inhibit cell division
until their chromosomes have been repaired. The bacterial checkpoint systems responsible for
these DNA damage responses are incompletely understood. Here, we show that Caulobacter
crescentus responds to DNA damage by coordinately inducing an SOS regulon and inhibiting the
master regulator CtrA. Included in the SOS regulon is sidA (SOS-induced inhibitor of cell
division A), a membrane protein of only 29 amino acids that helps to delay cell division
following DNA damage, but is dispensable in undamaged cells. SidA is sufficient, when
overproduced, to block cell division. However, unlike many other regulators of bacterial cell
division, SidA does not directly disrupt the assembly or stability of the cytokinetic ring protein
FtsZ, nor does it affect the recruitment of other components of the cell division machinery.
Instead, we provide evidence that SidA inhibits division by binding directly to FtsW to prevent
the final constriction of the cytokinetic ring.
52
Introduction
Cells frequently experience genotoxic stresses with potentially mutagenic or fatal
consequences. To maintain genome integrity, organisms respond by producing or activating
proteins with roles in translesion DNA synthesis, recombination, and repair. In addition,
organisms may delay the cell cycle and inhibit cell division until DNA repair and replication are
complete. In eukaryotes, these responses are orchestrated by elaborate checkpoint systems that
are well understood at the molecular level (Harper and Elledge, 2007). The bacterial mechanisms
of cell cycle and division control following DNA damage remain only partially understood, and
the extent to which different bacterial species use checkpoint systems is unresolved.
In bacteria, DNA damage triggers a change in gene expression called the SOS response
(Butala et al., 2009; Little and Mount, 1982). This gene expression program is initiated by the
autocatalytic cleavage of the transcriptional repressor LexA, an event stimulated by RecA, a
multifunctional recombinase and Rad51 homolog. Cleavage of LexA induces the SOS regulon,
which includes a battery of DNA repair genes, some of which are widely conserved in bacteria
(Erill et al., 2007). In addition, the Escherichiacoli SOS regulon includes sulA, which encodes a
cell division inhibitor (Huisman and D'Ari, 1981; Mukherjee et al., 1998). SulA directly inhibits
polymerization of FtsZ, a GTP-binding tubulin homolog that polymerizes at mid-cell and is
crucial to cytokinesis. As SulA is not conserved outside the y-proteobacteria, additional
mechanisms for inhibiting cell division likely exist in other bacteria.
Cytokinesis in bacteria requires the assembly of a large multicomponent complex of proteins
called the divisome. In E. coli, these components assemble in an ordered fashion. The first
protein to assemble is FtsZ, which polymerizes at mid-cell into a ring-like structure that is
tethered to the cytoplasmic surface of the inner membrane (Adams and Errington, 2009;
Lutkenhaus and Addinall, 1997). In addition to its direct role in constriction, the Z-ring also
nucleates assembly of the rest of the divisome, which ultimately coordinates constriction and
53
membrane invagination with construction of a septum, a process that requires localized
peptidoglycan synthesis (Errington et al., 2003). The initial formation of a Z-ring is mediated by
the proteins FtsA and ZipA, which stabilize FtsZ polymers and anchor the ring to the inner
membrane (Adams and Errington, 2009). In E. coli, additional components are then assembled in
a step-wise manner in the order FtsK, FtsQ, FtsL/FtsB, FtsW, FtsI, and FtsN (Goehring and
Beckwith, 2005; Vicente and Rico, 2006). The order of assembly is similar, but not identical,
in Caulobacter(Goley et al., 2011). Although the assembly of cell division proteins is relatively
well characterized, the execution and regulation of cell division remain poorly understood.
Previous studies of cell division regulation have focused almost exclusively on proteins that
modulate Z-ring assembly and stability. For instance, in E. coli, the regulators MinCD and SlmA
inhibit FtsZ polymerization at the poles and in proximity to chromosomal DNA, thereby
restricting Z-ring formation to mid-cell (Bernhardt and de Boer, 2005; Lutkenhaus, 2007);
MinCD and Noc play analogous roles in Bacillus subtilis (Wu and Errington, 2004). FtsZ is also
a focal point of regulation following cellular stress. In B. subtilis, UgtP coordinates Z-ring
assembly with glucose availability (Weart et al., 2007), while MciZ inhibits FtsZ polymerization
during sporulation (Handler et al., 2008). In E. coli, DNA damage induces SulA, which inhibits
FtsZ polymerization and GTP hydrolysis to prevent cell division (Justice et al., 2000; Mukherjee
et al., 1998).
It is unclear whether the numerous essential components of the cytokinetic ring other than
FtsZ can also serve as points of control. The SOS-induced inhibitors YneA, DivS, and Rv2719c
in B. subtilis, Corynebacteriumglutamicum, andMycobacteriumtuberculosis, respectively,
appear not to inhibit Z-ring formation (Chauhan et al., 2006; Kawai et al., 2003; Ogino et al.,
2008). However, the direct targets of these inhibitors remain unknown.
The a-proteobacterium Caulobactercrescentus is an excellent model system for
understanding the regulation of cell division. Caulobactercells are synchronizable, and the cell
54
cycle is accompanied by a series of easily tracked morphological transitions (Fig. 2.1A). Motile
swarmer cells reside in a GI phase and cannot initiate DNA replication. Swarmer cells then
differentiate into stalked cells and, concomitantly, initiate DNA replication. As replication
proceeds, cells grow and elongate, build a Z-ring at mid-cell, and eventually form a visible
constriction at this future site of cell division. Once S phase completes, the cell can divide
asymmetrically to produce a stalked cell that immediately reinitiates DNA replication and a
swarmer cell that must again differentiate before initiating.
How Caulobacterdelays cell division after DNA damage is unknown. As noted,
Caulobacterdoes not encode a homolog of SulA or other known SOS cell division inhibitors.
The disruption of DNA replication was shown to down-regulate the activity of CtrA, a cell cycleregulated transcription factor that normally promotes the expression offitsA andftsQ (Wortinger
et al., 2000). However, it is unknown whether the inhibition of CtrA is either necessary or
sufficient to delay cell division following DNA damage.
Here, we identify sidA (SOS-induced inhibitor of cell division A), which encodes a small
inner membrane protein that is strongly up-regulated following DNA damage. Overproducing
SidA is sufficient to inhibit cell division, suggesting it plays an analogous role to SulA in E. coli.
However, unlike SulA, SidA does not interfere with the assembly of the Z-ring or the
recruitment of other essential cell division components. Instead, SidA inhibits constriction by
interacting directly with FtsW and FtsN. Furthermore, mutations that suppress the lethality of
overproducing SidA map toftsWandftsI. We present evidence that FtsW, FtsI, and FtsN form a
subcomplex within the cytokinetic ring. Although these proteins have been implicated in septal
peptidoglycan synthesis, SidA does not appear to inhibit this activity. Our results instead suggest
a second role for these proteins in triggering the final constriction of the cytokinetic ring, an
activity that is inhibited by SidA. The FtsW/I/N subcomplex thus represents a key regulatory
55
node within the cell division machinery, and SidA is, to our knowledge, the first endogenous cell
division inhibitor with a known binding target other than FtsZ.
56
Results
DNA damage induces cell cycle arrest and a change in global transcription patterns
To investigate the mechanisms coupling DNA integrity and replication status with cell
division, we first analyzed the response of Caulobactercells to mitomycin C (MMC) and
ultraviolet light (UV), which damage DNA, and to hydroxyurea (HU), which disrupts DNA
replication by depleting nucleotide pools. Each treatment caused cells to transiently arrest the
cell cycle; cells continued to elongate but failed to divide (Fig. 2.1B). Using whole-genome
microarrays, we analyzed global changes in gene expression after each perturbation. Wild-type
cells were grown to mid-exponential phase in either rich medium or minimal medium and were
exposed to one of the three agents (MMC, UV, or HU). Samples were collected immediately
prior to treatment and every 20 min up to 80 min. RNA from each time point was compared with
RNA from the pretreatment sample on DNA microarrays.
A total of 160 genes, or nearly 5% of the annotated genes in Caulobacter,were significantly
induced or repressed during at least one of these six time courses (for complete data, see
Supplemental Table S1 in (Modell et al., 2011)). To identify candidate regulators of cell division,
we focused on the gene expression changes that were common across all conditions. We found
that 74 genes changed consistently across the six time courses, with 28 being down-regulated
and 46 being up-regulated (Fig. 2.1C; Fig. 2.2). Of the 28 genes consistently repressed, 10 were
identified previously as direct targets of the master cell cycle regulator CtrA, and nine others are
likely indirect targets, as they showed significant decreases in expression in ctrAtS strains (Laub
et al., 2002). A previous study demonstrated a similar down-regulation of two CtrA
targets,ftsQ andftsA, after HU treatment (Wortinger et al., 2000). We also found
thatftsQ mRNA levels dropped after DNA damage, but just missed the thresholds set for
inclusion, while the probe forftsA did not provide reliable data.
57
A
*
~f~d
m.
SW
S
WNto
predivsion.l
(0
oell diok
(0
inhibilion
DNA damage
B
HU
MMC
rk m*"rchraala
1
uv
'w'
e
rchnink
-4 -2 0 2 4 s
DI
20
0
I
15
AkiA
10
r*CA
5
20
o
4
i0
time (min.)
58
Figure 2.1. DNA damage induces global changes in gene expression and inhibits cell division. (A) Schematic of
the Caulobactercell cycle with and without DNA damage. (B) Wild-type CB 15N grown to mid-exponential phase
was exposed to UV light, HU, or MMC for 80 min or left untreated, and was imaged by differential interference
contrast microscopy. Cells exposed to MMC were also examined after 160 min. Bar, 2 im. (C) Gene expression
profiles of DNA damage-regulated genes in Caulobacter. Profiles are shown for 74 genes significantly changed in
expression level after treatment with HU, MMC, or UV light in rich (PYE) and minimal (M2G) media. The column
labeled "LexA" indicates with a red box whether a gene has a LexA box upstream. The columns labeled "CtrAdirect" and "CtrA-indirect" indicate with a green or black box, respectively, whether a gene is a direct target of CtrA
or is indirectly affected by CtrA. For annotation of individual genes and complete data, see Figure 2.2 and
Supplemental Table S1 in (Modell et al., 2011). Expression ratios are shown relative to untreated cells and are
represented using the color scale shown. Gray blocks indicate missing data. (D) Graph showing induction
of sidAafter DNA damage, relative to other members of the SOS regulon: recA,lexA, and ssb. Response curves are
the average across all six time courses in C.
59
HU
rich
minimal
MMC
rich
minimal
UV
rich
mdnimal
cc~mre
=MU OsY~xadaniIpnei
ftYPrin
CCM7ll conserved hyo"""cei prtein
S cos211 DNA poiysrase MI *1ph esunit
protein
ccee hypoles
c2r NpSt.canpo uromep.n..ow.gor
CC232 bsOen.WNm.. daed domias- conteinng
cc30uypoeice cyoneft proi
c2eSUA1Psdependunt hake..e
cC31s 1 hypsOftN cylosoi protein
CC3044 IApoINNce protein
cc1ss raeod 8AM sufeftny protein
cC371S hypowdcd protein
cMiS antidn protein pwD-2
cC30m8 MIa tomin protein
czui
otun proteinrsW54
cC3a hypoOencd protein
cc82 oldolaite redalasse fgayproten
cC211Stoinprotein hE-3
cC=U3wetN DNA ceMylaee uperanhy proein
cUS SXzchclm
ABC SURnS A
c=e3 hoINcd cytooeno protein
CC1498 dInree-emnd DNA Inakg protein
CD3=S#OT4WdVNAMnakgprotein
cysasefo protein
cS212 ypoenM
C0S21 muC lay proten
ccOs2 D
MA-MIN-- Oycoeylase
ccim2lexA repressor
cOS hypolwdo cgtow.. p erasn
cim DNA hoelesel
CO2NI conserved hW*temei protein
CC3113 L.Uas.4Sa tcSi
ccim fpoelene protein
CCS213 conserved hypothecal protein
CC2272 rndonucesse IN
CCIGU3 mathvoodNrot Ilyse
OCU3lS g
out nlo ons 4rnwfse
Ccsi hypomecei protein
CCMSl genece ethongeprotein,putgae
CC2SiI gInl kOne 8nseMw rae, Metaxin sufneny
ccauestWe I reotdcfo-minoalonsystem
cW33R7A&Cbpuormspensein pfutein
ccSSueco.disero.i
eUS(NPanq
csss uM
pemmse elme-32 factor
c 32 conevs
hypothete
protein
CCIW clmotuds mOO pretein
cC12 hehene phemphwotngermse doma protein
ccaSm ioproetn
cCOm inft n.aW rod M
Meadon protein
ccift hypolhwce poten
CC2S162 mohM-McepUng clumotuds protein
cesi vins protn
cC-11StGc AAA-aiFy reopan.
regUkdor
cc0S probaes cwmoreoeptor YVA
C3SWtonespone
treceptor
cCesz cihwatseme recoiverdommin protein CheYI
ccm SA .Whaoeott.i.cmsaproein
ceini ~me b e NdeOMphII receptor
COW conraerved hypodecew
WC"SMhffooledeprotak
ccmadoeuscei
pran, DUFU2S
protein
cooe protein
CCOSMfo" n#w erplot
la
CC2MerjPMceehvpofuftd pi oto
proten
C1123 conserved nyponle
ccOSS conserved hypoehescal pten
ccOsVwnqein
cO2S9tWeolVPMl proein p4A
cc29Stty1i. l"piVn protein pA
Figure 2.2. Annotated gene expression profiles. Transcriptional profiles for the 74 genes regulated during DNA
damage (see Figure 2.1) are shown with the corresponding CC numbers and NA 1000 annotation.
60
Identification of sidA, a novel SOS-induced cell division inhibitor
Inhibiting CtrA may help block cell division by decreasing the expression offtsQandftsA.
However, many bacteria induce genes-such as sulA in E. coli and yneAin B. subtilis-following
DNA damage that post-transcriptionally inhibit cell division (Huisman and D'Ari, 1981; Kawai
et al., 2003). As noted, the Caulobactergenomedoes not contain homologs of these genes. To
identify a damage-inducible cell division inhibitor, we examined our expression data for genes
that (1) are rapidly and strongly up-regulated following DNA damage, (2) are predicted members
of the SOS regulon based on the presence of a consensus LexA-binding site (da Rocha et al.,
2008), and (3) have no known or predicted role in DNA repair. Based on these criteria, we
identified CC 1927, which we named sidA, which has a predicted LexA-binding site in its
promoter region and is induced nearly 14-fold within 20 min of DNA damage (Fig.
2.2D). sidA is predicted to encode a highly hydrophobic 40-amino-acid protein lying 39 bases
upstream of the replicative DNA polymerase III a subunit (dnaE). Despite their close
proximity, dnaE and sidA are likely not cotranscribed, as the expression levels
of dnaE and sidA are not strongly correlated in the experiments here or reported previously
(Laub et al. 2000).
SidA is the principal SOS-induced cell division inhibitor
To explore the function of sidA, we created a strain in which all but the first and last three
amino acids of the originally annotated coding sequence were deleted. This AsidA strain showed
no obvious growth or morphological defects in standard rich (Fig. 2.3A-B) or minimal medium,
suggesting that sidA is dispensable in undamaged cells.
To assess the ability of AsidA cells to withstand DNA damage, we first tested the growth of
wild type and AsidA on plates containing increasing concentrations of the damaging agent MMC.
Under such conditions of DNA damage, AsidA cells showed no major viability defect (data not
shown). To better characterize the DNA damage response of AsidA, we used time-lapse
61
microscopy to monitor cells grown on agarose pads containing MMC (Fig. 2.3C). For both wildtype and AsidA, we tracked >250 individual cells from three independent movies, noting for each
cell its initial cell cycle stage and subsequent cell division behavior. Among cells that had yet to
initiate division when first placed on the pad, 24.9% of AsidA cells divided compared with 15.1%
of wild-type cells. We also observed a sixfold increase in the number of minicells
in AsidA populations, which resulted from ectopic cell divisions occurring near the cell poles
(Fig. 2.3C). SidA thus appears to help prevent inappropriate cell division events following DNA
damage at both mid-cell and the poles.
A
B
1.4
U.
wt
-sd- AsidA
m.*.
CdA
doubling time
77 mil.
78 min.
MM
93 min.
0.8
0
0.
00.6
0.2
02
0
2
4
6
8
10
12
14
time (h)
AsidA
min,. In MMC:
0
100
200
300
400
S00
AsidA
S00
miicll6.8%
Wt
1.1%
division
n=
24.9%
15.1%
253
297
Figure 2.3. sidA is the primary SOS-induced cell division inhibitor. (AB) Micrographs (A) and growth curves (B) of
wild-type, AlexA, AsidA, and AlexA AsidAcells grown in rich medium. Bar in the top left panel, 2 pim. (C)
AsidA (tetR) cells were placed on agarose pads containing MMC and imaged for up to 600 min. Examples of
minicell formation resulting from division near a cell pole or division at mid-cell are shown. The percentage of cells
that produce minicells or divide medially are shown on the right and are compared with wild-type cells treated
identically.
62
Although these data indicate that SidA inhibits cell division following DNA damage,
most zsidA cells still eventually formed long filaments (Supplemental Movie SI in (Modell et
al., 2011)), indicating that an alternative means of division inhibition also functions following
DNA damage. Note that sulA in E. coli behaves similarly; it is the primary SOS-induced
inhibitor of cell division, but not all cells lacking sulAdivide prematurely following DNA
damage, presumably due to the action of another cell division inhibitor (Gottesman et al., 1981;
Hill et al., 1997).
To determine whether SidA is the primary division inhibitor within the CaulobacterSOS
regulon, we created a strain harboring deletions in both lexA and sidA. The lexA deletion alone
results in extreme cellular filamentation and a reduction in growth rate owing to constitutive
expression of the SOS regulon, even in the absence of DNA damage (Fig. 2.3A,B; (da Rocha et
al., 2008)). If SidA is the primary SOS-induced inhibitor of cell division, a deletion
of sidA should eliminate or alleviate the phenotypes of a AlexA strain. Strikingly, we found that
deleting sidAin a AlexA strain completely suppressed cellular filamentation (Fig. 2.3A),
demonstrating that SidA is the principal SOS-regulated division inhibitor in Caulobacter.
Whereas AlexA cells were often 10-20 times the length of wild-type cells, AsidA AlexA cells
were nearly indistinguishable from wild type. Deleting sidA also partially suppressed the growth
phenotype of a AlexA strain (Fig. 2.3B). A complete rescue of growth is likely not possible,
owing to the constitutive activation of DNA repair and recombination genes in the AlexA strain.
Taken together, our data thus far suggest that sidA (1) is strongly induced following DNA
damage as part of the CaulobacterSOS regulon and (2) plays a role in preventing cell division
post-damage.
SidA is sufficient, when overproduced, to inhibit cell division
Next, we wanted to test whether SidA was sufficient to inhibit cell division in the absence of
DNA damage by overexpressing sidA in undamaged, mid-exponential-phase cells. We fused the
63
annotated sidA coding sequence to the xylose-inducible promoter Pxy on a high-copy plasmid
and transformed this construct into the wild-type strain CB 1 5N. To overproduce SidA, cells were
grown in medium containing xylose. Within 1.5 h, cells were elongated relative to wild type or
cells harboring an empty vector. After 3 and 6 h, nearly every cell overproducing SidA had
formed a long filament, often with multiple pinch sites that likely represent nascent cell division
sites (Fig. 2.4A,B). These data indicate that SidA is sufficient to inhibit cell division even in the
absence of DNA damage, and that sidA may participate in a DNA damage checkpoint similar
to sulA.
Homologs of sidA are often predicted to start at a position corresponding to the second
methionine of the originally annotated sidA. Additionally, we noted that, unlike the annotated
start site, the second methionine in sidA lies downstream from an apparent Shine-Delgarno
sequence, suggesting that it is the primary translational start site. To test this prediction, we
mutated the first methionine on the Pyl:sidA overexpression plasmid from ATG to AAG. For
cells harboring this construct, growth in xylose also led to cell division inhibition, suggesting that
the second start site is functional (Fig. 2.4B). The cellular filamentation phenotype was,
however, diminished when overexpressing the shorter version of sidA. The more pronounced
phenotype seen when overproducing the longer version may indicate that the additional 11
amino acids at the N terminus either stabilize the protein or somehow render SidA more potent.
We favor the former, as overproducing SidA with an N-terminal M2-epitope tag, which is
unlikely to enhance potency, led to severe cellular filamentation, as with the original
Psyl:sidA construct (Fig. 2.4B). We could not, however, directly test the stability and size of the
native SidA protein, as it is difficult to raise antibodies against such short, hydrophobic proteins.
To more firmly identify the translational start site, we fused the coding sequence for green
fluorescent protein, gfp, to the sidA promoter at each of the two putative translational start sites
on a low-copy plasmid. Following MMC treatment, only cells bearing the fusion of GFP to the
64
second methionine exhibited significant fluorescence (Fig. 2.4C). We conclude that SidA
translation initiates primarily, and perhaps exclusively, from the second site to produce a 29amino-acid protein; hereafter, references to SidA imply this shorter version and SidA* refers to
the longer form.
C
A
- MMC
+ MMC
1.2
U2
1.0
0.
0.8
sidA*(T2A)
0
0.6
0.4
M2-sidA
sidA*
0.2
00
2
4
6
U2
0.
10
i
0..
time in xylose (h)
B
0.
eid#A*
aijEA*irlA I
AJ9_&iAA
glu
xyl
(3 h)
xyl
(6 h)
Figure 2.4. Overproducing SidA is sufficient to inhibit cell division. (A) Growth curve in rich medium of cells
carrying a high-copy plasmid on which the xylose-inducible promoter drives expression of the M2 epitope
only; sidA *, the originally annotated CC 1927 ORF; sidA *(T2A), which harbors a mutation in the first annotated start
codon; or M2-sidA, in which the M2 epitope is fused to the second start site in the originally annotated CC 1927
ORF. (B) Each strain from A was grown to mid-exponential phase in rich medium supplemented with glucose to
repress expression of the plasmid-encoded construct. Expression was then induced by adding xylose, and cells were
imaged by differential interference contrast (DIC) microscopy after 3 and 6 h. (C) The promoter and leader region of
CC1927, up through either the first or second annotated methionine, were fused to the coding region of EGFP to
generate PsdA*.egfp and Psja:egfp,respectively. Strains carrying each construct on a low-copy plasmid were grown
in the presence or absence of MMC, and expression was examined by epifluorescence microscopy. Bars, 2 tm.
65
SidA does not disrupt divisome assembly or localization
How does SidA inhibit cell division? We first tested whether overproducing SidA affected
patterns of gene expression. We compared RNA from cells overproducing M2-SidA for 45 or 90
min with cells grown in noninducing conditions on whole-genome DNA microarrays, but did not
observe any significant changes to the SOS or CtrA regulons (data not shown). Overproducing
SidA also did not affect chromosome content, as measured by flow cytometry, indicating that
SidA does not inactivate CtrA. Instead, SidA likely acts post-transcriptionally to inhibit cell
division.
In E. coli, the SOS-induced regulator SulA disrupts polymerization of the cytokinetic ring
protein FtsZ (Mukherjee et al., 1998). To determine whether SidA functions similarly, we
introduced the Pxy:M2-sidA overexpression plasmid into a strain in whichftsZ-eyfp is expressed
from the chromosomal vanillate promoter P,,n. Vanillate was added to cells 1.5 h prior to the
addition of xylose to allow FtsZ-EYFP to accumulate to sufficient levels for visualization by
epifluorescence microscopy. After growth in xylose for 1.5 h, the majority of cells had begun to
elongate, but FtsZ foci were still visible at the constriction site (Fig. 2.5A), indicating that SidA
did not disrupt the cytokinetic Z-ring. After 3 and 4.5 h in xylose, cellular filamentation became
more severe, but FtsZ foci were still present (Fig. 2.5A). In some cells, the FtsZ ring appeared to
move a short distance from the original constriction site (Fig. 2.5A), indicating that FtsZ rings
are also able to assemble de novo even in the presence of high levels of SidA. To confirm these
results, we used time-lapse microscopy of this same strain grown on an agarose pad containing
xylose (Supplemental Movie S2 in (Modell et al., 2011)). This analysis confirmed that FtsZ rings
typically remain at constriction sites in individual cells well beyond the time normally necessary
for septation, although occasionally the Z-ring relocated to a new position along the long axis of
the cell body. Collectively, these results demonstrate that SidA does not disrupt the
polymerization, assembly, or stability of FtsZ rings.
66
A
P,,,:M2-sidA
xyl (hours post-shift)
1.5
0
I.
91% (n=123)
U.
0
95% (n=155)
B
gfp-ftsi + Py:M2-sidA
xyl (hours post-shift)
glucose
0.5
1.0
1.5
3.0
4-5
CL
0.
0
79% (n=172)
C
gfp-ftsl+ Pr,,:M2-sidA
glu (minutes post-shift)
xylose,
10
20
30
40
50
60
70
CL
U
(D
67
Figure 2.5. SidA does not prevent assembly of the cell division machinery. (A) Subcellular localization of FtsZ was
examined in a strain expressingftsZ-eyfp from the vanillate-inducible promoter P,,, and overexpressing M2sidA from a xylose-inducible promoter on a high-copy plasmid. Cells were grown to mid-exponential phase in rich
medium with glucose and then shifted to xylose. Vanillate was added 1.5 h before shifting to xylose. At the times
indicated, samples were taken and cells were imaged by DIC and epifluorescence microscopy (top two rows). The
white asterisk indicates a FtsZ ring that is no longer associated with a constriction site and so likely moved and
reassembled. (Bottom two rows) Localization of FtsN was examined in a strain expressing an egfp-ftsN fusion at its
native chromosomal locus and overexpressing M2-sidA from a high-copy plasmid. The percentage of cells with foci
after 4.5 h in xylose is shown below the last panel. (B) Localization of FtsI was examined in a strain expressing gfpftsl at its native chromosomal locus and overexpressing M2-sidA from a xylose-inducible promoter carried on a
high-copy plasmid. Cells were grown to mid-exponential phase in rich medium with glucose and then placed on
agarose pads containing xylose to produce M2-SidA. Individual cells were followed by time-lapse microscopy, with
phase and epifluorescence images captured at the time points indicated. (C) Cells expressing gfp-ftsI and xyloseinducible M2-sidA were grown in the presence of xylose for 4.5 h to inhibit cell division and induce cellular
filamentation. Cells were then placed on an agarose pad containing glucose to repress M2-sidAexpression.
Individual cells with localized GFP-FtsI were then followed by time-lapse microscopy, with phase and
epifluorescence images captured at the time points indicated. Bars, 2 m.
SidA thus acts downstream from FtsZ assembly to affect cell division. In E. coli
and Caulobacter,the cell division machinery assembles in a stepwise manner, with the
recruitment of each component to mid-cell requiring recruitment and assembly of prior
components (Goehring and Beckwith, 2005; Goley et al., 2011; Vicente and Rico, 2006). Once
the divisome is fully assembled, FtsZ constriction and cell envelope septation proceed, although
the signals and mechanisms driving these processes are poorly understood. As FtsZ recruitment
is one of the earliest steps of cell division, SidA could block the recruitment of a late-arriving
cell division protein. Alternatively, SidA may not affect divisome assembly and may instead
inhibit the constriction process.
To distinguish between these two possibilities, we examined the localization of the cell
division protein FtsN, which is one of the late-arriving essential components recruited to the
division site (Costa et al., 2008; Goley et al., 2011; M611 and Thanbichler, 2009). The Pxy,:M2sidA overexpression plasmid was introduced into a strain expressing egfp-ftsN from its native,
chromosomal locus. When grown in the presence of glucose, EGFP-FtsN foci were seen at midcell only in very late, deeply pinched predivisional cells. When grown in xylose to induce SidA
production, these FtsN foci remained intact, always localizing at or near the constriction site
(Fig. 2.5A). Similar results were obtained with Venus-FtsW (see below; Fig. 2.9B). The
68
persistence of FtsZ, FtsW, and FtsN foci suggests that SidA does not disrupt the assembly of the
cell division apparatus; instead, SidA likely inhibits the active process of constriction.
Despite the persistence of FtsN foci, SidA could somehow prevent the localization of
another divisome component while still allowing FtsN assembly. We therefore tested whether
SidA overproduction affected the localization of Ftsl, another essential, late-arriving cell division
protein that participates in septal peptidoglycan synthesis. For these experiments, the Py,:M2sidA overexpression plasmid was introduced into a strain expressing gfp-ftsI from its
chromosomal locus. Cells were grown initially in glucose and then shifted to xylose to induce
sidA and were followed by time-lapse microscopy. As expected, cells became filamentous owing
to the accumulation of SidA. GFP-FtsI foci were seen localized to mid-cell and were maintained
through the duration of the 4.5 h cells were imaged (Fig. 2.5B).
Taken together, our data suggest that, in cells overproducing SidA, the cell division
machinery is fully assembled but somehow inactivated. To test whether cell division can occur
once SidA is removed, we examined the behavior of cells harboring the Psy,:M2-sidA plasmid
and expressing gfp-ftsl. Cells were grown initially in xylose for 4.5 h to inhibit cell division and
were then returned to medium containing glucose to shut off sidA expression. Within 30 min,
many cells were noticeably more pinched at the same sites initially containing GFP-FtsI foci,
and, after an hour, these cells had divided (Fig. 2.5C). Importantly, at time points during and
immediately preceding division, the GFP-FtsI foci did not appear significantly brighter than the
first time point, regardless of initial intensity. These analyses suggest that SidA does not drive
delocalization of FtsI and that cells are poised, once SidA is cleared, to complete cell division.
Mutations inftsW and ftsI suppress the effects of overproducing SidA
To identify the protein(s) targeted by SidA, we screened for mutations that suppress the
growth defect of sidA overexpression by isolating mutants that form colonies despite SidA
overproduction, which is normally lethal. Wild-type cells were transformed with a high-copy
69
plasmid harboring Py,:M2-sidA and grown on plates containing xylose. This process yielded 80
colonies; an identical transformation with an empty vector yielded tens of thousands of colonies,
indicating that suppressors of SidA overproduction are rare. To ensure that suppression did not
result from mutations on the sidA overexpression plasmid, we tested whether plasmids from each
colony could, in a clean background, still disrupt cell division. We also confirmed by Western
blotting that putative suppressor strains still produced high levels of M2-SidA in xylose. In total,
we identified nine strains that supported growth despite the production of high levels of M2SidA. Strikingly, six of the nine suppressors mapped toftsI orfts W, both of which encode lateacting, transmembrane components of the cell division machinery. FtsI participates directly in
the synthesis of septal peptidoglycan as a periplasmic transpeptidase, while FtsW is a polytopic
membrane protein that likely translocates peptidoglycan precursors from the cytoplasm to the
periplasm (Errington et al., 2003; Hltje, 1998; Mohammadi et al., 2011). The amino acid
substitutions in FtsW that suppress sidA overexpression-A3IK, F145L, and T180A (found
twice)-are predicted to lie adjacent to the cytoplasmic end of transmembrane helix 1, the
cytoplasmic end of transmembrane helix 4, and the periplasmic end of transmembrane helix 6,
respectively (Fig. 2.6A). The substitutions identified in FtsI-145V and F58V-are located in the
cytoplasmic tail and the center of FtsI's single-transmembrane helix, respectively (Fig. 2.6A).
For the remaining three suppressor strains, no mutations were found in any known
divisome/fts gene.
Eachfts W andftsl suppressor mutation was introduced into a clean wild-type genetic
background by allelic replacement, and no growth or morphological phenotypes were seen
during growth in rich medium (Fig. 2.7). We then transformed each strain with the M2sidA overexpression plasmid and inoculated serial dilutions on rich-medium plates containing
xylose. Each suppressor mutation conferred a significant growth advantage on xylose as
compared with wild-type cells transformed with the same plasmid (Fig. 2.6B), and each mutation
significantly reduced the cellular filamentation that results from overexpressing sidA (Fig. 2.6C).
70
B
A
serial dilutions
Ftsl
FtsW
Wt
!sW(A31K)
transpeptid ase
T180A
dimerization
sW(F145L)
U)
A 31K
4
U
145V
F145L
C
periplasm
F58V
n
-
9
~ftsW(TI80A)
9.
ftsl(145V)
cytoplasm
SidA
fts(F58V)
Wt
ftsW(A31K)
ftsW(F145L)
ftsW(TI80A)
ftsl(14V)
ftsi(F58V)
wt
ftsW(A31K)
ftsW(F145L)
fsW(TI8OA)
1tsl(145V)
ftsl(F58V)
126'
98'
90'
97'
92'
98'
ki
C.
D
q)
doubling
time (min.)
E
sidA
F
TM
Pxy,:M2-sidA
C. crescentus .I
S-AL
Caulobacter K31
I
S
S
P. zucineum
kLA
8V
Brevundimonas sp. p LS
0. alexanddi RNLIRDTSAAVI
. maris RVVQDV
T
Nitrobacter sp. A P. bermudensis SPTI
RDL
C. crescentus
Caulobacter K31
P. zucineum
Brevundimonas sp.
0.alexandrii
M.mars
Nitrobactersp.
bermudensis
P.
ftsI
*L
GF
CT
CS
LVA.
$671--QNG---
C
IT
ISVTAAAVALSLSAALAG- IAMWGSVLGAAAV gQLIPQLS
iAVLGVI INFVA--
S
M
i"iwl*
CckA-GFP --
xyl
S
M
O0
QVS
GIG
a
T
TR
LIIJ SC
A
LRVNGLSLAL
-VL
RVEL
LSVIGLVLS
-IS
RAVQI
VGPfII
SLI AVIAARLVMFU
LILCAVAPGLOGLLLRLGNVS
G
FNAIG
I
L
glu
CtrA -*
W
M2-SidA --
TM
Figure 2.6. Mutations infis W andftsI suppress the SidA overproduction phenotype. (A) Summary of a sidA
overexpression suppressor screen. The location of mutations identified in FtsW and Ftsl are indicated on schematics
representing the domain structure of each protein. (B) Each suppressor mutation was introduced into a clean wildtype background by allelic replacement, followed by transformation with the M2-sidA overexpression plasmid. Each
strain was then grown to mid-exponential phase, and serial dilutions were plated on PYE supplemented with
chloramphenicol to maintain the plasmid and with xylose to induce SidA. (C) Cellular morphology of strains
71
harboring each suppressor mutation and overexpressing M2-sidA for 6 h. Bars, 2 pm. (D) Cellular morphology of
strains harboring each suppressor mutation and a deletion of lexA. The doubling time of each strain in rich medium
is indicated below the corresponding micrograph. (E) The location of a predicted transmembrane domain
within sidA is shown schematically at the top and directly below in an alignment of the coding region of SidA
orthologs. The location of a single predicted transmembrane domain withinftsI is shown schematically at
the bottom and directly above in an alignment of the transmembrane domains of FtsI orthologs. The alignments are
aligned to each other, using the last predicted amino acid as an anchor point. The arrow indicates a conserved
phenylalanine in both alignments. Black and gray shading indicate residue conservation and similarity, respectively,
found at that position in >50% of aligned sequences. (F) Subcellular fractionation of cells overexpressing M2sidA from a xylose-inducible promoter on a high-copy plasmid for 1.5 h and expressing cckA-gfp from the
chromosome. Samples from cells grown in either glucose or xylose, as indicated, were fractionated into soluble (S)
and membrane (M) fractions. Samples were separated by SDS-PAGE, transferred to a PVDF membrane, cut into
three pieces, and probed with antibodies specific for GFP, CtrA, or the M2 epitope.
Each mutation also significantly reduced the cellular filamentation of a AlexA strain (Fig. 2.6D),
which, as noted (see Fig. 2.3), is strictly dependent on sidA. The FtsW(F145L) and FtsI(145V)
mutations led to the most complete suppression of both growth and morphological phenotypes
(Fig. 2.6C,D). These results suggest that SidA inhibits cell division at a late stage by modulating
an activity of FtsW or FtsI.
wt
ftsW(A31K)
ftsitF145L)
ftsW(T18OA)
fti3(145V)
ft(F5#V)
Figure 2.7. ftsW and ftsl suppressor mutations display wild-type morphology during growth in rich medium. Each
suppressor mutation was introduced into a clean wild-type background by allelic replacement. Cells were grown in
log phase and imaged by DIC microscopy.
SidA resides in the inner membrane and interacts directly with FtsW and FtsN
As the suppressors of sidA overexpression mapped near or within the transmembrane
domains of FtsW and FtsI, we considered whether SidA is also localized to the inner membrane.
An analysis using TMHMM (Krogh et al., 2001) predicted that residues 9-29 of SidA constitute
a transmembrane domain. In the middle of this domain is a highly conserved phenyalanine; a
conserved phenylalanine is also found in the middle of the transmembrane domain of FtsI, which
was also the site of the mutation F58V in FtsI that suppressed sidAoverexpression (Fig. 2.6E).
72
To determine the subcellular localization of SidA, we fractionated cells overproducing M2SidA and probed both membrane and soluble fractions with an a-M2 antibody. SidA was found
exclusively within the membrane fraction after 1.5 h of overproduction (Fig. 2.6F). As controls,
we confirmed that the cytoplasmic response regulator CtrA was found predominantly in the
soluble fraction, while the sensor histidine kinase CckA was found exclusively in the membrane
fraction.
To determine whether SidA interacts physically with FtsW, FtsI, or any other divisome
component, we used a bacterial two-hybrid system based on the reconstitution of a split
adenylate cyclase (Karimova et al., 2005). We found that, of the essential divisome components
FtsZ/A/K/Q/L/B/W/I/N, SidA interacted most strongly with FtsW and to a lesser extent with
FtsN (Fig. 2.8A). Interestingly, FtsW and FtsN also interacted strongly with each other, as
reported recently in E. coli (Alexeeva et al., 2010). Although SidA did not interact with fulllength FtsI, neither did FtsW (data not shown), as expected based on studies in E. coli (Mercer
and Weiss 2002). We therefore tested a truncated version of FtsI (FtsIAC) lacking its periplasmic
catalytic domain. This construct did interact weakly with FtsW and FtsN, but did not interact
with SidA. However, an interaction between FtsIAC and SidA was observed when full-length,
untaggedfts W was cotranscribed withftsI C, suggesting that FtsIAC and SidA can bind FtsW
simultaneously and likely do not compete for the same site. The strength of the SidA-FtsW and
FtsW-FtsN interactions were approximately fivefold lower than a soluble zip-zip
homodimerization-positive control. However, divisome components must interact in the
membrane, where protein levels and molecular orientations may be more limited than with the
cytoplasmic zip-zip interactions. Collectively, these interaction studies demonstrate that (1)
FtsW binds to FtsI and FtsN, with these three proteins likely forming a subcomplex within the
cytokinetic ring, and (2) SidA directly binds FtsW and FtsN and is brought in close proximity to
FtsI via FtsW. We also found that, in cells synthesizing low levels of M2-EGFP-SidA,
fluorescent foci were frequently seen at pinch sites within cells (Fig. 2.9A). These foci were
73
often relatively dim, similar to the foci of Venus-FtsW observed in cells synthesizing M2-SidA
(Fig. 2.9B) and in contrast to FtsZ-YFP foci (Fig. 2.5A). These data are consistent with a model
in which SidA binds FtsW, a low-abundance divisome component.
A
14 -1
F
0
11
12
10
E
0
4,
171311. L
0
T25: FtsZ
FtsA
4~ ~L~rhILiA
FIK FtsQ
FtsL
FtsB
Ft*W
77
FtWAC FtsN FtsIAC FtSW
FtsN
FtSW
(+FtsW)
TI:
B
M2-SidA
C
120
FtsIAC FtSIAC FtsN
160
100
120
so
100
10
60
40
40
20
20
0
0
T25:
T18:
Ft*W
FtsW FtsW FMaW
(A31K) (F145L)(TI8OA)
M2-SidA
T25: FtsW
T18:
FtsW RaW Ft&W
(A31K) (F145L)(T18OA)
M2-SidA
Figure 2.8. SidA interacts with the late-arriving divisome components FtsW and FtsN. (A) Bacterial two-hybrid
analysis of interactions between M2-SidA and cell division proteins fused to T18 and T25, as indicated. Interactions
were quantified using a Miller assay and are reported relative to empty vector controls, which yielded 60 Miller
units. Each interaction was measured in triplicate; error bars represent the standard error of the mean.
FtsA# indicates the FtsA-MalF(TM) fusion described in the Materials and Methods. FtsIAC was tested alone and
while producing untagged FtsW, as indicated. Asterisks indicate a statistically significant difference (P < 0.05, onesided t-test) relative to empty vector controls. (B,C) Bacterial two-hybrid analysis of interactions between M2-SidA
(B) or FtsN (C) fused to T18 and mutants of FtsW fused to T25. Interactions are reported as a percentage of that
measured for wild-type FtsW with M2-SidA (B) or with wild-type FtsN (C).
74
A
P4,:M2-egfp-sidA
0.
U-
B
Pv,,:Venus-ftsW; PxM2-sidA
(+ vanlglu, 4.5 h)
(+ vanlxyl, 4.5 h)
0.
UL
a.
Figure 2.9. SidA and FtsW localize to constriction sites. (A) Subcellular localization of SidA was examined in a
strain expressing M2-egfp-sidA from a xylose-inducible promoter on a high-copy plasmid. Cells were grown to midexponential phase in rich media supplemented with glucose and imaged by phase and epi-fluorescence microscopy.
(B) Localization of FtsW was examined in a strain expressing yfp-fts W from the vanillate promoter P,,an and
overexpressing M2-sidA from a xylose-inducible promoter carried on a high-copy plasmid. Cells were grown to
mid-exponential phase in rich media with glucose, shifted to xylose and vanillate for 4.5 hours and imaged by phase
and epifluorescence microscopy.
We also assessed the ability of the FtsW suppressor mutants to interact with SidA. The
suppressor mutations F145L and T180A in FtsW both resulted in a weaker interaction with SidA
(Fig. 2.8B), but did not disrupt interaction with FtsN (Fig. 2.8C). The FtsW mutant A3 1K did not
interact with SidA or FtsN, suggesting that it may be misfolded or mislocalized in E. coli.
Notably, the substitution F145L significantly reduced the binding of FtsW to SidA, and this
75
substitution yielded the most complete suppression of sidA overexpression (Fig. 2.6C,D). We
conclude that SidA likely inhibits cell division through its direct interaction with FtsW, and that
mutations in FtsW suppress this inhibition by decreasing its affinity for SidA.
SidA prevents constriction of the Z-ring without disrupting synthesis of nascent septal
peptidoglycan
Collectively, our data indicate that SidA targets an activity, but not the localization, of the
late-stage cell division proteins FtsW, FtsI, and FtsN. Each of these proteins is thought to
participate in the synthesis of septal peptidoglycan, a critical aspect of cell division. FtsW likely
translocates peptidoglycan precursors into the periplasm, where they are incorporated into the
nascent septum by transglycosylases and the transpeptidase FtsI (Mohammadi et al., 2011). The
role of FtsN is less clear, although it has been proposed to stabilize the divisome and recruit
several nonessential proteins involved in cell wall remodeling (Gerding et al., 2009; Rico et al.,
2010). Given its affinity for FtsW, SidA could inhibit cell division by preventing the
translocation of septal peptidoglycan precursors into the periplasm. To test this possibility, we
examined cells stained with fluorescein-tagged vancomycin (Van-FL), which is thought to label
sites of nascent peptidoglycan synthesis by binding peptidoglycan precursor monomers and
growing chains in the periplasm (see the Materials and Methods; Fig. 2.11; (Daniel and
Errington, 2003; White et al., 2010)). In wild-type cells, Van-FL weakly stained the cell
periphery, with strong foci visible at deep constriction sites in late predivisional cells and weaker
transverse bands at mid-cell in early predivisional cells (Fig. 2.1 GA). No transverse bands or
bright foci of Van-FL were seen in cells (n = 57) depleted offtsZ and hence lacking an
assembled cytokinesis complex (Fig. 2. 1A). These results are consistent with Van-FL
recognizing nascent septal peptidoglycan in C. crescentus.
76
A
AftsZ, P ,,,:ftsz
+ mul 1I& h
Aft'W, Pvan:ftsW(- van, 7.5 h)
4. -h. I A h
U
P,,,:M2-sidA
B
+ xyl
+ gIu
70
90
75
95
min. post-synchrony
4
(U
4
P~y,:M2-sidA (+ xyl, 4.5 h)
Figure 2.10. Overproducing SidA does not inhibit the translocation of septal peptidoglycan precursors. (A) Van-FL
staining of wild-type, ftsZ depletion, andfis W depletion. Wild-type cells were grown to mid-exponential phase in
rich medium. TheftsZ depletion strain was grown to mid-exponential phase in the presence of xylose, washed, and
then grown in the presence of xylose or glucose for 1.5 h before imaging. Thefts W depletion strain was grown to
mid-exponential phase in the presence of vanillate, washed, and then grown without vanillate for 7.5 h before
imaging. (B) The M2-SidA overproduction strain was synchronized, released into rich medium containing either
glucose or xylose, and imaged at the times indicated. (C) A mixed population of the M2-SidA overproduction strain
was imaged after growth in xylose for 4.5 h. In all panels, cells were stained with Van-FL and were imaged by DIC
or phase and epifluorescence microscopy. Arrowheads indicate Van-FL staining in transverse bands and foci that are
at least 1.5-fold over the cell background (see the Materials and Methods; Fig. 2.11). Bars, 2 tm.
77
3803q*~u
S30M
20
40
0
s6
Distance (pixels)
Figure 2.11. Quantification of Van-FL foci. An example of the focus scoring metric is shown for a single cell. A line
was drawn through the center of the cell and its intensity values plotted in ImageJ. Values corresponding to the
focus maximum, cell background, and black background were scored (top, middle, bottom red lines respectively).
Focus intensity fold change above cell background (F) was calculated as the ratio of focus maximum to cell
background after the black background was subtracted from each. For the cell shown, the focus maximum, cell
background, and black background values were 3863, 2089, and 1202, respectively, resulting in F = 3-fold.
To test whether mid-cell Van-FL foci are dependent on the presence of FtsW, we examined
afts W depletion strain in which the only copy offts W is controlled by the vanillate-inducible
promoter Pvan on the chromosome. Cells grown initially in the presence of vanillate were washed
and then grown in the absence of vanillate for 7.5 h to deplete FtsW. As expected, these cells
showed a marked cell division defect, forming long filaments by 7.5 h that were comparable in
length with cells overproducing SidA for 4.5 h (Fig. 2.1 OA). However, unlike cells
overproducing SidA, cells depleted of FtsW were noticeably less pinched and formed long,
smooth filaments. Accordingly, Van-FL foci were observed only in 10% of cells (n = 70) after
7.5 h without vanillate (Fig. 2. 1OA, arrowheads); these rare foci may arise from incomplete FtsW
depletion or compensation by another translocase. We conclude that FtsW is required for cell
pinching, cell division, and mid-cell staining with Van-FL, consistent with its proposed role in
translocating peptidoglycan precursors to the periplasm.
To examine the effects of SidA on Van-FL staining, we synchronized cells containing
the M2-sidA overexpression plasmid and released swarmer cells into either glucose or xylose. In
glucose, transverse bands and foci of Van-FL were seen in predivisional cells; after 90 min, these
cells had divided and the daughter cells did not have foci, as expected (Fig. 2.1 OB). In xylose,
bands and foci of Van-FL were also seen in predivisional cells despite the overproduction of
78
SidA (Fig. 2.1OB). After 95 min, cells grown in xylose had not divided and most cells still
contained a focus of Van-FL at mid-cell. We also examined a mixed population of cells
overproducing SidA for 4.5 h and found that Van-FL foci were seen at pinched sites in 70% (n
83) of these filamentous cells (Fig. 2.1OC). Although we cannot rule out that SidA has a mild
effect on the flippase activity of FtsW, these data suggest that SidA likely does not inhibit cell
division by abolishing the synthesis of septal peptidoglycan. Instead, our results collectively
suggest that the FtsW/I/N subcomplex is critical in triggering the final stages of cytokinesis, and
that SidA blocks this step of cell division through a direct interaction with FtsW.
79
Discussion
Following DNA damage, cells typically must inhibit cell division to allow time for DNA
repair. In bacteria, the paradigm for damage-induced inhibition of cytokinesis has been the
depolymerization of FtsZ by SulA (Justice et al., 2000; Mukherjee et al., 1998).
However, sulA homologs are found only in y-proteobacteria, and it has been unclear whether
other damage-induced inhibitors also target FtsZ. Here, we identified sidA and showed that it
encodes the primary SOS-induced cell division inhibitor in C. crescentus. Unlike SulA in E. coli,
as well as many other known regulators of cell division, SidA does not directly disrupt FtsZ
polymerization or stability. Cells overproducing SidA retain a clear FtsZ ring at mid-cell, but
cannot constrict (Fig. 2.5). Overproducing SidA also did not affect the subcellular localization of
FtsW, FtsI, or FtsN, suggesting that SidA does not interfere with divisome assembly. Instead, our
results indicate that SidA inhibits constriction by targeting an activity of the late-acting cell
division proteins FtsW, FtsI, and FtsN (Fig. 2.12). Mutations in both fts W andftsI were identified
in an unbiased screen for suppressors of SidA overproduction. In addition, SidA was found to
bind directly to FtsW and FtsN, but, importantly, not to the FtsW suppressor mutants.
What are the activities of FtsW, FtsI, and FtsN in bacterial cytokinesis and how does SidA
affect them? These three essential cell division proteins are among the last components recruited
to the cytokinetic ring, where they likely form a subcomplex. Furthermore, each component has
been implicated in septal peptidoglycan synthesis. FtsW is a polytopic membrane protein and has
been proposed to translocate or "flip" peptidoglycan precursors from the cytoplasm to the
periplasm, where they are incorporated into existing peptidoglycan strands (Mohammadi et al.
2011). FtsI, also known as penicillin-binding protein 3 (PBP3), harbors transpeptidase activity in
its periplasmic domain and cross-links septal peptidoglycan (Spratt, 1977). The precise role of
FtsN in cytokinesis is unknown, but its essential function resides within its periplasmic linker
domain (M611 and Thanbichler, 2009). Our results suggest that SidA does not inhibit the putative
flippase activity of FtsW. In cells overexpressing sidA, the presence of Van-FL staining at
80
constriction sites indicates that septal peptidoglycan precursors are present in the periplasm.
SidA could target septal peptidoglycan modification activities of FtsW, FtsI, or FtsN that occur
in the periplasm after precursor flipping. However, this possibility seems unlikely, given that
SidA comprises only 29 amino acids and so lies almost entirely within the inner membrane.
Moreover, the mutations infts W andftsl that suppress sidA overexpression are all found within
the transmembrane or cytoplasmic domains.
- DNA damage
Y
= FtsZ ring
+ DNA damage
Y
FtsI
FRON
SidA
FtZW
LexA
NA damage
LeXA
sidA
L-- r*
,
SidA
:LI
Figure 2.12. A DNA damage checkpoint regulating cell division in C. crescentus. Model for regulation of cell
division by SidA following DNA damage. The schematic at the top shows the Caulobacter cell cycle and indicates
the progression of cell division, beginning with assembly and initial stabilization of FtsZ rings in stalked cells,
followed by constriction in late predivisional cells, and resulting finally in cell division. When DNA damage occurs,
the FtsZ ring is still assembled, but cell division is inhibited while cells continue to elongate. SidA inhibits cell
division by inserting into the membrane and binding FtsW and FtsN. The expression of sidA is under SOS control,
and hence is induced following DNA damage and cleavage of the LexA repressor.
81
Our results suggest that, in addition to controlling septum synthesis, FtsW, FtsI, and FtsN
may also play central roles in triggering the final constriction of the cytokinetic ring, and that
SidA may disrupt this function to inhibit cell division. As noted, we identified mutations
infts W andftsI that suppress the effects of overproducing SidA. The substitutions in FtsW likely
suppress by preventing the binding of SidA to the cytokinetic ring. However, a similar
explanation does not hold for the substitutions in FtsI, as SidA did not bind FtsI in our twohybrid system. Instead, these mutations may allow FtsL to help trigger constriction even when
SidA is bound to FtsW, thereby bypassing cell division inhibition. FtsI has also been suggested
to serve as a checkpoint signal that triggers Z-ring constriction in E. coli, possibly via
interactions with FtsA (Corbin et al., 2004), and FtsW and FtsI form a ternary complex with FtsZ
in M tuberculosis (Datta et al., 2006). Although we did not observe an interaction
between CaulobacterFtsI and FtsA or FtsZ, FtsI may act through an as-yet-unidentified
component of the cytokinetic ring. The identification of sidA overexpression suppressor
mutations that do not map to thefts genes should prove illuminating in this regard. Finally, we
note that FtsW, FtsI, and FtsN may not trigger constriction, and SidA could simply be using
FtsW and FtsN as a docking site from which it directly inhibits the activity of another divisome
component, such as FtsZ. Although we did not observe strong interactions between SidA and any
of the other essential Fts cell division proteins, this possibility cannot be ruled out completely.
The identification of SidA underscores how little is known about the execution of
cytokinesis in bacteria. The hierarchical dependencies underlying assembly of the cell division
apparatus have been well documented in recent years, largely through the study of fluorescently
tagged proteins. While a map of localization dependencies is useful, it remains a significant
challenge to define the biochemical activities of each cell division protein and to understand their
interdependencies and their roles in regulating cell division.
82
Is SulA an outlier?
SulA and the depolymerization of FtsZ has long been the paradigm for regulated cell
division inhibition in bacteria. Consequently, the characterization of other SOS-induced cell
division inhibitors has focused on Z-ring assembly and stability. However, in many cases, such
as YneA in B. subtilis (Kawai et al., 2003), DivS in C. glutamicum (Ogino et al., 2008), and
Rv2719c in M tuberculosis (Chauhan et al., 2006), a direct interaction with FtsZ has not been
documented. Moreover, in none of these cases did inhibitor overproduction result in complete Zring dissociation, as with SulA. In light of our results, we speculate that these inhibitors could
target cell division steps downstream from Z-ring assembly or stability, as has been proposed
recently for YneA and Rv2719 (Chauhan et al., 2006; Kawai and Ogasawara, 2006; Mo and
Burkholder, 2010). Consistent with this possibility, YneA, DivS, and Rv2719c are each singlepass transmembrane proteins like SidA but in contrast to the cytoplasmic SulA. In addition,
Rv2719c and YneA contain periplasmic domains with putative peptidoglycan-binding motifs,
and Rv2719c exhibits cell wall hydrolase activity in vitro. It is thus quite plausible that these
inhibitors also target FtsI, FtsW, or another late-acting divisome component, almost all of which
possess transmembrane and/or periplasmic domains. The three-protein complex SpoLID, SpolIM,
and SpoIIP in B. subtilis is also thought to block cytokinesis at a late stage in mother cells during
sporulation (Eichenberger et al., 2001).
Why target divisome components other than FtsZ? We speculate that the regulation of late
cell division events may be more effective at inhibiting division in cells that already have fully
formed, stable FtsZ rings. Targeting a late-acting divisome component may be easier for the cell
than dismantling an existing cytokinetic ring. Interestingly, Z-ring assembly occurs much earlier
during the cell cycle in Caulobacterthan in E. coli, perhaps necessitating a target other than FtsZ
(Quardokus et al., 2001). Alternatively, this strategy may prevent the wasteful disassembly and
reassembly of the cell division machinery during periods of transient DNA damage. Modulating
the activity of cell division proteins may also be a more effective and reliable means of inhibiting
83
cytokinesis than blocking component localization or assembly. The efficacy of inhibiting the
activity of late-stage cell division proteins is reinforced by the fact that many antibiotics, such as
cephalexin, target FtsI activity to effectively disrupt cell division in many bacteria.
Redundancies in cell division regulation
Is SidA the only mechanism for inhibiting cell division following DNA damage in
Caulobacter?The fact that a sidA deletion almost completely suppressed the cellular
filamentation of AlexA mutants argues that SidA is the primary SOS-induced cell division
inhibitor. However, the increase in aberrant cell divisions following DNA damage in AsidA cells
was modest and the majority of AsidA cells did not divide, suggesting that additional
mechanisms for inhibiting cell division must exist. As noted, a similar situation exists in E. coli,
where sulA mutations suppress the filamentation of AlexA strains but sulA mutants still filament
following DNA damage (Hill et al., 1997).
One possible additional mechanism is the damage-dependent down-regulation of CtrA,
which controls the expression of the cell division genesftsQ andftsA (Fig. 2. 1B; (Wortinger et
al., 2000)). The down-regulation of CtrA may prevent divisome assembly in cells that have yet to
synthesize FtsQ or FtsA, while SidA may block constriction in damaged cells that have already
assembled the divisome. Consistent with this model, we found that mixed populations of cells
treated with DNA-damaging agents exhibited varying levels of pinching, possibly reflecting
whether a divisome was fully formed when DNA damage occurred (Fig. 2. 1B).
The presence of unreplicated DNA at mid-cell following DNA damage may also prevent
cell division. Such a mechanism, termed "nucleoid occlusion," has been described in E.
coli and B. subtilis, where the proteins SlmA and Noc, respectively, bind throughout the
chromosome to inhibit FtsZ assembly in nucleoid-proximal areas (Bernhardt and de Boer, 2005;
Wu and Errington, 2004). In Caulobacter,the nucleoid occlusion model was initially dismissed
because Z-rings form when the nucleoid is still present at mid-cell. However, a nucleoid
84
occlusion-like mechanism could exist that inhibits later stages of cell division rather than FtsZ
assembly.
Finalperspectives
The discovery of SidA also underscores the increasingly recognized role of small proteins in
bacterial regulatory responses. Bacterial genomes encode scores of small (<50 amino acids)
proteins that have been missed in genome annotation projects or eluded identification in genetic
and proteomic screens. Recent studies have begun to systematically identify these small proteins,
and, intriguingly, a disproportionate number are predicted to be single-pass transmembrane
proteins that accumulate in stress conditions (Hemm et al., 2010; Hemm et al., 2008). In nearly
all cases, the functions of these small proteins are unknown, but some could target the cell
division machinery like SidA does.
Finally, while cell division in Caulobacterhas been known to depend on DNA replication
and chromosome segregation (Degnen and Newton, 1972), dependencies are not necessarily
enforced by dedicated surveillance or checkpoint systems (Hartwell and Weinert, 1989). The
identification of SidA demonstrates that Caulobacterdoes, in fact, employ checkpoint systems to
regulate its cell cycle under times of stress and DNA damage. In E. coli, SulA and UmuD
mediate DNA damage checkpoints (Huisman and D'Ari, 1981; Opperman et al., 1999), and, in B.
subtilis, a developmental checkpoint couples the status of DNA replication to the initiation of
sporulation (Burkholder et al., 2001). Cell cycle checkpoints that coordinate the timing and order
of cell cycle events to maintain genome integrity thus appear to be a common and powerful
regulatory strategy in bacteria, as in eukaryotes.
85
Materials and Methods
Strains, plasmids, and growth conditions
Strains and plasmids used are listed in Supplemental Table S2 in (Modell et al., 2011), with
construction details and growth conditions provided in the Supplemental Material.
DNA microarrays
RNA expression profiling was done as described previously (Biondi et al., 2006). Complete
data sets are provided in Supplemental Table SI in (Modell et al., 2011). Genes selected for
Figure 2.1 had expression values that increased or decreased by twofold or greater in at least four
of the six conditions tested.
Microscopy
For time-lapse microscopy, cells were immobilized on 1% agarose pads made with PYE
medium. Images were acquired on a Zeiss Axiovert 200M microscope with a 63 x phase or
aFluar 100x/1.45 objective using an Orca II ER camera. For epifluorescence, illumination was
provided by an HB0103 arc lamp using the following emission/excitation filters: YFP, S500/20x
and S535/30m; GFP, S470/40x and 520/40m; mCherry, S572/35x and S632/60m. All image
capture and processing was done through Metamorph software (Universal Imaging Group).
Immunoblots and biochemical fractionations
Immunoblots were performed as follows: Samples were normalized in sample buffer to 0.5
OD 600 /50 pL, resolved on 12% sodium dodecyl sulfate-polyacrylamide gels, and transferred to
polyvinylidene difluoride transfer membrane (Pierce). Membranes were probed with polyclonal
rabbit a-CtrA and a-GFP (Invitrogen) at a 1:5000 dilution and monoclonal mouse a-Flag
(Sigma) at a 1:1000 dilution. Secondary HRP-conjugated a-rabbit (Pierce) or a-mouse (Pierce)
were used at a 1:5000 dilution.
86
Biochemical fractionation was performed as described (Mbll and Thanbichler, 2009) with
several modifications. Samples from each culture were adjusted to total 8 mL*OD 600 units,
washed once with 0.2 M Tris-HCl (pH 8), and resuspended in 1 mL of 60 mM Tris-HCl (pH 8),
0.2 M sucrose, 0.2 mM EDTA, 200 tg mLU lysozyme, and 5 U of DNase I. The resuspension
was incubated for 10 min at room temperature, frozen at -80'C, thawed, and sonicated for 10
sec. Intact cells and chromosomal DNA were pelleted by centrifugation at 4000g for 10 min.
Membranes were then pelleted by centrifugation at 126,000g for 1 h at 4'C. Two-hundred
microliters of the supernatant representing the soluble fraction was diluted in 4x SDS sample
buffer. The membrane pellet was washed once with 0.2 M Tris-HCl (pH 8) and resuspended in
200 pL of 1 x SDS sample buffer. Equal volumes from each fraction were loaded for immunoblot
analysis.
Identification of SidA overproduction suppressors
Wild-type cells were transformed with plasmid pML1716-sidA and plated on PYE
supplemented with chloramphenicol and xylose. Single colonies were grown overnight in PYE
containing chloramphenicol and xylose and samples were taken for immunoblots, plasmid
preparations, and archiving. To isolate chromosomal suppressor mutations and eliminate
mutations arising in the sidA overexpression plasmid, we screened for colonies that met three
criteria. (1) We used immunoblotting to check that SidA production in each suppressor strain
was similar to that seen in wild-type cells transformed with the pMLI716-sidAplasmid and
grown in xylose for 1.5 h. (2) Plasmids from the suppressor strains were transformed into wildtype cells and plated on PYE supplemented with xylose or glucose. The presence of thousands of
colonies on glucose plates and few colonies on xylose indicated a functional plasmid. (3)
Plasmids from the suppressor strains were sequence-verified.
87
Nine strains fulfilled the three criteria above and were deemed likely to harbor chromosomal
suppressors. We sequenced the essentialfts genes from each strain. Mutations identified
infts W andftsI were then engineered into a clean wild-type background by allelic replacement.
Bacterial two-hybrid analysis
Two-hybrid complementation assays were performed essentially as described (Karimova et
al., 2005). OD 42 0 readings were obtained with a Spectramax 340PC 384 plate reader (Molecular
Devices), and the average rate (dOD 420/dt) during the 10- to 20-min time points was used to
calculate enzyme activity.
Van-FL labeling
Van-FL (Molecular Probes) was prepared in DMSO at 3 mg/mL and cells were labeled at a
final concentration of 3 pg/mL for 20 min. Cells were then pelleted at 13,000 rpm for 8 sec,
washed in PBS, and imaged directly by DIC or phase and epifluorescence microscopy.
Fluorescent bands and foci were measured in ImageJ (http://rsbweb.nih.gov/ij), and those with
intensities at least 1.5-fold over the cell background were scored as significant (see also Fig.
2.11).
88
Acknowledgements
We thank M. Prasol for help with the expression profiling, and A. Grossman, J. Dworkin,
and members of the Laub laboratory for comments on the manuscript. We acknowledge M.
Thanbichler and C. Jacobs-Wagner for strains. M.T.L. is an Early Career Scientist at the Howard
Hughes Medical Institute. This work was supported by an NIH grant (5R0 1 GM082899) to
M.T.L. and an NSF predoctoral fellowship to J.W.M.
Note added in proof
In B. subtilis, the cell division inhibitor Maf was also shown recently to directly target a
protein other than FtsZ, called DivIVA (Briley et al., 2011).
89
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92
Chapter 3
A DNA damage-induced, SOS-independent checkpoint regulates cell
division in Caulobacter crescentus
This work is in submission as Modell, J. W., Kambara, T. K., and Laub, M. T. A DNA damageinduced, SOS-independent checkpoint regulates cell division in Caulobactercrescentus.
JWM, TKK and MTL conceived and designed the experiments. JWM and TKK performed the
experiments.
93
Abstract
Cells must coordinate DNA replication with cell division, especially during episodes of
DNA damage. The paradigm for cell division control following DNA damage in bacteria
involves the SOS response where cleavage of the transcriptional repressor LexA induces a
division inhibitor. However, in Caulobactercrescentus, cells lacking the primary SOS-regulated
inhibitor, sidA, can often still delay division post-damage. Here we identify didA, a second cell
division inhibitor that is induced by DNA damage in an SOS-independent manner. DidA does
not disrupt assembly of the division machinery and instead binds the essential division protein
FtsN to block cytokinesis. Intriguingly, mutations in FtsW and FtsI, which drive the synthesis of
septal cell wall material, can suppress the activity of both SidA and DidA, likely by causing the
FtsW/I/N complex to hyperactively initiate cell division. Finally, we demonstrate that DidA and
SidA act synergistically to inhibit division as cells lacking both inhibitors divide prematurely
following DNA damage, with lethal consequences.
94
Introduction
Progress through the cell cycle requires the sequential execution of three fundamental
processes: DNA replication, chromosome segregation, and cell division. Maintaining the precise
order of these events is crucial to preserving genomic integrity, as any attempt to divide before
completing DNA replication or chromosome segregation could result in the scission of DNA and
a failure to endow each daughter cell with a complete genome. Coordinating DNA replication
and cell division is particularly challenging when cells encounter DNA damaging agents that
necessitate lengthy periods of chromosome repair before replication can finish and cell division
can resume. To ensure the order of cell cycle events and preserve genome integrity, many cells
employ checkpoints that actively halt cell cycle progression until DNA damage has been
repaired. While checkpoints are prevalent and well-characterized in eukaryotes (Harper and
Elledge, 2007), their role and significance in governing the bacterial cell cycle is less clear.
The ca-proteobacterium Caulobactercrescentus, which replicates once and only once per cell
division, is an excellent system for understanding the bacterial cell cycle. In particular, cells are
easily synchronized and DNA replication initiates only once per cell division, resulting in
distinguishable GI, S, and G2 phases. As with most bacteria, cell division in Caulobacter
involves the assembly of a large multi-protein complex at mid-cell that drives constriction of the
cell envelope and separation of daughter cells (Goley et al., 2011). The position of the division
machinery is established by the tubulin homolog FtsZ, which forms a ring-like structure at midcell and subsequently recruits other essential cell division proteins (Goley et al., 2011;
Quardokus et al., 2001; Wortinger et al., 2000). Once assembled, how these proteins coordinate
the various steps of cytokinesis is unclear and the factor(s) that ultimately trigger cytokinesis are
unknown.
95
Like eukaryotes, bacteria can inhibit cell division following DNA damage. The best-studied
mechanism involves the so-called 'SOS response' (Erill et al., 2007; Little, 1991) in which DNA
damage stimulates the recombinase RecA to trigger an auto-catalytic cleavage of the
transcriptional repressor LexA. This cleavage leads to induction of SOS genes, many of which
are involved in DNA recombination and repair (Butala et al., 2009; Little, 1991). The SOS
regulon also typically includes a cell division inhibitor that can delay cytokinesis until after
damage is cleared. The best characterized SOS-induced division inhibitor, E. coli SulA, disrupts
polymerization of FtsZ and thus inhibits assembly of the cell division machinery, known as the
"divisome" (Huisman and D'Ari, 1981; Mukherjee et al., 1998). However, sulA is not widely
conserved beyond the y-proteobacteria and recent studies have indicated that the SOS-induced
division inhibitors from several Gram-positive species do not target FtsZ, although in most cases
the direct target remains unknown (Chauhan et al., 2006; Kawai et al., 2003; Ogino et al., 2008).
In Caulobacterthe primary SOS-induced division inhibitor is a 27 amino acid inner membrane
protein called SidA that inhibits division by interacting directly with the late-arriving division
protein FtsW (Modell et al., 2011).
Although sidA is the primary SOS-induced division inhibitor in Caulobacter,cells lacking
sidA can still arrest division when grown in the presence of the DNA damaging agent mitomycin
C (MMC). An SOS-regulated endonuclease called BapE may indirectly contribute to inhibiting
division (Bos et al., 2012), but we conjectured that Caulobacterlikely encodes another direct cell
division inhibitor that is induced by DNA damage but in an SOS-independent manner. Here we
identify such an inhibitor, now named didA, that is induced following DNA damage even in
strains that cannot mount an SOS response, demonstrating the existence of an SOS-independent
transcriptional response to DNA damage. Like sidA, the overexpression of didA in undamaged
cells is sufficient to prevent cell division. DidA does not disrupt FtsZ ring formation or divisome
96
assembly and instead likely inhibits division through a direct interaction with the divisome
component FtsN. Intriguingly, point mutations infts W andftsI suppress the lethality that results
from overproducing either SidA or DidA. Our results suggest that these mutations hyperactivate
the cell division process and implicate the protein complex FtsW/I/N in the triggering of
cytokinesis. Finally, we show that Caulobactercells are highly sensitized to MMC only when
both sidA and didA are deleted indicating that under certain conditions of DNA damage, these
inhibitors are functionally redundant.
97
Results
Identification of didA, a DNA damage-induced, SOS-independent cell division inhibitor
Our previous work demonstrated that sidA is the primary SOS-induced division inhibitor in
Caulobacter.However, many AsidA and ArecA cells exposed to the DNA damaging agent
mitomycin C still become filamentous suggesting that another, SOS-independent inhibitor may
also prevent division following DNA damage (Fig. 3.1; (Modell et al., 2011)). To identify
candidate SOS-independent division inhibitors, we examined global gene expression changes
following mitomycin C (MMC) treatment of a ArecA strain, which cannot induce SOS genes.
Wild-type and ArecA cells were grown to mid-exponential phase in rich medium and exposed to
MMC for 30 minutes. RNA was then isolated and compared to mock treated cells on wholegenome DNA microarrays (Table S1 in submitted manuscript).
-MMC
+ MMC
wt
AsidA
ArecA
Figure 3.1. Filamentation in sidA and recA mutants. Wild-type, AsidA and ArecA cells were grown to midexponential in PYE and treated with 1 pg/mL MMC or left untreated. After 3 hours, cells were imaged by phase
microscopy.
98
Of the 50 most up-regulated genes following MMC treatment in wild-type cells, 44 were
recA-dependent, including 31 that are directly regulated by LexA (da Rocha et al., 2008; Modell
et al., 2011) (Fig. 3.2A, 3.3A). The remaining 6 damage-regulated genes showed similar
induction levels in both wild-type and ArecA backgrounds (Fig. 3.2A) and are thus likely
controlled by an SOS-independent mechanism. One of these genes, CCNA03212 in the NA 1000
(CB1 5N) genome, encodes a previously uncharacterized 71 amino acid protein with a single
predicted transmembrane helix flanked by short cytoplasmic and periplasmic domains (Fig.
3.2B). The open reading frame of CCNA03212 overlaps with the C-terminus of the open reading
frame of CC3114 from the annotated genome of CB 15. In our expression profiling experiments,
only those probes lying within the CCNA03212 coding sequence were significantly upregulated
by MMC in wild-type cells (Fig. 3.3B-C), suggesting that the NA1000 annotation is correct.
Based on the studies described below, we named this gene didA for damage-induced cell division
inhibitor A.
To confirm that the didA locus encodes a damage-inducible protein, we created a strain in
which the chromosomal didA coding sequence was fused at its C-terminus to the coding region
of the 3xFLAG (3xF) epitope. DidA-3xF was barely detectable in the absence of DNA damage,
but was strongly induced following MMC treatment with protein levels increasing nearly 20-fold
after 1 hour (Fig. 3.2C). Western blotting indicated a band at the size predicted for DidA-3xF
(~1 1 kDa) and not CC3114-3xF (-25 kDa) indicating that the larger gene product annotated in
CB 15 is likely not produced at significant levels. To test the SOS-dependence of DidA-3xF
synthesis following MMC treatment, we examined DidA-3xF production in a ArecA strain and in
a strain harboring lexA(K203A), which encodes a non-cleavable form of LexA that blocks the
induction of an SOS response. In each case, DidA-3xF was slightly elevated in untreated cells,
likely due to increased basal levels of damage in the absence of SOS-mediated repair (Fig. 3.2D).
Following MMC treatment, DidA-3xF was strongly induced in all strains (Fig. 3.2D), consistent
with an SOS-independent mode of regulation.
99
A
1.1
1.0.-
i0.6
.
~d~
0*
wt + MMC
log(fold Induction)
-0.26
B
6
sidA
CC3114CCNA03212
-- (ddA)--4
1
386
C
600
min. In MMC
0
16
30
4
60
90
120
160
DIdA-3xF
D
.
I
-
+
IxA(K203A)
AreMA
wt
MMC:
-
+
+
DidA-3xF
Figure 3.2. didA is induced by DNA damage and is not SOS regulated. (A) Wild-type and ArecA cells were grown
in a rich medium (PYE) to mid-exponential phase and treated with 1 pg/mL MMC for 30 minutes. Expression
values, calculated from the average of two biological replicates, are shown for the 50 most upregulated genes in
wild-type cells with fold-change ratios calculated in comparison to mock treated cells. Expression values in wildtype cells are plotted on the x-axis and those in ArecA cells are plotted on the y-axis. The dashed line corresponds to
fold-change values that are identical in wild-type and ArecA cells. For complete data, see Figure 3.3 and Table SI in
submitted manuscript. (B) CC3114 and CCNA03212 (didA) are shown schematically in their genomic context.
Nucleotide positions relative to the annotated CC3114 start site are shown below. The gray shaded region represents
a predicted transmembrane domain (TMHMM). (C) Western blot of cells expressing didA fused to a C-terminal
3xFLAG epitope at the native didA chromosomal locus. Cells were grown in PYE to mid-exponential phase and
treated with 1 pg/mL MMC for the times indicated. (D) Western blot of wild-type, ArecA and lexA (K203A) cells
expressing didA-3xFLAG from its native locus treated with 1 ptg/mL MMC for 1 hour. Membranes (C-D) were
blotted with the a-FLAG antibody.
100
A
4/
IT
+MMC
ArcA
wt
CC 1827
CC_3424
CC_1330
CC_1302
CC_1i
CCW382
CC_3710
CC_7?S
C C_1054
C C_33
CC_514
CCz15
CC_105?
CCCWO
SWdA
frily proton
SAM pupe sily protein
LGCArSpror
Hypoahetod proten
DNA mihyladonine glycoyWo
2-polyprenylphenol G-hydrox)409 WCo oryprotein ubIB
writoxin protein ra-3
wftoxin protein parD-2
TonrB.C
oier moribranersoptor
Hyp tilloo proton
Hypotreatod proton
RsoAprotein
toxin protein rUE-3
GIyoxves
Red"I
OcMne_212 dIdA
C C.248
Conservid hypolttllool protein
CC2
Endonucleie, III
CC_10$
Hypothlleod proton
CC_3038
toxnprotinhIgB
CC_27
Hypdh@od protein
CC_3213
Conwrd hypotbuiloI protein
C C3515
HypotheboU 0tooollo proton
CC_1531
Hypothlooll protoin
HypotietIool nombrano usoolded protein
CC2311
CC_;211
DNApolymffer
1101phe mtnft
CC_;M2
Bln syntrhe reaed domin corening preton
CC 0-37 Proedtransoriptiona reguldoe, ntitoxin proton nigA
CC_3W
Hypotllom cyobolic proton
Urvoll OKA glyoyeapraWIly protein
CC_2333
CC_34G7
Hypothetloi protein
CCW1S
ReoAprotemn
CC_;212
Hypothetioi cytoooll pretein
CC_0140 ATPove ried to mnlgnolumcholdmloosubudt Chi
CC_2?64 TrneorphIoni regAmor
CC34
Sholt-ohlin oyl- CoAhydrolam
CC_-271
I muCtomilly protein
CC_2M40
ATPd@PndMt hlo411
CC-""SS DNA reair protein rooN
CC-152
DNAhuoallS 11
CC_3131 toxin protein rolE-4
CC_2D41
pmAtaIVI ICC-11 phOph0tere
CC_1331
Mycoyttrun meInvolvd In call Yel biogeneele
CC_22
Gyoxioso fertiy proton
CC238
Crouow)Indion andodeoxyribonulodee ruC
Tio-0oMpone-nt reopones regmidor
CC_2?0O
Hypotfetlom protein
CC.0G20
CC_130
Hypthello proten
CC_33
Rhodmneu-rueloed mAfUrtrenWerer
CC_32
Hypothetilo protin
CC 2530 Exdrude
ABC mubunt A
log Told change
1.5i
0.5I
0.51
1.0
1.5
B
C
10
0.0
CC3114
CCNAM3212
'I-
Ii
I
385
"""
TM
600
0.6
OA
02
01
-02
.. M|"ss"""
Figure 3.3. Annotated gene expression profiles. (A) Transcriptional profiles for the 50 most up-regulated genes
during DNA damage in wild-type cells (see Fig. 3.2A) are shown with their corresponding CC numbers and
NA 1000 annotation. The 'LexA' column shows genes whose upstream region contains a sequence match to 7 of the
8 bases in the CaulobacterLexA consensus binding site (GTTCN 7GTTC) (da Rocha et al., 2008). Genes whose logfold changes post-damage in ArecA cells are below 50% of those in wild-type cells are marked as 'RecAdependent'. All other genes are marked as 'RecA-independent.' (B) The positions of microarray probes representing
101
CC3114 and CCNA03212 are shown below the genes as horizontal bars. The four right-most probes were used to
calculate expression values for CCNA03212 (didA). (C) The transcriptional profiles for each probe in panel B are
shown.
To test whether DidA is a cell division inhibitor, we fused the didA coding sequence to the
vanillate-inducible promoter Pvan and cloned this construct into both a low- and medium-copy
plasmid. We transformed wild-type cells with each plasmid and then grew cells in the presence
of vanillate to induce didA in the absence of a DNA damaging agent. Synthesis of DidA from a
low-copy plasmid resulted in mild cellular filamentation and a modest growth defect, while
overproduction from the medium-copy plasmid caused a more pronounced division defect with
nearly all cells demonstrating severe filamentation after 6 hours (Fig. 3.4A-B). Thus, DidA, like
SidA, is sufficient to inhibit cell division in the absence of DNA damage. Taken together, our
results suggest that following DNA damage, DidA accumulates in an SOS-independent fashion
to help prevent cell division.
A
B
1.2
Pva,:didA
1.0
low-copy
0.8
:
0
0
-0-
empty vector
-W-
P.:ddA
(low-copy)
0.6
medium-copy
0.4
- van
0.2
0
0-
2
4
6
8
10
12
time (h)
1.2
1.0
0.8
0
empty vector
+
+
-a- P,*:ddA
(medIum-copy)
0.6
van
(6 h)
0.4
0.2
0
2
4
6
8
10
12
time (h)
102
Figure 3.4. DidA is sufficient to inhibit cell division. Growth curves (A) and micrographs (B) of strains
overexpressing didA. Cells harboring a low- or medium-copy plasmid that expresses didA from the vanillateinducible promoter P, were grown in PYE in the presence or absence of vanillate for the times indicated. Bar, 2
[pin.
DidA interacts with the late-arriving divisome component FtsN
We next sought to establish whether DidA interferes with cell division directly, through an
interaction with the divisome, or indirectly by inducing the SOS regulon or inhibiting the cell
cycle regulator CtrA. To investigate the possibility of the latter, indirect mechanisms, we isolated
RNA from cells overproducing DidA from a medium-copy plasmid for 45 minutes and
compared it on DNA microarrays to RNA from similarly treated cells grown in the absence of
inducer. No significant gene expression changes were observed in the SOS or CtrA regulons
(data not shown) suggesting that DidA acts post-transcriptionally, and possibly directly, to
inhibit cell division.
To further explore how DidA inhibits cell division, we examined its subcellular localization.
In predivisional cells, the major components of the cell division machinery are located at midcell (Goley et al., 2011) where they synthesize a septum and drive invagination of the cell
envelope. To assess DidA localization, we transformed wild-type cells with a low-copy plasmid
harboring an M2-yfp-didA fusion under the control of a xylose-inducible promoter, Pry. After
induction for 3 hours, cells were typically more filamentous than those expressing untagged didA
from the same plasmid for 6 hours (Fig. 3.4B, 3.5A), suggesting that the YFP-DidA fusion is
functional and perhaps more stable or potent than DidA alone. Notably, YFP-DidA foci were
frequently observed at pinch sites near mid-cell (Fig. 3.5A) placing it in close proximity to the
cell division machinery. Further, the fractionation of cells overproducing M2-DidA from a
plasmid indicated that DidA is strongly enriched in the membrane where many of the middleand late-arriving cell division components also reside (Fig. 3.5B). These data are consistent with
a model whereby DidA inhibits division through a direct interaction with a component of the
division apparatus.
103
A
C
P,-M2-yfp-ddA
+
+ xyl (1.6 h)
glu
+xyl
(3 h}
T26-DIdA +
T18-
FIsZ
T18-
FtsW
Ft6A
FtsK
FISQ
FtSL
FtsB
4-
+1+
I
FI
Ft&N M2-SidA
T25-IdA +
B
P.:3xF-9dA
- van
PM
+ van
S
T25.t5N
D
M
P.:dfdA +
Pv*rfftZjYf
CckA-GFP
fiP-ftl
venus-ftsW
gtp41tN
I.
CtrA
3xF-DidA
Pme fAvn
a.
IA.
E
a.U.
Wt
I
w'.*PWf
IWI-t
9048
Figure 3.5. DidA is a small, inner-membrane protein that interacts directly with FtsN. (A) The subcellular
localization of DidA was examined in a strain expressing m2-yfp-didA from the xylose-inducible promoter Pxy on a
low-copy plasmid. Cells were grown to mid-exponential phase in PYE with glucose and then shifted to xylose. At
the times indicated, cells were imaged by phase and epifluorescent microscopy. (B) Subcellular fractionation of cells
overexpressing 3xFLAG-didA from the Pvan promoter on a medium-copy plasmid for 1.5 hours and expressing the
transmembrane protein cckA-gfp from the chromosome. Samples were fractionated into soluble (S) and membrane
(M) fractions and analyzed by Western blot. The membrane was cut into three pieces, indicated by dashed lines, and
probed with antibodies specific for the GFP, CtrA, or M2 epitope. (C) Bacterial two-hybrid analysis of interactions
between T25-DidA and cell division proteins fused to T18, as indicated. The interacting pair TI 8-M2-SidA and
T25-FtsN was included for comparison. E. coli strains harboring each pair of fusions were plated on LB, and
colonies were restruck on MacConkey plates containing maltose. Red streaks indicate positive interactions. (D) Subcellular localization of FtsZ, FtsW, FtsI, and FtsN were examined in strains expressing ftsZ-yfp from the
chromosomal P,,n promoter, or venus-fts W, gfp-ftsI or gfp-fisN from its native chromosomal locus. Each strain was
transformed with a medium-copy plasmid expressing didA from the Pvan promoter. Strains were grown to midexponential phase in PYE and samples imaged by phase and epi-fluorescent microscopy after addition of vanillate
for 4.5 hours. In the fluorescent images, cell outlines were drawn based on the phase micrographs. Bars, 2 pm. (E)
The strains from panel D were grown to mid-exponential phase and 10-fold serial dilutions were plated on PYE
supplemented with vanillate to induce didA expression.
104
Although many regulators of cell division disrupt positioning or assembly of the FtsZ ring,
several SOS-induced inhibitors, including SidA, do not. To test for direct interactions of DidA
with the known set of critical Caulobactercell division components (Goley et al., 2011), we
performed a bacterial two-hybrid analysis with the reconstituted adenylate cyclase system used
previously for SidA (Karimova et al., 2005; Modell et al., 2011). When expressed from the lowcopy plasmid pKT25, a T25-DidA fusion interacted almost exclusively with the late-arriving cell
division protein fusion T18-FtsN (Fig. 3.5C, 3.6). Identical results were obtained with a T18DidA fusion on the high-copy plasmid pUT I8C and individual division proteins produced from
the low-copy plasmid pKT25, although the DidA-FtsN interaction was slightly weaker in this
orientation (data not shown). SidA, whose primary target is FtsW, also interacts, to some extent,
with FtsN; this interaction was at least as strong as the DidA-FtsN interaction, as judged by this
two-hybrid assay (Fig. 3.5C). In sum, our data suggest that DidA is an integral membrane protein
that localizes to mid-cell where it may disrupt cell division through a direct interaction with
FtsN.
T25:DidA
T18:
T18:
FtsN
FzIA
DipM*
KidO
FtsX
FzIC
FtsE
ZapA
/-
+/+
T25:DidA
Figure 3.6. DidA interacts directly with FtsN. Bacterial two-hybrid analysis of interactions between T25-DidA and
cell division proteins fused to T18, as indicated. Each pair was plated on LB, and colonies were restruck on
MacConkey plates containing maltose.
105
FtsN is among the last cell division proteins to arrive prior to cytokinesis. Although its
precise molecular function is unknown, FtsN interacts with multiple division proteins and is
thought to play a key role in stabilizing the assembled divisome (Goehring et al., 2007;
Karimova et al., 2005; Rico et al., 2010). To ask whether DidA destabilizes or blocks assembly
of the cell division machinery through its interaction with FtsN, we examined the localization of
early- and late-arriving division proteins during DidA overproduction. Cells producing
fluorescently tagged FtsZ, FtsW, FtsI, or FtsN were transformed with a plasmid for
overexpressing didA and then grown in the presence of vanillate to induce DidA synthesis. After
4.5 hours of induction, cells expressingftsZ-yfp, venus-fts W, or gfp-ftsI were inhibited for cell
division, but 89%, 95% and 85% of cells, respectively, contained fluorescent foci at or near
visible pinch sites. These results indicate that DidA likely does not disrupt the localization of cell
division proteins or drive the disassembly of division protein complexes (Fig. 3.5D).
Additionally, we noted that many cells displayed multiple foci of the FtsZ, FtsW, or FtsI
fluorescent fusions, a pattern rarely seen in the absence of DidA; this result suggests that DidA
also does not prevent the formation of new division assemblies.
Intriguingly, cells expressing gfp-ftsN were noticeably shorter and more pinched than those
expressing the other fluorescent fusions (Fig. 3.5D). Further, cells expressing gfp-ftsN were able
to robustly form colonies despite DidA overproduction, in contrast to cells expressing the other
fluorescent fusions or the wild type (Fig. 3.5E). The GFP-FtsN fusion therefore acts as a DidA
suppressor, possibly by decreasing its affinity for DidA or by stabilizing FtsN and thereby
increasing FtsN levels. Whatever the case, these data further support a model in which DidA
interacts with FtsN to block cell division but without disrupting assembly of a full, intact division
apparatus.
106
Mutations in ftsW can suppress the division inhibition caused by either SidA or DidA
To further explore the mechanism by which DidA inhibits division, we screened for
spontaneous mutations that suppress the lethality of overproducing DidA. Wild-type cells
carrying a medium-copy plasmid expressing 3xF-didA from P..n were grown on plates
containing vanillate to induce M2-DidA synthesis. Because wild-type cells overproducing M2DidA cannot form colonies (Fig 3.7B), those rare colonies arising on vanillate plates represent
strains harboring putative suppressor mutations. From roughly 3 x 107 plated cells, 34
suppressors were identified but only one strain retained high levels of functional M2-DidA.
Whole genome resequencing identified a putative suppressor mutation infls W, which would
produce the substitution A246T in the predicted large periplasmic loop of FtsW (Fig. 3.7A). This
fts W mutation was created de novo in a clean CB 1 5N background and confirmed to suppress the
lethality of overproducing DidA. As noted, no interactions between DidA and FtsW were
observed in our two-hybrid analysis; thus FtsW(A246T) likely suppresses DidA overproduction,
not by preventing binding of the inhibitor, but by promoting an activity of FtsW (Modell et al.,
2011).
Intriguingly, we had previously found mutations infts W that suppress the lethality of
overproducing SidA. We therefore reasoned that SidA and DidA may function similarly to
inhibit cell division. To explore this possibility, we asked whether other, previously identified
suppressors of SidA overproduction could also suppress DidA overproduction, and vice versa.
Two mutations primarily suppressed the lethality of only one of the inhibitors; the FtsW(T 1 80A)
strain strongly suppressed overproduction of M2-SidA but not DidA, whereas the strain
producing a GFP-FtsN fusion suppressed the activity of DidA but not M2-SidA. These inhibitorspecific suppressors may prevent binding of their respective inhibitors. The other mutations
examined showed varying abilities to suppress the lethality associated with overproducing either
inhibitor. In particular, the strains producing FtsW(F 145 L) or FtsW(A246T) showed robust
suppression of both inhibitors.
107
A
FtsW
FtsN
FtsI
FISdA
F14L
GFP
wt
ftW(F145L)
ftl(545V)
ftsW(A31K)
ftsW(T18OA)
ftsW(A246T)
gfpfN
Figure 3.7. Mutations infts W suppress SidA and DidA overproduction phenotypes. (A) Schematic showing the
membrane topology of the division proteins FtsW, FtsI, and FtsN, as well as the division inhibitors SidA and DidA.
Missense mutations and the GFP-FtsN fusion that suppress the activities of SidA or DidA, or both, are listed in red.
(B) The suppressor mutations from panel A were introduced into a wild-type strain by allelic replacement followed
by transformation with a medium-copy plasmid expressing either M2-sidA from the Pry promoter or didA from the
Pva promoter. To induce M2-sidA, strains were grown in PYE supplemented with glucose and then plated on PYE
supplemented with xylose. To induce didA, strains were grown in PYE and plated on PYE containing vanillate.
Each strain was plated in 10-fold dilutions.
The ability of these single substitutions, F145L and A246T, to suppress the lethality of
overproducing either SidA or DidA could indicate that the inhibitors share a binding site within
FtsW that is disrupted by the suppressor mutations. However, we viewed this as unlikely given
that (1) DidA does not bind FtsW in our bacterial two-hybrid system and (2) M2-SidA binds to
FtsW(A246T) as strongly as it does to wild-type FtsW (Fig. 3.8). As an alternative explanation,
we hypothesized that the sub-complex of late-arriving division components FtsW, FtsI, and FtsN
could exist in one of two conformations: an active conformation that promotes constriction of the
inner membrane and septum, and an inactive conformation that is promoted or stabilized by
108
SidA and DidA. In this model, the suppressor mutations infts W andftsI would promote the
active conformation and thus enable cell division even in the presence of SidA and DidA.
T1S:M-SidA
T26:
FtsW
FtSW
FtsW
FtBW
(A31K)
(F146L)
(TISOA)
FSW
(A246T)
Figure 3.8. SidA interacts directly with FtsW. Bacterial two-hybrid analysis of interactions between T 18-M2-SidA
and FtsW mutants fused to T25 as indicated. Colonies were grown to exponential phase in LB and 5 ptL aliquots
plated on MacConkey agar containing maltose.
SidA and DidA suppressor mutations drive hyperactive cell division
If the FtsW(F145L) and FtsW(A246T) mutations promote an active conformation of a
subcomplex of cell division proteins, cells harboring these mutations, but not producing SidA or
DidA, may attempt division at an earlier stage of the cell cycle than wild-type cells, even in the
absence of DNA damage. To explore this possibility, we grew strains harboring one of the
suppressor mutations infts W, or inftsl, into mid-exponential phase in rich medium and measured
cell lengths in a large population of cells. Indeed, many of the suppressor mutations resulted in
cells that were significantly shorter on average than the wild type even though their growth rates
were not substantially different (Fig. 3.9A-B, 3. 1A-B). The degree of shortening roughly
correlated with the strength of suppression, as cells harboring the mutationsfts W(A246T),
ftsW(F145L), andftsI(I45V) that were best able to suppress both SidA and DidA were also the
shortest. ForftsW(A246T), which yielded the strongest suppression, all cell types were, on
average, ~0.3 microns shorter than wild-type cells (Fig. 3. 10B). These observations suggest that
thefts W(A246T) strain is not simply enriched for swarmer cells, the shortest Caulobactercell
type, but that these mutant cells divide as shorter predivisional cells resulting in shorter swarmer
and stalked daughter cells.
109
A
B
cell length (%wt)
88
wt
ftsW(A246T)
92
96
100
wt
ftsW(A246T)
fti(145V)
ftW(Fl45L,)
ftW(TI80A)
ftWs(A31K)
fti8(F58V)
C
+ MMC
+ cephalexin
+ novobiocin
wt
fsW(A246T)
AsIdA AdIdA
Figure 3.9. Mutations that suppress sidA and didA overexpression likely hyperactivate cell division. (A-B) The sidA
and didA suppressor mutations, each introduced into a clean wild-type background by allelic replacement, were
grown to mid-exponential phase in PYE and phase images were taken. Bar (A), 2 pm. Cell lengths, relative to wildtype, were calculated for each strain (n > 440 for each strain); error bars (B) represent standard error of the mean.
(C) Wild-type, fts W(A246T), and AsidA AdidA cells were grown to mid-exponential phase and plated in 10-fold
dilutions on PYE containing 0.35 pg/mL MMC, 0.35 pg/mL novobiocin, or 7.5 pig/ml cephalexin, as indicated.
B
A
1.2
1.2.
1.0
-0- wt
1.0
0
0.8
gfp-ftmN
p0 ftW(fl8OA
0.8
+
a, 0.6
0
--
0.4!
0.2
ftW(F145L)
ftW(A31K)
Sft.W(A246)
0
0
2
4
6
hours
8
10
12
---
0.6
-4o- ftI(F56V
sW(A246)
0.4
C
0.2
0
1.9
2.5
3.1
3.7
4.3
4.9
5.6
6.2
microns
Figure 3.10. Suppressor mutant growth properties. (A) Wild-type andfts W(A246T) cells were grown to midexponential phase and imaged by phase microscopy. Cell lengths were quantified from 491 wild-type and 610
fts W(A246T) cells using MicrobeTracker and summarized as a histogram with the maximum frequency for each
strain normalized to 1. (B) Growth curves for the strains from Fig. 3.9C grown in PYE.
110
Another prediction of our model is that cells with the mutations causing hyperactive division
may be sensitive to other stresses that, like DNA damage, require a transient inhibition of cell
division. Novobiocin inhibits DNA gyrase and causes replication arrest without directly
damaging DNA. Cells must therefore inhibit division to prevent scission of unreplicated DNA
that remains at mid-cell. Cephalexin inhibits penicillin-binding proteins and prevents the
crosslinking of septal cell wall material. An attempt to activate divisome constriction without the
corresponding synthesis of septal cell wall could result in membrane or envelope stress. We
focused onfts W(A246T), the strongest suppressor of SidA and DidA overproduction, and found
that cells expressing this allele exhibited a viability defect when grown on plates containing
MMC, novobiocin, or cephalexin (Fig. 3.9C). These results are consistent with a model in which
fts W(A246T) cells cannot properly delay division when stressed.
SidA and DidA redundantly regulate division during MMC treatment
Given that SidA and DidA are induced concomitantly following MMC treatment, what are
the relative contributions of each inhibitor toward regulating cell division? Our previous work
demonstrated that AsidA cells show modest division defects during DNA damage, perhaps
because DidA can compensate functionally. To examine the roles of SidA and DidA during
DNA damage, we constructed a strain in which all but the first and last three amino acids of didA
were deleted. As with the sidA deletion strain, AdidA cells grown on plates containing MMC
showed no major viability defects (Fig. 3.1 IA). However, a strain lacking both sidA and didA
showed a pronounced viability defect with a nearly 100-fold decrease in plating efficiency (Fig.
3.1 IA) suggesting that at least one of the inhibitors is necessary to properly control cell division
post-damage. This viability defect was rescued by the presence of either inhibitor on a low-copy
plasmid (Fig. 3.11 B) confirming that in the presence of MMC, SidA and DidA are, to some
extent, functionally redundant.
111
- MMC
A
+ IWIVC
C
z
Wt
5
.1
AsIdA
I
AdidA
I
I
lididA tSIsdA
3.
- MMC
B
4
"V
aOIOA
+MMC
AsidA
Wt
D
30.
~i25-
tAdIdA tl8IdA
j20.
AdIdA AsidA + sidA
AdIdA AsIdA
+
*10
didA
5.
..
0.
00"M^
Issid
E
time grown on MMC pad (h)
0
I
2
3
4
5
6
wt
AdIdA
AsIdA
Figure 3.11. Cells lacking sidA and didA cannot properly regulate cell division following DNA damage. (A) Wildtype, AsidA, AdidA, and AsidA AdidA cells were grown to mid-exponential phase and plated in 10-fold dilutions on
PYE containing 0.35 ptg/mL MMC. (B) Wild-type and AsidA AdidA cells carrying an empty plasmid, and AsidA
AdidA cells carrying a plasmid with either sidA or didA driven by their native promoters were plated as in panel A.
(C-E) Synchronous populations of swarmer cells from the strains in panel A were placed on agarose pads containing
PYE and MMC and then imaged for 8 hours by time-lapse microscopy. (C) The time to first mid-cell division and
(D) the percentage of cells that stopped growing following division are shown (for criteria on calling division events
and growth cessation, see Supplemental Materials and Methods). Asterisks represent a statistically significant (p <
0.01) difference relative to the wild type. Error bars represent standard error of the mean. (E) Representative fields
of wild-type and AsidA AdidA swarmer cells grown on pads containing MMC are shown at the time points indicated
in hours. Black arrows indicate cells that divided. Red arrows indicate cells arrested for growth following division.
Bar, 2 pim.
To better understand the nature of MMC sensitivity in AdidA AsidA cells, we used timelapse microscopy to examine synchronous populations of swarmer cells during growth on
agarose pads containing MMC. Wild-type swarmer cells grown at this concentration of MMC
112
did not divide for -5 hours on average, which is significantly longer than the standard doubling
time of~90 minutes in MMC-free media (Fig. 3.11 C, 3.1 lE). Roughly 5% of wild-type cells
arrested growth following a cell division event (Fig. 3.11 D), indicating that division may have
been premature or inappropriately executed and consequently was lethal. The single deletion
strains, AsidA and AdidA, were also able to delay division in the presence of MMC, although
they divided slightly earlier than the wild type by -30 (p = 0.02) and ~15 (p = 0.34) minutes,
respectively. These single deletion strains also had twice as many growth arrested cells following
division events compared to the wild type, although these defects were apparently insufficient to
produce a gross viability defect (Fig. 3.11 A). In contrast, AdidA AsidA cells lacking both
inhibitors divided ~1.25 hours earlier than wild-type (p = 6.9 x 10-0), and more than 25% of
cells exhibited growth defects following a division event (Fig. 3.11 C-E). Taken together, our
data suggest that the lethality experienced by AdidA AsidA cells in the presence of MMC results
from an inability to appropriately delay cell division.
We also assessed the viability of AdidA AsidA cells on plates containing novobiocin or
cephalexin. Neither of these conditions directly damage DNA nor do they strongly induce sidA
or didA expression (data not shown). Accordingly, AdidA AsidA cells were not sensitive to
novobiocin and were only weakly sensitive to cephalexin suggesting that DidA and SidA do not
play significant roles during these stress responses (Fig. 3.9C). These findings are consistent with
our model in which the sensitivity offts W(A246T) cells during these same stresses results from
an abnormal propensity to divide and not a reduced affinity for DidA or SidA.
113
Discussion
SOS-independent regulationof the DNA damage response
During episodes of DNA damage, cells often use checkpoint systems to transiently inhibit
the cell cycle and prevent cell division (Hartwell and Weinert, 1989). In bacteria, the regulatory
paradigm for responding to DNA damage has long been the E. coli SOS system in which
cleavage of the repressor LexA drives the transcription of DNA repair genes as well as the cell
division inhibitor sulA (Huisman and D'Ari, 1981; Mukherjee et al., 1998; Opperman et al.,
1999). SOS-induced division inhibitors have subsequently been identified in a range of other
bacteria, including sulA homologs in y-proteobacteria and the unrelated genes yneA, divS, chiZ
and sidA in various other species (Chauhan et al., 2006; Dullaghan et al., 2002; Kawai et al.,
2003; Modell et al., 2011; Ogino et al., 2008). Although these SOS-dependent regulators are
often assumed to be the primary, or even sole, mechanism for inhibiting division post-damage,
there have been hints of SOS-independent division regulation. For instance, in E. coli, B. subtilis,
and Caulobacter,cells lacking their SOS-induced inhibitors, or that cannot induce an SOS
response, can still become filamentous following DNA damage indicating an alternative means
of blocking cell division (Hill et al., 1997; Howe and Mount, 1975; Huisman et al., 1980; JaffM et
al., 1986; Liu et al., 2001). However, to the best of our knowledge, no damage-induced, SOSindependent division regulators have been previously documented. Here, we identified didA in
Caulobacteras one such regulator.
How do Caulobactercells recognize and respond to DNA damage to induce didA if not
through the canonical derepression of SOS genes? One possibility is that cells monitor the same
signal, but with a different protein sensor. The SOS system measures the accumulation of
ssDNA, which stimulates RecA to trigger the autocatalytic cleavage of LexA (Craig and Roberts,
1980; Little and Mount, 1982; Slilaty et al., 1986). Another protein, such as the RecA homolog
RadA, could also recognize ssDNA, but ultimately affect the activity of a different transcription
114
factor. Alternatively, a DNA damage sensor unrelated to RecA could recognize a distinct type of
DNA damage or DNA structure. In B. subtilis, the diadenylate cyclase DisA monitors genome
integrity and may recognize branched DNA structures that arise during the recombination-based
repair of double-strand breaks (Witte et al., 2008). When paused at such DNA structures, DisA is
prevented from synthesizing cyclic-di-AMP (c-di-AMP), a diffusible molecule required for the
activation of the transcription factor SpoOA, thereby coupling DNA damage with transcription
(Bejerano-Sagie et al., 2006; Oppenheimer-Shaanan et al., 2011; Witte et al., 2008). It remains
unclear precisely how c-di-AMP affects SpoOA activity in B. subtilis and whether a c-di-AMPbased response to DNA damage extends to other organisms. Nonetheless, didA transcription
could follow a similar regulatory strategy that relies on c-di-AMP, or another damage-regulated
second messenger.
Another potential mechanism driving the SOS-independent induction of didA could involve
the conserved replication protein DnaA that also serves as a transcription factor. In B. subtilis,
replication stress somehow activates DnaA to transcriptionally regulate ~50 genes (Breier and
Grossman, 2009; Goranov et al., 2005). In E. coli, DnaA may also help respond to replication
stress by inhibiting cell division in an SOS-independent manner, but this pathway remains poorly
characterized (Fujimitsu et al., 2008; Gon et al., 2006). It is unclear whether the transcriptional
activity of CaulobacterDnaA is affected by DNA damage and it is unknown whether DnaA
regulates didA. Nonetheless, didA transcription could, in principle, be driven by DnaA or another
transcription factor that responds to replication stress and stalled forks, which often accompany
DNA damage.
The execution and regulation of cell division
Many cell division inhibitors, including E. coli SulA, block cell division by disrupting FtsZ
polymerization. FtsZ is an effective target as it recruits most other cell division proteins.
However, neither DidA nor SidA affect the assembly or stability of FtsZ rings in Caulobacter.
115
Instead, these inhibitors appear to block cell division by targeting FtsW, FtsI, and FtsN. Bacterial
two-hybrid studies indicated that DidA interacts with FtsN. Additionally, a chromosomally
encoded GFP-FtsN fusion was able to suppress the effects of DidA overproduction, supporting a
model in which DidA inhibits cell division by binding to FtsN. SidA interacts with FtsW and
FtsN in a two-hybrid system, and the lethality of overproducing SidA can be suppressed by
mutations in either FtsW or FtsI (Modell et al., 2011). Although DidA and SidA bind different
proteins, these two inhibitors may ultimately inhibit division in similar ways as two mutations in
fts W, and one inftsI, can suppress the effects of overproducing either SidA or DidA.
FtsW, FtsI and FtsN are among the last essential proteins recruited to the cytokinetic ring.
These proteins physically interact with each other and likely form a sub-complex within the
divisome that drives the synthesis and remodeling of the septal cell wall. FtsW translocates cell
wall precursors into the periplasm at mid-cell which are then incorporated into a mature septum
by transglycosylases and the transpeptidase domain of FtsI (Mohammadi et al., 2011; Tipper and
Strominger, 1965; Wise and Park, 1965). The function of FtsN is less clear, although in
Caulobacter,its essential activity is located within a periplasmic linker domain (Moll and
Thanbichler, 2009). In both Caulobacterand E. coli, FtsN also recruits proteins involved in cell
wall remodeling to the division site (Bernhardt and de Boer, 2003; Goley et al., 2010; Moll et al.,
2010; Peters et al., 2011; Poggio et al., 2010).
How do single mutations in FtsW and FtsI prevent the inhibition of cell division by both
SidA and DidA? One possibility is that these mutations reduce the affinities of SidA and DidA
for their division protein targets. However, SidA binding to FtsW was unaffected by the A246T
mutation and DidA binds FtsN, not FtsW or FtsI, at least in our bacterial two-hybrid system. A
second possibility is that SidA and DidA block the recruitment of even later arriving proteins. As
noted, FtsN may help recruit cell wall remodeling factors such as the peptidase DipM and the
peptidoglycan amidase AmiC (Bernhardt and de Boer, 2003; Moll et al., 2010). Although the
116
genes encoding such proteins are individually dispensible, it is formally possible that SidA and
DidA disrupt the recruitment of multiple peptidoglycan remodeling factors, thereby preventing
division. However, given that the inhibitory activity of both SidA and DidA can be suppressed
by mutations in FtsW and FtsI, this model seems unlikely.
Instead, we favor a model in which the FtsW/FtsI/FtsN subcomplex exists in two
conformations: an inactivate conformation that is stabilized by SidA or DidA, and an active
conformation which drives cytokinesis (Fig. 3.12A). We propose that the substitutions that
suppress both SidA and DidA, such as FtsW(A246T), may effectively lock FtsW/FtsI/FtsN in the
active state allowing cells to bypass the block in division normally caused by an accumulation of
these inhibitors. On their own, these suppressor mutations cause cells to initiate division
hyperactively. In support of this model, cells with the suppressing mutations were reproducibly
shorter than wild-type cells (Fig. 3.9A-B), likely because they divide at a slightly earlier stage of
the cell cycle. Additionally, FtsW(A246T) cells are sensitive to novobiocin and cephalexin,
treatments that do not cause DNA damage but nonetheless require cells to delay division.
A
B
outer
mem brane
DNA damage
SOS-dependent
(LexA, RecA)
SIdA
SOS-independent
peptidoglycan
DidA
inner
membrane
Ftsw SidA
FtsW/IIN
}Dd
Ftaz
FtsW/IIN*
LexA
cell division
DNA damage
didA
117
Figure 3.12. Parallel pathways regulate cell division in Caulobacter following DNA damage. (A-B) Two cell
division inhibitors are induced following DNA damage in Caulobacter.sidA is induced by cleavage of the SOS
repressor LexA while didA is induced by unknown regulatory factors. SidA and DidA are both small transmembrane
proteins that can block cell division by preventing the divisome sub-complex FtsW/I/N from assuming an active
conformation, FtsW/I/N*. FtsW/I/N* could promote division by enhancing peptidoglycan synthesis and remodeling,
by triggering FtsZ constriction, or by coordinating these activities.
Taken together, our results suggest that the DNA damage-induced division inhibitors in
Caulobactertarget the FtsW/FtsI/FtsN subcomplex to block constriction of the division
machinery and cell envelope. Precisely how SidA and DidA block division is not yet clear, in
part because the execution of cytokinesis remains poorly characterized at a molecular level. The
synthesis of septal cell wall material could provide the force and directionality for cellular
constriction, with FtsZ required mainly for mid-cell positioning of division proteins. In such a
case, SidA and DidA could prevent division by directly blocking a critical or rate-limiting
peptidoglycan modifying activity of the FtsW/FtsI/FtsN subcomplex. Alternatively, GTP
hydrolysis by the FtsZ ring may provide the energy for, and directionality of, constriction,
effectively pulling the rest of the cytokinetic ring along with it (Li et al., 2007). Assembly or
activity of the FtsW/FtsI/FtsN sub-complex could somehow trigger FtsZ constriction, and the
inhibitors SidA and DidA may block this step of division. Finally, it is possible that Z-ring
constriction and septum synthesis combine to drive cytokinesis. As FtsW, FtsI, and FtsN are
transmembrane proteins with cytoplasmic and periplasmic domains, they are each well
positioned to coordinate the Z-ring and nascent septum, and SidA and DidA could disrupt this
coordination. Distinguishing between these various models for cytokinesis and elucidating the
precise mechanisms of action for SidA and DidA will ultimately require more detailed studies of
the FtsW/I/N sub-complex; the mutants identified here, such as FtsW(A246T), may prove
particularly useful in these efforts.
118
FinalPerspectives
Our results highlight the FtsW/Ftsl/FtsN subcomplex as an important regulatory node in the
control of cell division. Following certain types of DNA damage, DidA and SidA appear to
function together to prevent inappropriate cell divisions (Fig. 3.12A-B). Such redundancy may
afford cells with a fail-safe survival mechanism. Alternatively, or perhaps in addition, SidA and
DidA may be differentially induced following different types of DNA damage, providing
independent routes to the inhibition of cell division under different conditions. Additionally, we
note that although cells lacking both sidA and didA divide prematurely during DNA damage,
many still filament to some degree, suggesting that yet other mechanisms of division inhibition
exist in Caulobacter.Finally, DidA is the latest in a growing class of small, stress-induced
membrane proteins that play critical regulatory roles (Fontaine et al., 2011; Hobbs et al., 2011).
These proteins are often missed or incorrectly annotated in genome sequences, but many, like
SidA and DidA, clearly play critical roles in regulating cellular processes, including cell division.
119
Materials and Methods
Strains, plasmids, and growth conditions
Strains and plasmids used in this study are listed in Table S2 in the submitted manuscript
with construction details and growth conditions provided in the Supplemental Materials and
Methods.
DNA microarrays
RNA expression profiling was done as described (Biondi et al., 2006). Expression
experiments were performed in duplicate and the results for each gene were averaged.
Immunoblots and biochemical fractionations
Samples for immunoblots were normalized in sample buffer to 0.5 OD600 /50 PL, resolved on
12% sodium dodecyl sulfate-polyacrylamide gels and transferred to polyvinylidene difluoride
transfer membrane (Pierce). Membranes were probed with polyclonal rabbit a-CtrA and a-GFP
(Invitrogen) at a 1:5000 dilution and monoclonal mouse a-FLAG (Sigma) at a 1:3000 dilution.
Secondary HRP-conjugated a-rabbit (Pierce) or a-mouse (Pierce) were used at a 1:5000 dilution.
Biochemical fractionation was performed as described (Modell et al., 2011).
Microscopy
All phase contrast images were acquired on a Zeiss Observer ZI microscope with a 100x/1.4
oil immersion objective and an LED-based Colibri illumination system. For additional
information on image analysis and time-lapse microscopy, see the Supplemental Materials and
Methods.
Bacterial two-hybrid analysis
Two-hybrid complementation assays were performed essentially as described (Karimova et
al., 2005). BTH101 cells harboring plasmids with the T25 and T18 fusion constructs were grown
120
to single colonies on LB agar plates and restruck or spotted on MacConkey agar plates
supplemented with maltose for imaging.
Identification of DidA overproduction suppressors
Wild-type cells were transformed with a Pvan.3xF-didA overproduction plasmid and plated
on PYE agar in absence of vanillate to allow colony formation. Single colonies were grown
overnight in PYE and plated on PYE agar supplemented with vanillate at roughly 2x 106 colony
forming units per 10 cm plate. Rare colonies were grown overnight in PYE supplemented with
vanillate and samples were taken for immunoblots, plasmid preparations and archival. To isolate
chromosomal suppressor mutations and eliminate mutations arising in the 3xF-didA
overproduction plasmid, we screened for colonies that met two criteria. (1) We used
immunoblotting to check that 3xF-DidA production in each suppressor strain was similar to that
seen in wild-type cells transformed with the same plasmid and grown in vanillate for 1.5 h. (2)
Plasmids from the suppressor strains were transformed into wild-type cells and plated on PYE
agar supplemented with or without vanillate. The presence of thousands of colonies on plain
plates and few colonies on vanillate indicated a functional plasmid. The mutation in the
fts W(A246T) suppressor was identified by whole-genome sequencing.
121
Acknowledgements
We thank C. Tsokos, C. Aakre and A. Podgornaia for their comments on the manuscript. We
thank C. Aakre for assistance in overlaying cell boundaries on fluorescent images.
Financial Disclosure
M.T.L. is an Early Career Scientist at the Howard Hughes Medical Institute. This work was
supported by a National Institutes of Health grant (RO 1 GM082899) to M.T.L and an NSF
Graduate Fellowship to J.W.M.
122
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125
Chapter 4
Conclusions and future directions
126
Conclusions
In this work, I identified and characterized two regulators that together allow Caulobacterto
delay cell division following DNA damage (Fig. 3.12). sidA is an SOS-induced division inhibitor
and functional analog of sulA in E. coli and yneA in B. subtilis. Given that a sidA deletion
significantly reduced filamentation in a AlexA background, which causes constitutive SOS
induction, we suspect that sidA is the primary, if not only division inhibitor within the SOS
regulon. didA also accumulated following DNA damage, but its induction occurred even in
strains lacking recA or harboring a non-cleavable allele of lexA, suggesting that its regulation is
SOS-independent. Strains lacking either inhibitor still typically delayed division and remained
viable during exposure to Mitomycin C (MMC). However, strains lacking both inhibitors divided
prematurely and suffered a viability defect under the same conditions. These results illustrate
some degree of redundancy within the CaulobacterDNA damage response and underscore the
presence and importance of an SOS-independent transcriptional response to DNA damage.
Once induced, SidA and DidA are both inserted into the inner membrane at mid-cell where
they prevent division. Unlike SulA which completely dissolves Z-rings or the Gram-positive
SOS-induced inhibitors which have mild effects on Z-ring assembly, SidA and DidA did not
noticeably disrupt Z-ring formation nor any subsequent step in divisome assembly. Instead I
provided evidence that each inhibitor disrupts divisome constriction by targeting components of
the late-arriving divisome sub-complex FtsW/FtsI/FtsN. By a bacterial two-hybrid analysis, I
showed that SidA bound strongly to FtsW and secondarily to FtsN while DidA bound
exclusively to FtsN. Furthermore, thefts W(T180A) mutation and an N-terminal GFP fusion to
FtsN selectively suppressed the activities of SidA and DidA respectively, likely by reducing the
affinity of each inhibitor for their target. To our knowledge, these are the first examples of
endogenous, regulatory division inhibitors with known targets other than FtsZ.
127
Strikingly, several mutations infts W andftsl suppressed the effects of both inhibitors with
fts W(F145L) andfts W(A246T) demonstrating the most complete suppression. An explanation of
reduced affinity is unlikely for these suppressors given that by our bacterial two-hybrid analysis,
SidA bound FtsW(A246T) as strongly as wild-type FtsW and DidA did not bind FtsW. Instead,
we hypothesize that these suppressors cause cells to divide hyperactively. In support of this
model, cells harboring suppressor mutations divided prematurely in the absence of DNA damage
resulting in a small cell phenotype. Importantly,fts W(A246T) cells were also sensitive to several
additional stresses which likely require division inhibition but do not cause SidA or DidA
induction. Taken together, our results highlight FtsW/FtsI/FtsN as a critical regulatory node in
Caulobacterand suggest a new role for the sub-complex in triggering divisome constriction.
New Roles for FtsW/FtsI/FtsN in cytokinesis
What are the activities of FtsW and FtsN that are inhibited by SidA and DidA respectively?
In E. coli, FtsW forms a complex with FtsI in the absence of other divisome components
(Fraipont et al., 2011), and it is required for the proper recruitment of FtsI (Mercer and Weiss,
2002) and FtsN (Addinall et al., 1997). In Caulobacter,although FtsW may not be strictly
required for the recruitment of FtsI or FtsN (Goley et al., 2011), all three proteins interact with
each other and likely form a sub-complex within the divisome. The essential function of FtsN is
unknown, but it may help to stabilize the assembled divisome (Goehring et al., 2007; Rico et al.,
2010). I showed that during SidA or DidA overproduction, FtsZ, FtsW, FtsI and FtsN still
localize to nascent division sites indicating that the assembly, recruitment and stabilization
activities of FtsW and FtsN are unaffected. A second proposed function of FtsW is to translocate
the cell wall precursor lipid II from the cytoplasm into the periplasm at mid-cell where it is
incorporated into the invaginating septum. Biochemical evidence for this activity was recently
provided as purified FtsW allowed the translocation of lipid II in artificially constructed singlemembrane liposomes (Mohammadi et al., 2011). Using a fluorescently-labeled vancomycin
reporter, I showed that SidA does not prevent the septal translocation of cell wall precursors into
128
the periplasm and thus does not inhibit the translocase activity of FtsW. Given that SidA and
DidA do not disrupt any known function of FtsW or FtsN in divisome assembly or septum
synthesis, we propose that these components likely play previously unappreciated roles in
promoting constriction of the assembled divisome.
The identification of mutations infts W andftsI that suppressed the effects of both inhibitors
indicated that despite having distinct binding partners, SidA and DidA could prevent division by
inhibiting the same general divisome function. Furthermore, our data suggest that these
suppressor mutations do not simply decrease inhibitor affinity, but instead cause divisomes to
execute this function hyperatively. We have proposed a unified model whereby this unknown,
critical activity could be considered as a single output performed by the FtsW/Ftsl/FtsN
complex. Accordingly, the complex could exist in one of two conformations: an inactive
conformation which is stabilized by the presence of SidA or DidA and an active conformation
which is stabilized by the suppressor mutations infts W andftsl. Understanding the precise nature
of this divisome function should provide mechanistic insights into the molecular events that
mediate cytokinesis and help to distinguish between the proposed models of divisome
constriction.
Characterizing the FtsW/FtsI/FtsN activity pathway
In the cell wall-centric model of division, septum synthesizing proteins provide the force
and directionality for divisome constriction and cytokinesis. Intriguingly, hyperactive suppressor
mutations were found inftsl, which encodes the only essential divisome component with a wellestablished catalytic activity in septum synthesis. The transpeptidase domain of FtsI introduces
crosslinks into the peptide chains that project from the glycan backbone, resulting in mature
septal cell wall (Adam et al., 1997). Accordingly, SidA and DidA could inhibit this
transpeptidase activity through the intermediates FtsW and/or FtsN. Alternatively, the inhibitors
could prevent FtsW, FtsI or FtsN from activating one or more septal transglycosylases or
129
amidases or from performing yet other functions that are required for synthesizing and shaping
the septal cell wall. Several currently available techniques might afford a more complete
assessment of peptidoglycan dynamics during SidA and DidA overproduction. Pulse chase
experiments with D-Cysteine, which is readily incorporated into peptidoglycan peptide chains,
have allowed new areas of cell wall growth to be monitored with high temporal resolution
(Aaron et al., 2007). The fluorescent penicillin derivative Bocillin-FL has been used to directly
assess of FtsI activity (Eberhardt et al., 2003). Another peptidoglycan labeling technique, using
fluorescently tagged tripeptides, can be potentially used to determine regions of amidase activity
(Olrichs et al., 2011). Each of these techniques monitor the synthesis of septal cell wall at
different stages and might lend support to cell wall-centric regulation by SidA, DidA and the
suppressor mutations.
In the FtsZ-centric model of division, FtsZ and perhaps FtsA drive divisome constriction
and guide the inward synthesis of the septal cell wall. Under this model, SidA and DidA could
prevent FtsW/FtsI/FtsN from triggering a divisome assembly checkpoint in which the latearriving sub-complex could somehow enable FtsZ constriction. The FtsQ/FtsL/FtsB complex
forms contacts with both early- and late-arriving divisome components and is thus an attractive
candidate for mediating such a checkpoint (Di Lallo et al., 2003; Karimova et al., 2005).
Interestingly, interactions within the divisome occur in part through contacts between the
transmembrane domains (van den Berg van Saparoea et al., 2013), and most of the SidA and
DidA suppressor mutations mapped within the transmembrane domains of FtsW and FtsI.
Alternatively, FtsW/FtsI/FtsN could directly trigger Z-ring constriction and each component has
demonstrated direct binding to FtsZ or FtsA in various organisms (Busiek et al., 2012; Datta et
al., 2002; Datta et al., 2006; Di Lallo et al., 2003; Karimova et al., 2005), although evidence for
such interactions in Caulobacteris lacking. Few techniques exist to experimentally assess FtsZ
dynamics in vivo. Fluorescence recovery after photobleaching with a fluorescently-labeled copy
of FtsZ could allow Z-ring turnover to be examined during SidA and DidA overproduction.
130
However, it would be difficult to determine whether any observed effects were due to direct
regulation on FtsZ or to an indirect consequence of inhibiting any step in cell wall synthesis.
The identification of additional, hyperactive suppressors of SidA and DidA could help to
distinguish between the models of division. During our suppressor screens, I isolated only one
suppressor twice (fts W(T180A)) and thus have not screened to saturation. To more readily
identify additional mutations that suppress both inhibitors, I have recently developed a plasmid
which overproduces SidA from the P y/1 promoter and DidA from the Pvan promoter. Colonies
allowing growth under one inducing condition can be restruck on the other condition providing a
fast and easy secondary screen for dual suppression. Using this method, I have already recovered
one of the original, hyperactive suppressor mutations (fts W(F]45L)). If additional suppressors
are found in the transpeptidase domain of FtsI or the periplasmic domains of known or novel
divisome components, the cell wall-centric model of division and its inhibition would be
supported. Alternatively, suppressors within FtsZ, FtsA or other cytoplasmic FtsZ binding
proteins might support the FtsZ-centric model. Ultimately, such a screening strategy would more
thoroughly clarify which genetic determinants, both in terms of proteins and domains, constitute
the FtsW/FtsI/FtsN activity pathway.
A complementary approach to investigate the downstream effectors of FtsW/FtsI/FtsN
activity would be to expand upon our bacterial two-hybrid (BACTH) analysis. The suppressor
mutations infts W andftsI could expose or enhance binding sites within the FtsW/FtsI/FtsN
complex for other divisome components. SidA and DidA could then prevent division by
obstructing these binding sites. To detect such interactions, the FtsW/FtsI/FtsN complex, with or
without suppressor mutations, could be co-expressed and tested for interactions with each
divisome component. Similarly, a three-hybrid approach could be used to determine whether
SidA or DidA could disrupt any of these interactions. As with the saturating suppressor screen,
the identities of these interactions could reveal which additional divisome components, if any,
131
participate in the SidA/DidA signaling pathway. A major caveat with these experiments is that it
may not be possible to reconstitute activities of Caulobacterdivisome components within the
context of an F. coli host expressing its own divisome. It is also possible that additional
components of the Caulobacterdivisome would have to be co-expressed with FtsW/FtsI/FtsN in
order to reveal yet other interactions or achieve proper membrane insertion or orientation.
Cryo-election microscopy (Cryo-EM) would allow the direct visualization of the inner
membrane and septum during SidA and DidA overproduction. Abnormalities in either of these
layers could indicate a mechanism of action, although if both layers are similarly prevented from
constricting, it would be difficult to determine which one was targeted for inhibition. A more
informative Cryo-EM experiment would be one in which movements of the inner membrane and
septum were uncoupled. For instance, I observed that strains harboring the hyperactive
ftsW(A246T) mutation are sensitive to the FtsI transpeptidase inhibitor cephalexin (Fig). Cells
treated with cephalexin cannot insert crosslinks into new septal cell wall and thus likely cannot
support the growth of an invaginating septum. If FtsW(A246T) hyperactivity causes premature
constriction of the Z-ring, the inner membrane could become uncoupled from growth of the
cephalexin-inhibited septum. This result would potentially explain the viability defect of
fts W(A246T) cells treated with cephalexin and lend support to the model of a FtsW/Ftsl/FtsN
assembly checkpoint regulating a FtsZ-centric division event.
A puzzling aspect of our research is that suppressor mutations are found in divisome
components that do not appear to be directly targeted by either inhibitor; mutations inftsI
suppress SidA overproduction, and mutations infts W andftsI suppress DidA overproduction.
However, given the close association of each subunit within the FtsW/FtsI/FtsN complex with
the other two, it is not difficult to envision scenarios that explain suppressor activities and modes
of inhibition (Fig. 4.1). If we consider FtsW, FtsI and FtsN as individual components of a linear
activation pathway, an epistasis analysis would place FtsW and FtsI downstream from FtsN and
132
its inhibition by DidA. Similarly, FtsI would lie downstream from FtsW and its inhibition by
SidA with the final pathway appearing FtsN > FtsW > Ftsl. Alternatively, the FtsW/FtsI/FtsN
complex could harbor a single active site composed of features from each subunit. The binding
of SidA and DidA at distinct, possibly allosteric sites could ultimately inhibit the activity of a
single active site. Structural studies of FtsW/FtslI/FtsN, in complex with SidA or DidA, would
help to distinguish between these models. While it is still difficult to perform crystallographic or
structural NMR studies on membrane proteins, several recent technological and analytical
advances could facilitate such research (Ubarretxena-Belandia and Stokes, 2012).
DidA -- I FtsN
SidA
--
FtsW
rftsw*
ftsW
Fts I
Fts I
FtsW
FtsN
16 tsI*
SidA
constriction
--
DidA
constriction
Figure 4.1. Models of FtsW/FtsI/FtsN and SidA/DidA activity. On the left, FtsW, FtsI and FtsN are modeled as a
linear activation pathway. fts W* andftsl* denote hyperactive suppressor mutations. On the right, FtsW, FtsI and
FtsN are modeled as a complex with a singular function.
Why target FtsW/FtsI/FtsN?
Before the identification of SidA and DidA, it was unclear whether the targeting of FtsZ by
SulA in E. coli was representative of damage-induced cell division inhibitors in Gram-negative
bacteria. The Gram-positive SOS-induced division inhibitors, with transmembrane domains and
peptidoglycan-binding motifs, seemed to represent a mechanistically distinct group acting
downstream from Z-ring assembly. As mentioned, it is perhaps logical that Gram-positive
133
bacteria would regulate later steps in division given the presence of cell walls that are
considerably thicker and more complex that their Gram-negative counterparts. For these reasons,
it was surprising to find that two damage-induced division inhibitors in Caulobacter,a Gramnegative proteobacterium like E. coli, target divisome components implicated in cell wall
synthesis and remodeling, and thus may bear more similarity to the Gram-positive inhibitors.
Why do Caulobacterand E. coli employ different regulatory strategies to execute the same
task within a similar, rod-shaped envelope? One possibility is that despite their structural
similarities, the two proteobacteria execute division differently, and thus require different forms
of regulation. In Caulobacter,SidA and DidA suppressor mutations in fts W andftsl exhibit small
cell phenotypes indicating that these divisome components perform rate-limiting steps in
division. The concept of a rate-limiting step is difficult to imagine when considering a multifunctional division machine that is itself poorly understood. In strict temporal terms, the ratelimiting step could control the onset of division and dictate its timing within the cell cycle, or it
could determine the rate at which constriction proceeds once division is triggered. In biochemical
terms, there could be a rate limiting enzymatic step within a linear biosynthetic pathway, for
instance the translocation of cell wall precursors could be rate-limiting for the synthesis of the
septum. Alternatively, a rate-limiting step could serve as a checkpoint that activates several
downstream functions in parallel. Energetically, rate-limiting and force-generating steps are
frequently related (Brenner and Eisenberg, 1986; Smith et al., 2001), although this is by no
means a theoretical requirement. Whichever model or combination thereof is most correct, small
cell alleles have been observed in E. coli, with gain-of-function mutations in the FtsZ-associated
membrane tether FtsA (Geissler et al., 2007), and in B. subtilis, with N-terminal truncations in
FtsL (Bramkamp et al., 2006). Intriguingly, most known regulators of division in E. coli target
the Z-ring while in B. subtilis, FtsL has been implicated in mediating division inhibition in
response to both YneA and replication stress (Goranov et al., 2005; Kawai and Ogasawara,
2006). Much additional research will be required to precisely define rate-limiting steps as they
134
pertain to bacterial cell division. However, it is generally possible that the most critical steps in
division are performed by different divisome components or complexes in different species.
Although Caulobacterand the Gram-positives may have converged upon the inhibition of
septum synthesis following DNA damage, under the aforementioned divisome assembly
checkpoint model, SidA and DidA could instead target FtsW/FtsI/FtsN activities relating to Zring constriction.
In addition to the properties of the divisome itself, several lifecycle and evolutionary
distinctions could explain the targeting of different divisome components between the a- and yproteobacteria. In Caulobacter,the Z-ring is assembled in stalked cells, a cell cycle stage when
the nucleoid still occupies the mid-cell (Kelly et al., 1998). Z-ring formation at a similar stage is
prevented in E. coli by the nucleoid-associated division inhibitor SlmA (Bernhardt and de Boer,
2005). Therefore, a mixed population of Caulobactercells contains a higher percentage of Zrings compared to E. coli, and the inhibition of a later stage in division might be less temporally
and energetically costly than the complete dis-assembly and reassembly of Z-rings and
divisomes. Alternatively, the choice of a divisome target could reflect the type of DNA damage
most frequently encountered by that organism. For instance, if Caulobacterevolved during
transient episodes of minimal DNA damage, it might be desirable to momentarily prevent
divisome activity while leaving it fully assembled and ready to divide once the damage is
removed. Alternatively, the inhibition of Z-rings in E. coli could reflect a history of more severe
DNA damaging agents, during which the complete disassembly of Z-rings more robustly
prevented any chance of mistakenly attempting division.
A deeper understanding of these evolutionary questions will require the identification and
characterization of additional cell division inhibitors in other bacterial clades. Unfortunately, if
the Caulobacterinhibitors are any indication, such genes are not always easy to find. Unlike
SulA and the Gram-positive inhibitors, SidA and DidA are representative of a growing class of
135
small proteins which have been understudied as a result of incomplete or inaccurate genome
annotations (reviewed in (Hobbs et al., 2011)). Indeed, both genes were mis-annotated in
Caulobacter,and their homologs in other a-proteobacteria were often completely unannotated.
Directed informatic and biochemical approaches are proving useful in the search for small
proteins, and intriguingly, many in E. coli are single-pass transmembrane proteins that play
important regulatory roles (Fontaine et al., 2011; Hemm et al., 2010; Hemm et al., 2008). As the
repertoire of damage-induced inhibitors grows, it may be informative to determine the activity of
each in heterologous species and the consequences of substituting one for another. If some
inhibitors are broadly effective, it could be telling to determine consequences of substituting one
for another. This type of experimental strategy could be used to support evolutionary and
functional hypotheses regarding the selection of regulatory targets within the divisome.
The SOS-independent regulation of didA
The SOS system has long served as the lone example of a bacterial regulatory response to
DNA damage. However, the recent discovery of the di-adenylate cyclase DisA in B. subtilis
demonstrated that bacteria can in fact respond to damage with multiple recognition elements and
signaling pathways (Bejerano-Sagie et al., 2006). Furthermore, the modulation of the
transcriptional activities of DnaA in response to replication stress indicates that in B. subtilis,
several overlapping modes of regulation can address the downstream consequences of DNA
damage in addition to the lesions themselves (Goranov et al., 2005). In E. coli, many anecdotal
reports have suggested the existence of SOS-independent responses that help to prevent cell
division following damage, although mechanistic insights are lacking. I identified didA, which is
to our knowledge the first SOS-independent, damage-induced cell division inhibitor.
How is didA regulated if not by the SOS regulators RecA and LexA? I initially observed
DidA induction following exposure to the DNA damaging agent mitomycin C, an alkylating
agent that causes a variety of lesions arising from single-strand adducts, double-strand crosslinks
136
and the intermediate structures resulting from the repair of each lesion. DidA induction could
therefore be triggered by any of these structures or by indirect consequences of an MMC
treatment, such as replicative or oxidative stress. I am currently investigating the mechanisms of
DidA regulation, and our preliminary results have uncovered genetic networks involved in the
transcriptional induction of DidA and the stationary phase-specific induction of a phage-like
system that likely mediates horizontal gene transfer and induces DidA indirectly.
DriD is required for DidA induction
To better understand the signals responsible for DidA induction, I monitored a DidA
reporter following a variety of stresses, including DNA damaging agents, replication inhibitors,
antibiotics and other physical stressors. Strikingly, DidA was induced exclusively by DNA
damaging agents suggesting that its SOS-independent signal is likely derived from a damaged
DNA structure (Fig 4.2). I next sought to determine whether DidA was induced by the SOS
signal (ssDNA), albeit with different regulatory machinery, or by a distinct damaged structure.
DidA and SidA reporter strains were treated with a panel of DNA damaging agents, each with a
MMC
HU
HU
(0)
(hi)
ceph
nov
(10)
nov
(hi)
UV
(10)
(min in M2)
UV
M2G
(hi)
30
60
120
EtOH
(1o)
DIdA-3xF
-
MMC
EtOH
NaCI
(hi)
(10)
H2 02
(10)
H202
(hi)
Kan
Tet
Chlor
DidA-3xF
137
Figure 4.2. didA is induced specifically by DNA damaging agents. Western blot of a strain expressing didA from its
native promoter tagged at its C-terminus with the 3xFLAG epitope treated with mitomycin C (MMC), hydroxyurea
(HU), cephalexin (ceph), novobiocin (nov), ultraviolet light (UV), ethanol (EtOH), high salt (NaCl), hydrogen
peroxide (H202), kanamycin (kan), oxytetracycline (tet), and chloramphenicol (chlor). Where indicated, cells were
treated at high (hi) and low (lo) doses, and blots were probed with a-FLAG (Modell and Laub, unpublished).
different mechanism of action. Indeed, DidA and SidA showed differential induction levels with
maximal DidA induction occurring following exposure to zeocin which causes double-strand
breaks (Fig. 4.3). Within the eukaryotic DNA damage response, there are proteins that
specifically recognize double-strand breaks, and it is tempting to speculate that DidA could
participate in a similar network in Caulobacter.
Norfloxacin
Hydroxyurea
Mitomycin C
(DNA alkylator) (Cyrase inhibitor) (stalled forks)
-
low
high
low
high
low
P51 d-gfp
PdA-gfp
high
Zeocin
(ds breaks)
low
high
*9
4
4GFP
Figure 4.3. sidA and didA are differentially regulated. Western blots of strains expressing gfp from the sidA or didA
promoter treated with the DNA damaging agents as indicated at low and high doses were probed with a-GFP
(Modell and Laub, unpublished).
To identify the genes responsible for the transcriptional induction of didA, I constructed a
strain in which the promoter of didA is fused to the chromogenic reporter LacZ (Fig. 4.4). When
cells harboring this construct were plated in the presence of MMC at a low concentration,
colonies still formed slowly and appeared blue owing to the MMC-induced expression of the
DidA reporter. I next mutagenized this strain with the Tn5 transposon and isolated colonies that
were unable to induce DidA and thus appeared white. Such strains contained transposon
insertions in either the PdidA-lacZreporter or an un-annotated transcription factor which we
named driD. I constructed a clean, in-frame deletion of driD and confirmed that DidA was no
longer induced following an MMC treatment (Fig. 4.5A). I additionally observed that DriD
protein levels remain unchanged during DidA induction suggesting that its activity is regulated
post-translationally following DNA damage (Fig. 4.5B).
138
IacZ
repressors
inducers
0
0
0
0
o0o
00 0
000
0
0
*00
0
00
0 00
0
.0.0
0
00
no MMC
0
MMC
Figure 4.4. Screening for transcriptional regulators of didA. A strain expressing lacZ from the didA promoter was
used to screen for didA regulators. Repressors
MM are identified
+ as blue+colonies on plates containing no MMC while
inducers are identified as white colonies on plates containing MMC (Modell and Laub, unpublished).
A
AdriD
WTf
M
B
MCDidA-3xF
3xF-driD
MMC:
B
-
+
driD-3xF
+
Ox*-dofD
dr4D-rxF
4
4
DidA-3xF
-4 DidA-3xF
Figure 4.5. DriD is activated post-translationally to induce didA. (A) Western blot of wild-type and AdriD cells
expressing didA-3xF from its native locus. (B) Western blot of cells expressing 3xF-driD or driD-3xFas the only
copy of driD and didA-3xF from its native promoter. Cells in A and B were treated with MMC for 1 hour. Blots
were probed with cc-FLAG. (Modell and Laub, unpublished).
With the identification of driD, we have begun to unravel the genetic network responsible
for DidA induction, but many questions remain. Does DriD directly participate in didA
transcription, or does it recognize a damaged DNA structure? Are double-strand break ends
responsible for DriD activation or is there another damaged structure that is enriched during a
zeocin treatment? How is DriD activated post-translationally and what genes other than DidA are
contained within its regulon? ChIP-PCR and ChIP-seq, in conjunction with a site-specific
double-strand break generation system recently developed by our lab, will allow the
139
characterization of DriD binding sites within the genome, whether they are damaged sites,
promoters or both. If DriD indeed recognizes DNA damage directly, gel-shift assays with an
assortment of DNA structures should help to home in on the specific DNA damage signal.
Finally, microarrays and additional genetic screens will uncover other components of the DriD
regulatory network and perhaps shed light on the mechanisms of its post-translational activation.
The identification of a gene transfer agent (GTA)
Using the same PdidA-lacZ reporter, I performed a screen for transcriptional repressors of
didA (Fig. 4.4). Cells harboring this construct form white colonies when plated in the absence of
MMC. We conjectured that just as AlexA cells constitutively transcribe SOS genes in the absence
of DNA damage, transposon insertions in a didA repressor should constitutively transcribe the
lacZ reporter resulting in blue colonies. I isolated blue colonies and mapped the transposon
insertions sites, many of which were within genes participating in DNA synthesis and repair
which likely increase basal levels of DNA damage. However, several insertions mapped to an
un-annotated transcription factor which we named sprD. Surprisingly, cells lacking sprD only
showed didA induction when grown into late stationary phase, whereas wild-type cells showed
no detectable didA induction in any growth phase (Fig. 4.6A). I next tagged SprD with the
3xFLAG epitope and found that SprD is expressed at very low levels during exponential phase
but then accumulates significantly at higher ODs consistent with its ability to repress didA during
stationary phase in wild-type cells (Fig. 4.6B). To better understand the role of SprD, I
performed a microarray analysis comparing AsprD and wild-type cells grown into stationary
phase. Strikingly, JsprD cells showed the dramatic induction of a ~15 gene operon that bears
homology to a locus encoding a GTA in the a-proteobacterium R. capsulatus (Leung et al.,
2010). GTAs are phage-like particles with capsids that are too small to carry their own genomes
(Fig. 4.7; reviewed in (Lang et al., 2012)). Instead they are thought to incorporate random
fragments of the host genome into their capsids and then mediate horizontal gene transfer events
between the same or closely related species. We believe this genome fragmentation step, which
140
likely produces double-strand breaks, generates the signal causing didA induction in stationary
phase cells. In support of this model, cells lacking the GTA locus no longer significantly induce
the didA reporter construct when grown into late stationary phase (Fig. 4.8A). Furthermore, cells
lacking both sprD and driD no longer induce didA during stationary phase (Fig. 4.8B), indicating
that the DriD regulatory machinery lies downstream from the GTA induced by the sprD deletion.
A
AsprD
WT
OD:
0.3
0.6
0.9
1.2
0.3
0.6
0.9
1.2
4
LacZ
P-idacZ
B
OD:
0.3
0.8
1.1
1.2
"4 SprD-3xF
Figure 4.6. SprD represses didA during stationary phase. (A) Wild-type and AsprD cells expressing lacZ from the
didA promoter were grown from mid-exponential phase into stationary phase. (B) Cells expressing sprD-3xFas the
only copy of sprD were grown from mid-exponential into stationary phase. Samples were taken at the ODs indicated
for western blot and probed with a-LacZ (A) and ot-FLAG (B) (Modell and Laub, unpublished).
donor cell
recipientcell
'I
IN
IL
IVP - GTA genes
(
I
/
4,
'I
IL
k
(
HGT event
4-
I
Figure 4.7. Gene transfer agents (GTAs). The red chromosomal region denotes the GTA genome. When induced,
most GTA particles incorporate random fragments of the host genome (blue). Occasionally, a GTA particle will
incorporate a fragment of its own genome (red), but the capsid head is too small to fit the GTA genome in its
entirety. GTAs can inject their contents into recipient cells where they can be incorporated into the host genome
(green) by recombination (adapted from (Lang et al., 2012)).
141
A
B
WT AGTA
AsprD/AdriD
AsprD
AsprD
-
OD:
0.8
1.0
1.1
0.8
1.0
1.1
4
DidA-3xF
Pdd-acZ
Figure 4.8. GTA production in stationary phase induces didA. (A) In a AsprD, Pdd4-didA-3xFbackground, cells
harboring or lacking the GTA genomic region were grown into stationary phase. Samples were taken for western
blot and probed with a-FLAG. (B) In a AsprD, PdidA-lacZ background, cells harboring or lacking driD were grown
into stationary phase. Samples were taken at the indicated ODs for western blot and probed with a-LacZ (Modell
and Laub, unpublished).
As with driD, many questions remain regarding sprD and regulation of the GTA locus.
Several GTAs have been identified in other a-proteobacteria which are no longer capable
performing gene transfer (Lang et al., 2012). I am currently attempting to transfer antibiotic
resistance mutations and gene cassettes from AsprD donors which overexpress the GTA locus to
determine if the CaulobacterGTA has retained its function. I am also individually mutating the
genes within the GTA locus to determine which is responsible for generating the signal involved
in didA induction. Finally, our work with SprD has generated a number of new questions
regarding stationary phase in Caulobacter.Given that cells lacking sprD only induce didA during
late stationary phase, what is the stationary phase-specific signal responsible for the induction of
the GTA and didA? GTAs in other species can be regulated by quorum-sensing, although such a
system has not been found in Caulobacter.It is also unclear whether there are conditions that
allow GTA induction during stationary phase in cells with functional SprD. I am currently
devising genetic screens that will allow the dissection of the stationary phase pathways that
control SprD activity.
Concluding Remarks
The identification and characterization of sidA and didA have highlighted considerable
diversity within the regulation of bacterial division. One of the best ways to understand a system
is to perturb it, and through our work on division inhibitors, we have developed tools and ideas
142
that should help clarify the molecular mechanisms of division. Our genetic screens have opened
doors in novel and unexpected areas of Caulobacterbiology, and future work on the SOSindependent transcriptional response to DNA damage and the regulation of a gene transfer agent
will further illuminate the richness of regulation that constitutes the bacterial DNA damage
response.
143
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