Exopolysaccharide Synthesis in Escherichia coli in Response
to Osmotic Stress
Thesis submitted for the degree of
"Doctor of Philosophy"
By
Michael Ionescu
Submitted to the Senate of the Hebrew University
June 2008
This work was carried out under the supervision of
Prof. Shimshon Belkin
ACKNOWLEDGEMENTS
A very special thanks go out to Dr. Yael Rozen, who initiated this project and guided me
throughout my first steps in science. Others who were actually helping me to do the work and
are most appreciated for that are Na'ama Katz who was involved in wza-promoter cloning and
characterization, Dr. Alessandro Franchini (EAWAG, Switzerland) together we ran chemostat
cultures and Dr. Masahiko Imashimizu (NIG, Japan) who prepared the templates and proteins
for the in-vitro transcription experiment. Laboratory members are all appreciated for their
time, patience and help. I would like to thank especially Sahar Melamed for his help with
microscopy, Sharon Yagur, Tal Elad and Sagi Magriso for the many discussions and advises
and above all, Dr. Rachel Rosen for her support all along the way. From the department of
Plant and Environmental Sciences I would like to thank Danny Ionescu for listening to all of
my ideas and of course for his advises, and Doron Eisentsat for helping with oxygen
measurements. From the Faculty of Medicine, Prof. Shoshy Altuvia and Dr. Maya ElgrablyWeiss are most appreciated for their help with promoter characterization. I also grateful to
researchers (cited in the text) from all over the world for E. coli strains and plasmid gifts.
Very special thanks go out to Prof. Thomas Egli (EAWAG) and Prof. Nobuo Shimamoto
(NIG) for inviting me to their laboratories and for their hospitality during my visit to these
institutes.
My wife Karmit is highly appreciated for providing me the time and peace to concentrate in
my thesis and of course for her support during the hard times.
And last, I would like to express my gratitude to my supervisor Prof. Shimshon Belkin whose
expertise, understanding and patience contributed a lot to my graduate experience. I
appreciate his vast knowledge, skill, vision, ethics, interaction with participants and his
assistance in writing the many reports along the way including this thesis. I also want to thank
him for supporting my ideas, understanding my problems and of course for his many many
advices.
ABBREVIATIONS
ampR – ampiciline resistance
PCR – polymerase chain reaction
CA-EPS – colanic acid EPS
ppGpp - guanosine-tetraphosphate
catR = chloramphenicol resistance
RFU – relative fluorescence units
cAMP – cyclic adenosine mono
RLU – relative light units
phosphate
RQ – relative quantification
CFU – colony forming units
RNAP – RNA polymerase
CR – Congo red
ROS – reactive oxygen species
CRP – cAMP receptor protein
rpm – rotations per minute
Ct – threshold cycle
RT-PCR – real time PCR or reverse
D – flow rate
FDG - fluorescein-di-β-Dgalactopyranoside
transcription
strepR – streptomycin resistance
tetR – tetracycline resistance
EPS – exopolysaccharide
Tn10 – transposon10
IPTG - isopropyl-beta-D-
VBNC – viable but not culturable
thiogalactopyranoside
kanR – kanamycin resistance
LB – Luria Bertani
LPS – lipopolysaccharide
µ - specific growth rate
OD600 – optical density measured at
600 nm
OMP – outer membrane protein
PBS – phosphate buffered saline
UV – ultra violet
X-gal - bromo-4-chloro-3-indolyl-ȕ-Dgalactopyranoside
UTR – untranslated region
ABSTRACT
Non-disinfected wastewater, routinely disposed to the sea in many countries, can
cause outbreaks of gastrointestinal diseases and inflict damage on aquaculture and
fishing. Nevertheless, the factors determining the fate of enteric bacteria in seawater
are not fully understood. The marine environment is extremely different from the
natural habitat of enteric bacteria and in order to survive they need to undergo drastic
biochemical, physiological and molecular changes. One of the seawater stressors is
salt stress, a growth-arresting factor that not only imposes osmotic shock and cell
dehydration but also a specific toxic effect of Na+. Escherichia coli responds to salt
stress by accumulating osmoprotectans and expelling Na+ out of the cytoplasm and
also by some less understood actions like outer membrane proteins (OMPs; porins)
exchange and production of exopolysaccharides (EPSs).
In a previous study it was reported that a yjbF'::luxCDABE transcriptional fusion was
induced upon exposure of E. coli to seawater as part of the response to osmotic stress
and that its induction was markedly enhanced in a strain lacking the general stress
response sigma factor protein RpoS (38). yjbF is a member of the yjbEFGH operon,
which is implicated in the production of an EPS of an unknown function.
In this study, using whole genome microarray analysis, I screened for genes that are
expressed similarly to yjbF: induced by osmotic stress and repressed by 38. Among
187 genes that fit these criteria stood out members of the wca operon, which is
implicated in the production of the colanic acid EPS (CA-EPS). The screen results
were further verified by RT-PCR and cloning of the promoter region of the wca
operon to generate a wza'::luxCDABE transcriptional fusion.
Both yjb and wca promoters were found to have almost identical regulatory
fingerprint: both were identically induced by osmotic stress, regulated by temperature,
activated by RcsC and RcsB and partially dependent upon RcsA (activated) and σ38
(repressed). The only difference between the activity of these two promoters was in
their intensity, which was 4-folds higher in the wca promoter.
This apparent negative regulation of yjb and wca operons by σ38 may be the result of
sigma factor competition for the RNAP. According to this model, in an rpoS mutant
the sigma factor protein that activates the yjb promoter has an improved access to the
RNAP, and therefore drives a stronger expression of this gene.
The role of σ38 in E. coli's physiology has been extensively studied. Most studies have
focused on characterizing genes positively controlled by σ38 (~ 450; 10 % of E. coli
genome) and on the physiological implications of under-expression of these genes in
σ38 deficient strains. Only a few reports refer to genes negatively controlled by σ38
and to the physiological consequences of their over-induction.
Positive selection for loss of σ38 has been shown to take place in starved cultures and
it has been suggested to be an adaptive trade-off of the general stress response in σ70dependent nutrient scavenging systems. The response of such a culture to osmotic
stress is no longer reliance upon σ38-dependent functions and an alternative strategy
has to be employed. In accordance with that idea I show that these operons are not
only overinduced in an rpoS mutant but the EPSs they are involved in synthesizing,
are also overproduced in this strain during osmotic stress; both phenotypes were
suppressed by inactivation of wca operon and yjb operon, respectively. The
physiological consequence of this overproduction is still unknown.
Interestingly, the yjbEFGH rpoS double mutant exhibited a heavily mucoid
appearance (overproduction of CA-EPS) even without subjecting it to osmotic stress;
in addition, cell morphology changed from the normal sized rod shape of the parental
strain into giant filaments in the double mutant. At the molecular level, the two
operons, wca and yjb, were 10-100 folds overinduced in this strain in an almost NaClindependent manner. The two phenotypes were observed also in the yjbE rpoS
mutant and suppressed by introducing the plasmid carrying the yjbF'::luxCDABE
transcriptional fusion (which also carries the yjbE gene) and surprisingly, also by the
plasmid carrying the wza'::luxCDABE transcriptional fusion (but not by
lacZ'::luxCDABE). The reason for this occurrence is unknown.
As σ38 does not activate yjb induction, the question of which sigma does activate its
stress responsive promoter was addressed. E. coli possesses a “house keeping” sigma
σ70 factor, as well as six known alternative sigma factor proteins. I employed a set of
strains with modified expression of the different sigma factors, harboring a
yjbF'::luxCDABE transcriptional fusion, and show that in-vivo none of those activate
yjbF induction. σ70 - like σ38 – appears to repress yjbF induction. In contrast, primer
extension analysis combined with in-vitro transcription indicated that transcription
from one of the two promoters that are located in the only promoter region of the
yjbEFGH operon is driven by RNAP associated with σ70.
σ70 manipulation is problematic since the effect is universal. Hence, overinduction of
yjbF in σ70 manipulated strains can be attributed to many different mechanisms and
does not rule the possibility that it is transcribed by the RNAP-σ70. Based on these
data it can be hypothesized that σ70 is likely to be the sigma factor protein that
activates yjb promoter regardless of the in-vivo results. Less conservative explanations
might suggest that yjb transcription can be carried out by several sigma factors or by a
novel sigma factor(s) that has not been discovered yet.
In parallel to the genetic work, we also studied the factors that limit yjbF'::luxCDABE
induction to the early stages of growth in LB medium. Growth in a batch culture in a
rich medium is accompanied by increased cell density, decreased specific growth rate
and changes in the medium properties. Continuous and batch culture experiments
indicated that the yjb operon is induced during slow growth (=0.1 h-1) in an osmotic
stress- independent manner, and that in slow growing cells (=0.1 h-1) yjb induction
by osmotic stress is 5-folds stronger than in fast growing cells (>0.2 h-1). This
behavior is typical to many σ38 -dependent genes which are induced during entry to
stationary phase; unlike these general stress response genes, yjb induction was found
to be repressed by increased cell density. Further investigation indicated that yjb
induction is sensitive to oxygen concentration. When cell density increases, oxygen
consumption causes a depletion in oxygen that prevents yjb promoter activation.
Our results suggest that the yjb and wca operons are members of an alternative stress
response regulon that might compensate for σ70 -deficiency. The contributions of EPS
and other σ38-repressed characters to cell survival in σ38 deficient strains are still
needed to be studied. The proposed Yjb regulatory role, expressed only in σ38
deficient strain, together with the new insights into the positive selection for loss of
σ38 function, suggest that despite the importance of σ38, it is dispensable and in its
loss, other functions emerge and may provide different strategies to cope with
stresses.
CONTENTS
1. INTRODUCTION…………………………………………………………………9
1.1 When Escherichia coli goes to the beach…………………………………9
1.2 The model microorganism used in this study……………………………..9
1.3 The marine environment…………………………………………………10
1.4 Stress responses in E. coli………………………………………………..11
1.4.1 Hyperosmotic stress response………………………………….11
1.4.2 Maintenance of cytoplasm pH…………………………………13
1.4.3 Oxidative stress………………………………………………...14
1.4.4 Starvation………………………………………………………15
1.5 Role of sigma factor proteins: switching from growth to survival……….16
1.6 σ38, the general stress response master regulator………………………...18
1.7 Genes induced upon exposure to seawater……………………………….19
1.8 The role of σ38 in E. coli response to seawater exposure………………...20
1.9 Negative regulation by σ38…………………………….…………………21
1.10 Adaptive inactivation of rpoS gene…………………….……………….22
1.11 The fate of enteric bacteria in seawater………………….……………...22
1.12 The yjbEFGH operon of E. coli………………………………………...24
1.13 The Rcs phosphorelay pathway………………………….……………...25
1.14 EPSs production in E. coli K12…………………………………………29
2. OBJECTIVES OF THIS THESIS…………………………………………………30
3. MATERIALS AND METHODS………………………………………………….31
3.1 E. coli K12 strains and sub-strains used in this study……………………31
3.2 Monitoring plasmid copy number………………………………………..34
3.3 Media and growth condition……………………………………………..35
3.4 Allele distribution by P1 coliphage (transduction)……………………....36
3.5 Construction of new inactive alleles……………………………………..37
3.6 Chemical transformation of plasmid DNA………………………………38
3.7 Cloning (plasmid construction)…………………………………………..39
3.8 Monitoring promoter induction: bioluminescence assays………………..45
3.9 Monitoring promoter induction: colorimetric assays…………………….46
3.10 Total RNA extraction…………………………………………………...47
3.11 Whole cell DNA-microarray analysis…………………………………..47
3.12 Relative Real Time PCR analysis……………………………………....49
3.13 Primer extension ………………………………………………………..50
3.14 In-vitro transcription assays…………………………………………….50
3.15 Bacterial growth - continuous culture…………………………………..51
3.16 Modifying aeration regime and gas exchange capacity………………...52
3.17 EPS-related phenotype characterization………………………………...53
3.18 Light/fluorescence microscopy…………………………………………53
3.19 Characterization of bacterial pellet volume formed upon centrifugation...…...54
3.20 Buoyancy assay…………………………………………………………57
3.21 Total carbohydrates determination……………………………………...57
4. RESULTS………………………………………………………………………….58
4.1 EPSs synthesis in response to osmotic shock…………………………….58
4.1.1 The basic phenomenon: yjbEFGH is induced by elevated osmotic
pressure……………………………………………………….58
4.1.2 Screen for yjb like regulated genes……………………………..60
4.1.3 The yjb and wca promoters display similar induction patterns...........64
4.1.4 Overproduction of EPS in the background of RpoS inactive
strains………………………………………………………...69
4.1.5 Phenotypes in yjb rpoS::Tn10 double mutants……………….71
4.1.6 A novel method for EPS quantification………………………..74
4.1.7 Activity of the yjb and wca promoters is enhanced in the
yjbEFGH rpoS::Tn10 mutant………………………………76
4.1.8 Suppression of the yjbE rpoS::Tn10 mutant mucoid and
filamentous phenotypes………………………………………78
4.2 Characterization of yjb promoter region…………………………………80
4.2.1 Which sigma factor(s) control yjb operon induction?.................80
4.2.2 Identification of the promoter region, promoters and transcription
starts sites…………………………………………………….80
4.2.3 70 can drive transcription from yjb P1 in-vitro………………..85
4.2.4 Regulation of yjbF'::luxCDABE by 70 in-vivo………………..87
4.2.5 Regulation of yjbEFGH promoter by alternative sigma factors……..91
4.2.6 In-vivo characterization of transcription pausing sites in yjb………..94
4.3 Regulation of yjb promoter activity by growth conditions……………… 96
4.3.1 Induction of the yjb operon occurs in early batch growth……...96
4.3.2 Induction of the yjb operon is not quorum-sensing dependent……....98
4.3.3 Induction of the yjb operon is highest in minimal media and at
low growth rates……………………………………………...98
4.3.4 Induction of the yjb operon is cell density related…………….102
4.3.5 Induction of the yjb operon depends upon gas exchange rates…….103
4.3.6 Induction of yjbF is oxygen dependent……………………….108
5. DISCUSSION……………………………………………………………………110
5.1 What regulon does the yjb operon belong to?.........................................110
5.2 Comparison between the yjb and wca regulatory regions……………...111
5.3 Overproduction of EPSs in an rpoS deficient strain……………………114
5.4 The role of yjb genes in rpoS deficient strain…………………………..115
5.5 A novel method for EPS production detection and comparison………..118
5.6 The yjbEFGH promoter region…………………………………………118
5.6 Regulation of yjb operon by sigma factors……………………………..119
5.8 Regulation of yjbEFGH induction by oxygen and specific growth rate……...121
5.9 EPSs production in response to osmotic stress…………………………123
6. REFERENCES…………………………………………………………………..125
7. SUPPLEMENTARY INFORMATION…………………………………………149
1. INTRODUCTION
1.1 When Escherichia coli goes to the beach
Disposal of non-disinfected wastewater to the sea is a daily practice in many
countries. The resulting introduction of enteric bacteria to the marine environment can
cause outbreaks of gastrointestinal diseases, and inflict damage to aquaculture and
fishing (Belkin and Colwell, 2005). Nevertheless, the factors determining the fate of
such microorganisms in seawater are not fully understood.
The parameters controlling the survival of enteric bacteria in seawater have intrigued
scientists for decades, and numerous studies have explored the fate of enteric bacteria
following their exposure to seawater. Some studies were driven by public health
concerns and others by the will to expand our knowledge on bacterial stress responses.
In this study, motivated by both reasons, the role and contribution of two Escherichia
coli K12 gene clusters to seawater survival were put to the test. The two operons, yjb
and wca, are both involved in exopolysaccharide (EPSs) synthesis - a physiological
aspect that was poorly studied in the context of bacterial stress responses.
1.2 The model microorganism used in this study
E. coli is one of the first and the most intensively studied model microorganisms.
Undoubtedly more is known about this bacterium than about any other microorganism.
It was thus the major model for studying the fate of enteric bacteria in seawater as
well as to map the physiological and the molecular nature of the bacterial stress
response. E. coli was discovered by German pediatrician and bacteriologist Theodor
Escherich in 1885 and is now classified in the Enterobacteriaceae (enteric bacteria)
family of the gammaproteobacteria class. Enteric bacteria are gram negative,
nonsporulating, facultative anaerobic rods. Among those are many bacteria
pathogenic to humans, animals and plants.
E. coli is also a mesophile and a copiotroph. A mesophile is an organism that grows
best at temperatures ranging from 20-45 ºC and a copiotroph is a heterotrophic
microorganism that requires a high organic carbon concentration in order to grow.
While sporulating bacteria such as gram-positive bacilli can differentiate into stress
resistance forms (endospores) upon exposure to suboptimal environments,
nonsporulating bacteria like E. coli are forced to undergo a highly sophisticated
cellular reorganization in order to develop multistress resistance characteristics
through a series of physiological and even genetic changes.
In this study the commensal laboratory E. coli K12 strain was used as a model
microorganism. It is the most studied strain and the easiest one to handle and
genetically manipulate. In addition, there exists a vast collection of previously
characterized strains, mutants, sub-strains and plasmids available from bacterial stock
centers and active laboratories around the world. The strains used in this study are
described in Table 1.
1.3 The marine environment
The marine environment is extremely different from the natural habitat of enteric
bacteria. The transfer from the neutrally buffered (pH 7), dark, warm (37 ºC) and
nutrient rich human gut into seawater imposes the need to adapt to an alkaline (~ pH
8.2-4), sunlight-irradiated, colder, saline (3.6-4.2%) and nutrient deficient
environment. In addition to these abiotic parameters, there are also biotic stress
factors such as predation (by protozoa, phages and bacteria) and competition with
local bacterial flora for the very limited resources. Thus, to survive the shift from their
natural habitat into seawater, enteric bacteria need to undergo drastic biochemical,
physiological and molecular changes.
1.4 Stress responses in E. coli
Outside their host, enteric bacteria are challenged by changing environmental
conditions. To adapt to and survive these changes, these bacteria posses global
response systems that drive changes in gene expression, in cellular physiology,
metabolism and even in the very basic sequence of their genome. These responses are
controlled by master regulators such as sigma factor proteins, signaling systems, small
molecule effectors, small RNAs and also by some inorganic molecules (for review see
Foster, 2007). Various stresses are being sensed differently and require different
responses; in respect to seawater stress, four abiotic stresses are most relevant:
osmotic shock, pH elevation, oxidative stress and starvation.
1.4.1 Hyperosmotic stress response
Seawater induces a salt stress, a growth-arresting factor that not only imposes osmotic
shock and cell dehydration but also a specific toxic effect of Na+ on certain metabolic
reactions (for review see Padan and Schuldiner, 1994). E. coli is a moderately
osmotolerant bacterium that can cope with osmotic pressure and dehydration by
accumulating compatible solutes (osmoprotectants) through synthesis or transport (for
review, see Wood, 2006). Compatible solutes are osmolytes the cytoplasmic
concentrations of which can be modulated over a broad range without disrupting
cellular function. The bacterial cell is equipped with several kinds of osmosensing
transporter proteins that can detect increasing extracellular osmotic pressure and
respond by mediating the uptake of these solutes. Accumulation of those in the
cytoplasm restores cellular hydration and volume. The immediate response to high
osmolarity is by the uptake of K+, an inorganic compatible solute the availability of
which is sensed by the KdpDE two-component regulatory system. This system
activates the expression of the high affinity K+ transport system KdpFABC upon an
elevation in osmotic pressure (Walderhaug et al., 1992; Sugiura et al., 1994). In order
to maintain the membrane potential, increased K+ concentration must be balanced by
the accumulation of anions. The major accumulated counterion in E. coli is glutamate
(Measures, 1975), which is also an osmoprotectant. High cytoplasmic K+
concentrations can inhibit the functioning of key enzymes (Arakawa and Timasheff,
1985); therefore, as a secondary response to high environmental osmolarity, the cell
replaces much of it with organic compatible solutes such as glycine betaine, ectoine
and proline. Such compounds restore hydration more effectively and also act as
protein stabilizers (chemical chaperons). The organic compatible solutes are either
transported from the environment by transporters such as ProP (Grothe et al., 1986;
Jebbar et al., 1992; MacMillan et al., 1999) and ProU (encoded by proVWX; Lucht
and Bremer, 1994), or synthesized de-novo following an osmotic up-shift. Choline
and betaine are transported into the cell and converted into glycine betaine by the betgenes encoded system following osmotic shock (Styrvold et al., 1986). Similarly,
trehalose is synthesized from UDP-glucose and glucose-6-phosphate by OtsA and
OtsB (Stryvold and Ström, 1991).
Another membrane sensor kinase protein that senses osmotic elevation, EnvZ,
regulates the level of phosphorylated OmpR in the cell. OmpR in turn regulates
transcription of a number of genes, the best studied of which are ompF and ompC,
which encode the two major porins in the outer membrane. Low levels of
phosphrelated OmpR (low osmolarity) activate ompF transcription and high levels
(high osmolarity) repress ompF and activate ompC (Pratt et al., 1996; Lan and Igo,
1998). Although the adaptive consequence of this replacement is not clear, ompC
mutants exhibit sensitivity to osmotic stress at alkaline pH (Heyde et al., 1987;
Thomas and Booth, 1992).
1.4.2 Maintenance of cytoplasm pH
Seawater pH is alkaline (~8.2-8.4). E. coli must maintain a cytoplasmic pH
compatible with optimal function and integrity of cytoplasmic proteins and for that
purpose it possesses several adaptive strategies for alkaline pH homeostasis (for
review, see Padan et al., 2005).
Seawater contains over ~450 mM sodium ions (Na+) that are more toxic to cells than
potassium ions, making it crucial to expel them from the cytoplasm. The reason for
this enhanced toxicity is not fully understood but all cells have efficient Na+ pumping
systems to eliminate these cations from the intracellular environment (for review, see
Padan and Schuldiner, 1994). One of E. coli's sodium pumps is NhaA, a sodium
ion/proton antiporter that uses the proton electrochemical gradient built during
respiration to expel sodium ions from the cytoplasm and functions primarily in the
adaptation to high salinity at alkaline pH. Expectedly, it is induced by Na+ and
alkaline pH (Dover and Padan, 2001); protein activity is optimal at pH 9 and non
active below pH 6.5 (Taglicht et al., 1991). H+ entry in exchange for Na+ is essential
for acidification of the cytoplasm.
In addition to sodium ion/proton pumps, E. coli alters the activity of components
involved in proton metabolism (Maurer et al., 2005); flagella synthesis is repressed
since its function expends proton motive force while electron transport and ATPase
components that raise proton concentration inside the cell are stepped up in order to
accelerate proton import. If available, the bacterium is capable of importing and
utilizing sugars in order to generate a large burst of fermentation acids. Similarly,
amino acids can be deaminased in order to produce acidic compounds.
1.4.3 Oxidative stress
Oxidative stress in seawater is threat only in the upper layer of the water column that
is penetrated by the short wavelength fraction of solar radiation. As will be described
later (section 1.4.4 and 1.11), continuous nutrient deficiency also leads to oxidative
stress. Reactive oxygen species (ROS), such as superoxide (O2-), hydrogen peroxide
(H2O2), and the hydroxyl radical (OH·) are toxic by-products of aerobic metabolism
and may also be generated by radiation.
ROS can oxidize any cell component including nucleic acids, proteins and fatty acids.
When ROS levels are kept low enough, specific antioxidants enzymes eliminate them
and the oxidative damage is repaired (including DNA damage repair and aberrant
proteins removal). Superoxide dismutase (SodA or SodB) reacts with superoxide to
form hydrogen peroxide (Carlioz and Touati, 1986). Hydrogen peroxide can be
transformed into one of the most destructive free radicals, hydroxyl radical, by the
Fe2+-mediated Fenton reaction; to prevent this from occurring, it is further neutralized
by catalases (KatE and KatG) or peroxidases (Loewen, 1984; Imlay and Linn, 1987).
Another way to deal with oxidative stress is to avoid internal ROS production by
increasing anaerobic respiration (Iuchi and Weiner, 1996). These basic acts of
protection, however, are not sufficient to protect against sudden large increases in
ROS production.
1.4.4 Starvation
E. coli reacts to lack of nutrients in two steps (Matin, 1991). The first step is
scavenging; when a particular nutrient becomes limiting and needs to be foraged from
the environment. This scavenging is controlled by CRP (Cyclic AMP Receptor
Protein), which allows the use of alternative carbon sources, and the two-component
regulatory systems Ntr and Pho, which control scavenging for nitrogen and
phosphorus, respectively (for reviews see Reitzer, 2003; Wanner, 1993). CRP senses
carbon source status through intracellular cAMP level; NtrC responds to glutamine;
the Pho system monitors inorganic phosphate levels via the activity of the Pst
transport system (Wanner, 1996). When scavenging fails, as is bound to happen when
the supply of nutrients is finite, the cells starve and enter the second step, stationary
phase.
During starvation, E. coli downregulates rRNA biosynthesis (Gourse et al., 1986 and
1996; for review see Gralla, 2005) and ribosome production, a phenomenon initially
associated with amino acids starvation and known as the stringent response (Irr, 1972;
Atherly, 1974). The molecule that regulates this response is the small nucleotide,
guanosine tetraphosphate, ppGpp, which is produced in response to diverse types of
nutrient limitations and conditions that cause growth arrest (for review see Cashel et
al., 1996; Potrykus and Cashel, 2008). E. coli uses two different pathways to
synthesize ppGpp: the RelA and the SpoT-dependent pathways. RelA is a protein that
is associated with the ribosome and produces ppGpp in response to uncharged tRNA
that approaches the ribosomal translation initiation site during amino acid starvation
(Haseltine et al., 1972; Haseltine and Block, 1973; Pedersen et al., 1973). Less is
known about SpoT-dependent production of ppGpp (which is also responsible for
hydrolyzing ppGpp; Hernandez and Bremer, 1991; Xiao et al., 1991) and how SpoT
senses starvation conditions. Yet it appears that SpoT is responsible for the
accumulation of ppGpp in response to most stresses and nutrient limitations apart
from amino acid starvation. ppGpp binds to the and subunits of the RNA
polymerase (RNAP) core enzyme (Reddy et al., 1995; Chatterji et al., 1998) and
affects gene expression: it represses rRNA synthesis and in parallel activates large
number of stress related genes (see section 1.5 for more details).
Long incubation in a non-proliferating state due to nutrient depletion leads to a
gradual loss of the ability to recover and reproduce (see section 1.11). These sterile
cells initially remain intact as individuals but may eventually lose their membrane
integrity and life supporting activities (Ericsson et al., 2000).
1.5 Role of sigma factor proteins: switching from growth to survival
The main responses to changing conditions take place at the transcriptional level,
where unnecessary genes are silenced and relevant genes are activated or stimulated.
This action is mediated by signaling pathways, various transcription regulators and
finally by the RNAP itself and associated factors.
E. coli RNAP consists of five subunits (2’) that make up the core enzyme, and
accessory replaceable sigma subunits, which together form the RNAP holoenzyme.
The sigma subunit recognizes the promoter element of a gene and directs RNAP to
transcribe it. Seven different sigma factor proteins have been described in E. coli,
each with a different affinity to the RNAP core enzyme and a different concentration
in the cytoplasm. Both the affinity and concentration are important for the
intracellular competition for free RNAP (Maeda, et al., 2000; Hiroto et al. 2000;
Farewell et al. 1998).
The seven known sigma factors are generally considered to control different regulons;
σ70 (RpoD), the dominant amongst them, regulates the induction of "housekeeping
genes" which fulfill the essential metabolic needs of the bacterium; σ38 (RpoS)
controls the general stress response regulon (Weber et al., 2005); σ32 (RpoH), the heat
shock response regulator (Grossman et al,. 1984; Rosen and Ron, 2002), controls a set
of chaperons and proteases which manage protein folding and refolding or proteolysis
of damaged proteins; σ54 (RpoN) regulates genes mostly involved in nitrogen
assimilation (Hunt and Magasanik, 1985); σ24 (FliA) regulates flagella synthesis; σ28
(RpoE), another heat shock response regulator (Rouvière et al., 1995), responds to
defectively folded proteins in the periplasm space; σ19 (FecI) regulates the induction
of a single operon, fecABCDE, involved in iron metabolism (Van Hove et al., 1990;
Braun, 1997).
Despite this seemingly straightforward categorization, a surprisingly extensive
functional overlap exists between σ70 promoters and those of σ32 and σ28 (Wade et al.,
2006). Furthermore, some of the sigma factors were shown to activate their regulons
under conditions other than the anticipated ones, or to appear to regulate genes that do
not fall into their expected metabolic context. For example, the heat shock regulons
were found to be induced by hyperosmotic shock (Bianchi and Baneyx, 1999) as well
as by a long list of toxic chemicals (Van Dyk et al., 1994). In addition to genes
involved in nitrogen metabolism, σ54 also controls formate, acetoacetate and
propionate catabolic genes and others that respond to phage shock, zinc and lead
(Reitzer and Schneider, 2001). σ70 itself plays a dual regulatory role: it regulates
essential (house keeping) genes during growth, but is redirected to activate a parallel
stress response regulon when the bacterium is challenged by suboptimal environments
and growth is inhibited. This transcriptional switch is mediated by the synthesis of the
regulatory nucleotide, ppGpp (see section 1.4.4), which binds RNAP and (a) reduces
the affinity of the normally dominant σ70 to the RNAP core, (b) allows alternative
sigma factors such as σ38 to compete successfully for RNAP and, in parallel, (c)
changes RNAP-σ70 conformation to activate promoters of genes involved in cell
maintenance rather than proliferation (Chang et al., 2002; see Nyström, 2004, for
review).
1.6 σ38, the general stress response master regulator
During adaptation to stress, σ70 and σ38 are the most dominant sigma factors in the
cytoplasm. Although most studies have focused on the contribution of σ38 and its
regulon to E. coli stress response, σ70 still performs the majority of transcription
actions in the cell even when σ38 level increases (Jishage and Ishihama, 1995).
σ38 is subject to complex regulation at all levels (transcription, translation, activity,
and protein degradation). During growth, σ38 levels in the cell are very low and its
transcription is driven from the promoters of the upstream gene nlpD. Upon growth
arrest, the transcription is reinforced by the activity of promoters located inside the
nlpD gene; both are σ70 and ppGpp dependent (Lange et al., 1995). Its mRNA
contains a long 5' untranslated region (UTR) which plays an important role in the
regulation of its stability and translation by proteins and small RNA molecules (OxyS,
DsrA and RprA; for review see Repolia et al., 2003). The σ38 level is regulated by
ClpXP protease degradation (Schweder et al., 1996).
The rpoS gene is strongly induced when E. coli cells are exposed to various stress
conditions, which include starvation, hyperosmolarity, pH downshift, or nonoptimal
temperatures (for review, see Hengge-Aronis, 2000). Under these conditions σ38 was
found to activate the expression of 10% of E. coli's genes (Weber et al., 2005), a
regulon that exhibits extensive regulatory overlaps with other global regulons such as
the CRP-cAMP regulon. Many of the genes were found to be induced in response to
several stresses, allowing cross protection against additional potential stressors.
Among the genes regulated by σ38 are genes that encode antioxidants,
osmoprotectants, membrane proteins involved in scavenging of various nutrients,
efflux pumps that may increase resistance against various toxic compounds and genes
involved in "housekeeping" actions like glycolysis, fermentation, anaerobic
respiration, electron transport and the pentose phosphate pathway.
1.7 Genes induced upon exposure to seawater
Adaptive physiological changes are usually linked to changes in gene expression
profile. Rozen et al. (2001, 2002) used an E. coli 687-member promoter-reporter gene
fusion library (Van-Dyk et al., 2001) to assay the immediate gene induction following
exposure of E. coli to low-nutrient seawater and a DNA microarray chips (Wei et al.,
2001) to reveal changes in RNA patterns following longer incubation in seawater. In
the latter case it was assumed that the “emergency response” is over and that the only
genes being transcribed are involved in continuous maintenance of viability.
In the immediate response experiment (Rozen et al., 2001), the strains each harboring
a different promoter::luxCDABE fusion, were shifted from rich medium (LB) into
artificial seawater supplemented with 10% of LB nutrients. These unfavorable
conditions led to a reduction in light production by 90% of the strains and to a
significant increase in light production of only 3% (22 promoters). When the
combined effect of artificial seawater was broken into its different environmental
factors (salinity, nutrient limitation and alkaline pH) it appeared that salinity or
osmolarity were involved in only four cases, among them the expected osmY, which
encodes hyperosmotically inducible periplasmic protein of an unknown function (Yim
and Villarejo, 1992; Weichart et al., 1993), yjbG that was recently reported to be
involved in EPS synthesis (Ferrièrs et al., 2007) as well as two previously
“unassigned” genes. Among the other promoters, the most important environmental
factor appeared to be nutrient limitation. This observation included otsA (encoding
trehalose 6-phosphate synthase; Giaever et al., 1988) and treA (encoding periplasmic
trehalase; Boos et al., 1987), both known to be osmotically regulated, which seemed
in this system to be induced by nutrient limitation rather than by salinity.
In the longer incubation experiment (Rozen et al., 2002), a culture was incubated in
artificial seawater for 20 hours. Analysis of the expression of functional gene groups
(Serres and Riley, 2000) indicated that the expression of gene groups involved in cell
division and in nucleotide biosynthesis was down-regulated following seawaterexposure while an increase was observed in the expression of genes the products of
which are involved in either carbon compound degradation or in respiration,
signifying the dearth of carbon and energy sources under such conditions. An
additional potential that was built within this period was motility, potentially
equipping the cells either for a chemotactic search for nutrition or for flight from the
inhospitable conditions imposed upon them.
1.8 The role of σ38 in E. coli's response to seawater exposure
In studies attempting to determine the fate of enteric bacteria in seawater, die-off rates
based on colony formation capabilities were often the main parameter used to
characterize bacterial survival. Based on survival rates of mutants in comparison to
their wild types, only six genes were revealed as crucial for seawater survival when
the experiment was performed in the dark. The most dominant among them was rpoS
(Gourmelon et al., 1997; Munro et al., 1994, 1995; Troussellier et al., 1998). The
other genes in which mutations were reported to be significant were otsA (Munro et
al., 1989), relA, spoT on top of a relA deletion (Munro et al., 1995) and both outer
membrane porins ompC and ompF encoding genes (Gauthier et al., 1992a).
As already mentioned above, RpoS is the general stress response sigma factor protein
(38), involved in the stress-activation of more than 10% of E. coli K12 genes (Weber
et al., 2005). Indeed, Rozen et al. (2001) reported that 17 out of the 22 promoterelements that were induced following exposure to seawater are positively regulated by
σ38. Among those was otsA (Hengge-Aronis et al., 1991; Fang et al., 1996; Schellhorn
et al., 1998; Weber et al., 2005) that was also listed among the genes affecting E. coli
viability in seawater (Munro et al., 1989). Interestingly, the promoter element of yjbF
(mistakenly designated by Rozen et al. (2001) as yjbG) stood out for being negatively
dependent upon σ38. This observation was one of several that made yjbF the focus of
this PhD thesis.
1.9 Negative regulation by σ38
This apparent negative regulation of yjbEFGH by σ38 may be the result of a direct
effect of a σ38-dependent repressor. Another explanation, in complete accordance with
all of our data, is based on the sigma factor competition model (Maeda et al., 2000;
for review see Nyström, 2004), which claims that in the absence of a sigma factor,
other sigma factors may compete more efficiently for the RNAP. According to this
model, in an rpoS mutant the sigma factor protein that regulates yjbEFGH has an
improved access to the RNAP, and therefore drives a stronger expression of this gene.
1.10 Adaptive inactivation of rpoS gene
The role of σ38, the general stress response sigma factor protein, in E. coli's
physiology has been extensively studied. While most studies have focused on
characterizing genes positively controlled by σ38 and on the physiological
implications of the under-expression of these genes in σ38-deficient strains (e.g.
Hengge-Aronis et al., 1993), only few refer to genes negatively controlled by σ38 and
to the physiological consequences of their overexpression. It has been shown that in a
strain lacking σ38, genes controlled by the vegetative sigma factor RpoD (70) are
overexpressed as a consequence of enhanced availability of sigma-free RNA
polymerases (Farewell et al., 1998). Other reports indicated that the rpoS gene tends
to undergo frequent mutations that lead to activity loss, and that the mutated forms
appear to spread and become dominant in glucose-limited chemostat cultures (NotleyMcRobb et al., 2001; Chen et al., 2004; Ferenci, 2007) or during prolonged incubation
in stationary phase (Zambrano et al., 1993). It was also shown that the rate of rpoS
mutations spreading throughout the population decreases when osmotic stress is being
applied to the system. These observations suggest that rpoS inactivation may be
beneficial to a starved cell due to overexpression of σ70-dependent nutrient sensing
and uptake systems but when σ38-dependent genes are required (e.g. osmotic stress
response genes) the rate of inactivation declines.
1.11 The fate of enteric bacteria in seawater
Incubation of enteric bacteria like E. coli or Salmonella typhimurium in seawater is
accompanied with a decay in their culturability, an observation that was initially
thought to indicate death of the microorganisms (Grimes et al., 1986) but later shown
to be mainly due to entering into a "dormant" state termed VBNC (Viable But Not
Culturable; Roszak and Colwell, 1987; Munro et al., 1987; Duncan et al., 1994). The
same observation is valid for other hostile conditions like the stationary phase of
growth as defined for laboratory cultures (Roszak and Colwell, 1987; Dukan and
Nyström, 1999). The main factor pushing cells into a VBNC state is nutrient
deprivation; interestingly, the rate of culturability loss can be altered based on culture
growth history and genotype. Stationary phase cultures or cultures grown in high
osmolarity lose their culturability at a slower rate than exponential growth phase
culture or a culture grown in low osmolarity (Munro et al, 1995). Monitoring the first
events leading to loss of culturablity in stationary phase suggests that it occurs as a
consequence of oxidative damage (Cuny et al., 2005). Another important discovery
was that in a starved culture only a sub-population becomes non-culturable. This
population undergoes drastic morphological changes that allow it to be separated from
the culturable sub-population based on its small cell-size and cell-spherical shape.
Desnues et al. (2003) have recovered both sub-populations from radioselectan-density
gradient formed upon ultra-centrifugation and showed that loss of culturablity is
accompanied with an increased level of carbonylated proteins, a result of oxidative
stress. The same authors suggest that the oxidative stress is a result of a decline in
sodA and sodB expression level (superoxide-dismutase A and B; antioxidant defense
system) expression level and that the accumulation of carbonylated proteins is
counteracted with an increased expression of rpoH, chaperons and proteases
(oxidized-protein removal by the heat shock response system). These data suggest that
VBNC may not be an adaptive strategy (differentiated physiological state like an
endospore) but a rather a stochastic deterioration of cell components.
1.12 The yjbEFGH operon of E. coli
yjbF is a member of a four genes operon, yjbEFGH. Ferrièrs and Clarke (2003) listed
it among genes positively dependent upon RcsC, the inner membrane sensor kinase of
the Rcs phosphorelay pathway. The Rcs-system, to be described in details later,
controls a variety of physiological functions in prokaryotes such as extracellular
polysaccharides synthesis (Stout and Gottesman, 1990; Virlogeux et al., 1996; Kelm
et al., 1997; Rahn and whitfield, 2003), biofilm formation (Prigent-Combaret et al.,
2000; Danese et al., 2000; Ferrièrs and Clarke, 2003), cell division (Carballes et al.,
1999) and motility (Tekada et al., 2001; Francez-Charlot et al., 2003). Ferrièrs et al.
(2007) have shown that the four genes of the yjb operon are transcribed together and
that operon is induced during growth on solid surface and is involved in the
production of an unknown EPS. Nevertheless, inactivation of any of the yjb genes did
not reduce biofilm formation ability of the bacterium (Ferrièrs and Clarke, 2003;
Ferrièrs et al., 2007).
Few other studies provided additional information regarding gene members of the
yjbEFGH operon. yjbE was found to be repressed by YdjG, a biofilm-induced protein
(Herzberg et al., 2006) and to be induced by ampiciline and ofloxacin (Kaldalu et al.,
2004). The same dependency upon the Rcs phosphorelay system was also
demonstrated for yjbH in Salmonella enterica (García-Calderón et al., 2007). YjbH
was predicted to be a -barrel shaped (Bigelow et al., 2004) outer membrane protein
(Ying et al., 2005).
Based on the Expasy protein server (http://www.expasy.ch/) the four proteins encoded
by the yjbEFGH operon belong to a family of uncharacterized conserved proteins. All
of those contain a signal peptide, that relates them to the cell envelope. YjbH is
predicted to be an uncharacterized lipoprotein (has a cystein in position 18 that
potentially binds a lipid molecule and forms N-palmitoyl cysteine or S-diacylglycerol
cysteine). No further functional annotations were available for those proteins.
1.13 The Rcs phosphorelay pathway
The Rcs phosphorelay system was first identified to be a positive regulator of the
production of an EPS called colanic acid (CA-EPS) in E. coli (see Figure 1 for a
model). This EPS, barely produced in the wild type under normal growth conditions,
was originally found to be overproduced in cells carrying an inactive lon gene
(Markovitz, 1964). Lon mutants produce mucoid colonies, a definition referring to a
thick cover of CA-EPS that endows the colony with a shiny and slimy appearance.
Lon mutants were also found to be sensitive to UV radiation, forming long filaments
after UV treatment. The lon gene encodes an ATP dependent protease (Charette et al.,
1981; Chung and Goldberg, 1981) that in addition to its role in misfolded proteins
degradation during heat shock, also has a regulatory function (for review see
Tsilibaris et al., 2006). The two phenotypes, mucoidity and UV sensitivity were
linked to an overexpression of several proteins (Markovitz, 1964), two of which were
later identified as Lon specific substrates RcsA (Torres-Cabassa and Gottesman,
1987) and SulA (Mizusawa and Gottesman, 1983).
SulA is a cell division inhibitor induced during SOS response (error-prone DNA
damage repair; Schoemaker et al., 1984). It binds to FtsZ, a key protein in cell
division which is responsible for septum formation (Bi and Lutkenhaus, 1991;
Higashitani et al., 1995), and represses the cell cycle (Mukherjee et al., 1998).
Figure 1: A model describing the major components involved in the Rcs-signaling pathway. H,
histidine; D, aspartate. Arrows indicate a positive effect; Blocked lines indicate a negative
effect.
RcsA was identified together with RcsB and RcsC only after the locus responsible for
CA-EPS production was found in a lon mutant; it overinduced a chromosomal
cpsB'::lacZ transcriptional fusion generated by random insertional mutagenesis that
also suppressed the mucoid phenotype. The locus was designated as cps, capsule
polysaccharide synthesis (Trisler and Gottesman, 1984) and it was recently renamed
wca so as to conform with the nomenclature of other capsules. This transcriptional
fusion was further used to screen for genes necessary for CA-EPS synthesis revealing
three mutations: rcsC, rcsB and rcsA (Gottesman et al., 1985). rcsC encodes the
sensor kinase of the signaling pathway that possesses a phosphoryl group receiver
domain in addition to its His-kinase domain, rcsB encodes the response regulator
composed of two domains, a receiver domain and a DNA binding domain (Brill et al.,
1988; Stout and Gottesman, 1990) and rcsA encodes the unstable auxiliary DNA
binding protein that is degraded by Lon (Torres-Cabassa and Gottesman, 1987).
In addition to RcsC, several proteins have been found to play a role in the signaling
pathway. The important ones are RcsF (Hagiwara et al., 2003; Castanié-Cornet et al.,
2006), DjlA (Clarke et al., 1997) and RcsD (Takeda et al., 2001). RcsD is an inner
membrane
protein
that,
like
RcsC,
possesses
a
histidine-containing
phosphotransmitter domain. Overproduction of the two other proteins, RcsF, an outer
membrane lipoprotein or DjlA, an inner membrane chaperon-like protein, was shown
to activate RcsC but the way the signal is transmitted is unknown.
RcsB, the response regulator, is a DNA binding protein that phosphorelated and
activated by RcsC, the inner membrane sensor kinase via RcsD (Takeda et al., 2001),
in response to an unknown cue. RcsB may then interact with RcsA to form a DNA
binding heterodimer, but since RcsA levels are generally low due to Lon activity, CA-
EPS is then produced at a low level. When RcsA levels increase (such as in a lon
mutant), the cells become mucoid.
The RcsAB heterodimer recognizes and binds a DNA consensus sequence designated
"RcsAB Box" (TaAGaatatTCctA; Wehland and Bernhard, 2000). This consensus
sequence was found upstream of wza (the first gene of the wca operon; Stevenson et
al., 1991; Stout V, 1996), rcsA (Ebel and Trempy, 1999) and yjbE (Ferrièrs et al.,
2007) promoters where RcsAB binding activates transcription. It was also found
downstream of the flagella master regulators (flhDC) operon promoter, where it
represses transcription (Francez-Charlot et al., 2003). Defective motility and flagella
synthesis were reported to take place in mdo (genes involved in membrane derived
oligosaccharide synthesis; Fiedler and Rotering, 1988) mutants, which also
overinduce the wca operon (Ebel et al., 1997). So far these are the only genes that
where found to be regulated by RcsAB. RcsB also regulates an RcsA-independent
regulon. The first RcsB-dependent RcsA-independent gene to be discovered was ftsZ
(Gervias et al., 1992; Carballes et al., 1999) and later RcsB was also shown to
positively regulate osmC and the small RNA rprA (Davalos-Gracia et al., 2001). RcsC
has been found to affect the expression of more than 150 genes (Ferrièrs and Clarke,
2003), half of which are predicted to encode proteins with functions related to the
bacterial surface. Based on the function of the Rcs-system, the RcsC-dependent genes
are expected to be also RcsB-dependent. However, the number of RcsC-dependent
genes (>150) is much greater than has been demonstrated for RcsAB and RcsB
together, indicating that the system and its regulatory targets are still poorly
understood.
1.14 EPSs production in E. coli K12
E. coli K12 possesses five sets of genes promoting EPS production: (a) genes
involved in O-antigen synthesis; (b) the CA-EPS synthesis genes (the wca operon;
Gottesman et al., 1985); (c) the pgaABCD operon which encodes genes involved in
the production of poly--1,6-N-acetyl-D-glucosamine polysaccharide that has been
shown to be crucial for biofilm formation (Wang et al., 2004); (d) the dfc pseudooperon (composed of gfcABCDE, etp and etk), responsible for the production of type
IV capsule in E. coli O127:H6 but is non-functional in E. coli K12 due to the presence
of an IS1 element in its promoter region (Peleg et al., 2005); and (e) the yjbEFGH
operon which is also a paralogue of dfcABCD (Ferrièrs et al., 2007).
Ferrièrs et al. (2007) demonstrated that overexpression of the whole yjbEFGH operon
but not of any of its individual gene members, results in the production of an
unknown EPS. Based on dye staining properties (Weiner et al., 1999), the yjbEFGH
operon has been suggested to be involved in the production of a polysaccharide that is
not cellulose and does not contain any (13)-- and (14)–-D-glucopyranosides
(does not bind calcofluor), could be helical (binds Toluidine blue-O) and possible
contains (14)--glucopyranosides units (binds Congo-red) (Ferrièrs et al., 2007).
2. OBJECTIVES OF THIS THESIS
This thesis focuses on the yjb operon, the promoter of which was found previously to
be induced upon a shift from rich medium into seawater, the regulon it belongs, its
regulation and other aspects like the consequence of its activation with special
emphasis on the nature of the negative regulation by σ38 (RpoS).
The following specific aims were addressed:
1. Identifying genes which are being co-transcribed with yjb in response to
osmotic stress (induced by salt stress and repressed by σ38).
2. Mapping the regulatory factors that affect yjb expression: molecular (sigma
factors, signaling pathways), physiological (growth phase, rate and culture
density) and environmental (nutrients availability).
3. Studying EPS synthesis overinduction in an rpoS deficient strain and its
physiological consequences.
3. MATERIALS AND METHODS
3.1 E. coli K12 strains and sub-strains used in this study
Several E. coli K12 strains are common among laboratories studying their biology,
including MG1655, W3110 and MC4100. Since being isolated in 1922 (Bachman,
1996), the K12 derivative MG1655, which was cured of its endogenous lambda phage
(Lam-) and its conjugal plasmid (F-), went through numerous manipulations including
X-ray, UV, chemical mutagenesis and genetic crossing to generate strain MC4100
that, on top of various point mutations, contains four deletions ranging from 1 to 97
kb in size (Peters et al., 2003). The strains used in this study are listed in Table 1. All
strains were first grown in LB broth supplemented with the appropriate chemicals
(antibiotics and strain specific essential metabolites) to an optical density of 1-2
(OD600; Spekol 1200, Analytik Jena, Germany) and then mixed with 50% glycerol
and kept at -80°C. Cells were recovered from freezing by overnight growth on LB
agar plates. Plates were kept up to one month at 4°C.
Table 1: E. coli strains used in this study
STRAIN
GENOTYPE
REFERENCE
MG1655 derived strains
MG1655
F-, Lam-
Laboratory stock
QC2410
MG1655 rpoS::Tn10
Loewen and Triggs, 1984
MK1399
MG1655 yjbEFGH (kanR)
This work
MK1310
MG1655 yjbE::kanR
This work
MK1320
MG1655 yjbF::kanR
This work
MK1330
MG1655 yjbG::catR
Yael Rozen PhD thesis
MK1340
MG1655 yjbH::kanR
This work
MG1655 fecI::kanR
MK176
This study
(allele introduced from MO704; Enz et al., 1995)
MG1655 fliA::kanR
This study
MK175
(allele introduced from WI11; Park et al., 2001)
MG1655 rpoN::kanR
MK152
This study
(allele introduced from ; Gardner et al., 2003)
MG1655 Pwza (kanR)
This work
MK1351
(allele introduced from MS1651; Peleg et al., 2005)
MK2499
QC2410 yjbEFGH (kanR)
This work
MK2431
QC2410 yjbE (yjbE::kanR)
This study
MK2432
QC2410 yjbF (yjbF::kanR)
This study
MK2433
QC2410 yjbG (yjbG::catR)
This work
MK2434
QC2410 yjbH (yjbH::kanR)
This work
MK2411
QC2410 Pwza (kanR)
This work
CF6343
MG1655 lacIZ(MluI)
Conter et al., 2002
SK1150
CF6343 rcsA::kanR
Francez-Charlot et al., 2003
SK1158
CF6343 rcsB::Tn10
Conter et al., 2002
SK1154
CF6343 rcsC::Tn10
Conter et al., 2002
MK2411
QC2410 Pwza (kanR)
This study
MK2421
MK2433 Pwza (kanR)
This study
MK2431
QC2410 rcsA::kanR
This study
MK2441
MK2433 rcsA::kanR
This study
MC4100 derived strains
-
MC4100
-
F araD139 D(argF-lac|)205 I flhD5301 fruA25 relA1
rpsL150(strR) rbsR22 deoC1
AG226
MC4100 groE-constitutive
Laboratory stock
Kusukawa and Yura, 1988
AG226 rpoH30::kanR
CAG9333
temperature sensitive (25ºC)
CAG16037
MC4100 rpoHP-lacZ (strepR)
Zhou et al., 1988
Rouvière et al., 1995
CAG16037 rpoE::catR
CAG22188
Rouvière et al., 1995
temperature sensitive (30ºC)
Lonetto et al., 1998
LM100
MC4100 Ptrp-rpoD, catR
LM1009
LM100 rpoS::Tn10 (allele introduced from QC2410)
This work
AF633
MC4100 uspB'::lacZ
Farewell et al., 1998
KK358
AF633 relA spoT (markerless)
Jishage et al., 2002
KK373
KK358 rpoD-DSA (536-538), (rpoD40; tetR)
Jishage et al., 2002
Magnusson et al., 2003
W3110 derived strains
W3110 cI857(cro-bioA)
DY378
Yu et al., 2000
(permissive strains for gene inactivation)
Table 2: Plasmids that were used in this study
PLASMID
GENOTYPE
REFERENCE
promoterless multi cloning site - luxCDABE,
Van Dyk and Rosson, 1998
ori pMB1 (pBR322), rop, ampR
Van Dyk et al., 2001
pDEW201
pDEW/yjbF
Van Dyk and Rosson, 1998
(pDEW/yjbF-1;
pDEW201, yjbF'::luxCDABE (pgi-yjbF)
Van Dyk et al., 2001
pDEW609)
pDEW/yjbF-2
pDEW201, yjbF'::luxCDABE (no RcsAB Box)
This study
pDEW/yjbE27-1
pDEW201, yjbE27::luxCDABE (pgi-yjbE)
This study
pDEW201, yjbE76::luxCDABE
pDEW/yjbE76-1
This study
(RcsAB Box-yjbE)
pDEW/yjbE27-2
pDEW201, yjbE27::luxCDABE (pgi-yjbE)
This study
pDEW201, yjbE76::luxCDABE
pDEW/yjbE76-2
This study
(RcsAB Box-yjbE)
pDEW/wza
pDEW201, wza'::luxCDABE
This study
pDEW/dfcB
pDEW201, dfcB'::luxCDABE
This study
pDEW/lacZ
Van Dyk and Rosson, 1998
pDEW201, lacZ'::luxCDABE
(pDEW229)
Van Dyk et al., 2001
Van Dyk and Rosson, 1998
pDEW/lon
pDEW201, lon'::luxCDABE
Van Dyk et al., 2001
Van Dyk and Rosson, 1998
pDEW/osmY
pDEW201, osmY'::luxCDABE
Van Dyk et al., 2001
pMLZ609
pDEW609 yjbF'::lacZ
This study
pAR1
pACYC184, araC::rpoH, catR
Aviram Rasuley PhD thesis
pKD13
Template plasmid for kanR amplification
Datsenko and Wanner, 2000
pES2
recA'::gfpUV, ori pUC, ampR
Sagi et al., 2003
3.2 Monitoring plasmid copy number
Since the transcriptional fusions employed in the present study were plasmid based,
plasmid copy number was monitored routinely. Plasmids were extracted from culture
aliquots removed at intervals during experiments using QIAprep spin Miniprep kit
(Qiagen) according the manufacturer's instructions. Weight of plasmid DNA was
determined with a ND-1000 spectrophotometer (NanoDrop Technologies Inc.,
Wilmington, DE USA), and plasmid copy number per cell was calculated by dividing
plasmid DNA content by calculated plasmid weight and by cell number in the sample.
Linearity of the measurements was tested by determining and plotting the quantity of
plasmid DNA in different volumes of the same culture (10-180 ng; R2=0.98).
3.3 Media and growth condition
Except for continuous cultures, all experiments were carried out in Luria-Bertani (LB)
medium (5 gr/l yeast extract, 10 gr/l bacto-trypton and 5 gr/l NaCl). To ensure similar
plasmid copy numbers, the lag phase was eliminated by initiating the experiments
with an actively growing culture. Unless mentioned differently, a single colony was
grown to an OD600=0.3 on LB broth supplemented with the appropriate antibiotic
(100 g/ml ampicillin, 10 g/ml tetracycline, 30 g/ml chloramphenicol, 50 g/ml
kanamycin) 37ºC (except for the temperature-sensitive mutants CAG22188 (30°C),
CAG9333 (25ºC) and their corresponding wild types), diluted 10-fold in fresh
medium and then re-grown to the desired optical density (for most experiments,
OD600=0.2).
In strains LM100 and LM1009, production of RpoD is driven by the trp promoter
(Magnusson et al., 2003). The addition of tryptophan (0.1 mg/ml) along with either
0.2 mM or 0.02 mM indol-acrylic acid (IAA) leads in this strain to either normal or
reduced production of RpoD, respectively. IAA was dissolved in acetone, the final
concentration of which in the assay mixture was always 1%. Before performing
experiments with these strains they were centrifuged (4000 rpm, 10 min; Eppendorf
centrifuge 5810 R) and re-suspended in a fresh medium containing the appropriate
IAA concentration. In strains containing plasmid pAR1, overproduction of RpoH was
achieved by adding 0.2% L-arabinose (Aviram Rasuley PhD Thesis, Tel Aviv
University).
3.4 Allele distribution by P1 coliphage (transduction)
Several mutants (Table 1) were constructed by replacing the native gene with a
defective allele by P1 transduction as described by Shively et al. (1984).
Donor P1 lystate production: the donor strain carrying a defective allele was grown
overnight at 37°C, shaken (200 rpm) in LB broth supplemented with the appropriate
antibiotics and then diluted 1:100 into fresh 10 ml LB supplemented with 5 mM
CaCl2. The culture was allowed to grow for 40 min before adding 100 l P1 coliphage
(reproduced on a wild type genetic background and kept at 4°C in a glass tube) and
than incubated at 37°C and shaken (200 rpm) for 3 hours until the culture was
completely lysed by the phage. Cells that survived the lysis were killed by adding 500
l chloroform and incubating for additional 10 min. Cell debris were removed by
centrifugation for 10 min at maximum speed and the upper phase, containing the new
phages, was collected and used for introducing the defective allele into the acceptor
strains.
Transduction: The acceptor strain was grown overnight at 37°C shaken in 200 rpm in
LB broth. The culture was centrifuged (Eppendorf centrifuge 5417 C), concentrated
x2 in 10 mM MgSO4, 5 mM CaCl2, and divided into 5 aliquots of 100 l each. Four
samples (the fifth served as a control) were then mixed with 10, 25, 40 or 100 l
donor P1 and incubated at 37°C with shaking (200 rpm) for 40 min. P1 infection was
stopped by adding 100 l 0.1 M Na3Citrate to each sample and positive transductants
were selected on LB agar plates supplemented with the appropriate antibiotics. To
remove P1 traces, positive colonies were selected and isolated on LB agar plates
containing the appropriate antibiotic.
3.5 Construction of new inactive alleles
New deletions of genes were carried out as described by Datsenko and Wanner (2000)
and by Yu et al. (2001) using the primers listed in Table 3.
Generation of PCR product: pKD13 (Datesnko and Wanner, 2000) was used as a
template DNA for PCR reaction with 60 base long primer pairs designed to amplify
the kanamycin resistance gene from the plasmid (20 bases at the 3' end of each
primer) and to undergo a recombination process with the edges of the chromosomal
target site (40 bases at the 5' side of each primer identical to the recombination sites in
the chromosome). PCR (TGradient, Biometra, USA; see details in section 3.7) was
carried out with proofreading Bio-X-Act DNA polymerase in the presence of 200 nM
template and primers using the manufacturer's reagents and instructions. The ~1500
bp product was analyzed by electrophoresis on a 1 % agarose gel, gel-purified and
desalted using QIAquick gel extraction kit (Qiagen), digested with DpnI to eliminate
template plasmid traces and desalted again using the same kit.
Preparation of recombination permissive electrocompetent cells: DY378 is a
recombination permissive strain harboring a lysogenic defective lambda phage. An
overnight culture grown at 30°C with shaking (200 rpm) in LB broth was diluted
1:100 into 100 ml fresh medium and regrown to OD600=0.6. The shaking flask was
then incubated in a 42°C pre-heated water bath for 15 min to induce the lambda PL
promoter (controlling the genes exo, beta and gam that encode recombination
promoting factors) and then cooled for 5 min on ice. Cells were washed 4 times in
cold DDW and finally resuspended in 1 ml. 100 l aliquots were mixed on ice with 50
l of purified PCR product, transferred into an electrophoration cuvette and electrical
DNA transformation was carried out at 2.5 mV (Electro Cell Manipulator ECM 935,
BTX, USA). The cuvette was then flooded with LB broth pre-warmed to 37°C,
transferred into a culture tube and grown for 1 hour. Positive recombinants were
selected on LB agar plates supplemented with the appropriate antibiotic and verified
by PCR and sequencing.
Controls: To ensure no DNA contamination, the electrical transformation was
performed without adding DNA; the original component cells were also plated to
ensure no antibiotic resistant bacterial contamination.
3.6 Chemical transformation of plasmid DNA
Plasmids were distributed into the various bacterial strains using the following
protocol:
Competent cells preparation: Single colony was grown at the appropriate temperature
(usually 37°C unless the strain is temperature sensitive; see Table 1), shaken (200
rpm) in 100 ml LB broth supplemented with the appropriate chemicals (antibiotics
and strain specific essential metabolites) to an OD600=0.5 and immediately
centrifuged in a 4°C pre-cooled centrifuge (Eppendorf centrifuge 5810R) at 4000 rpm
for 10 min. The growth medium was disposed of and the bacterial pellet was
resuspended in 50 ml cold 0.1 M CaCl2. After 30 min on ice, the culture was once
again centrifuged and resuspended in 3 ml cold mixture of 20% glycerol with 0.1 M
CaCl2. For maximum efficiency, competent cells were used immediately or either
kept in 200 l aliquots at -80°C for future use.
Chemical transformation: Competent cells were incubated on ice for 30 min and then
transferred into a 15 ml culture tube and gently mixed with 1-5 l of plasmid DNA.
The mixture was incubated on ice for 30 min, heat shocked at 42°C for 2 min, and
cooled again on ice for 2 min. Each sample was then supplemented with 0.8 ml LB
broth and allowed to grow for an hour at the appropriate temperature with shaking
(200 rpm). Positive colonies were selected on LB agar plates supplemented with the
appropriate antibiotic (usually ampiciline). Control: To ensure no antibiotic resistant
bacterial contamination, the original competent cells were similarly plated, expected
not to show any growth.
3.7 Cloning (plasmid construction)
Most of the plasmids used in this study are pDEW201 (Van Dyk and Rosson, 1998)
derivatives. Plasmids are listed in Table 2 and primers in Table 3.
Table 3: Primers that were used in this study.
Primer Name
Sequence (5' ĺ 3')
Use
Primers for gene inactivation
Transferring Pwza allele
Pwza-F
CGTTGGTGATCGCCTGTTGACCGG
from MS1651 to MG1655
Transferring Pwza allele
Pwza-R
GCCGTTCGTTCAGCTCCATCGTGG
from MS1651 to MG1655
yjbE-F
ATTTTTGCCATATCTGCGCTTGCGGCGACTT
Inactivating of yjbEFGH
CTGCGTGGGCGTGTAGGCTGGAGCTGCTT
and yjbE
C
GTCGTGGTTGTGGTGGTCCCGGTATTAGAA
Inactivating of yjbE
yjbE-R
CCATCACCGCATTCCGGGGATCCGTCGACC
TAAAGAGCTGGGCAATTCACTGTGGGACAG
Inactivating of yjbF
yjbF-F
TCTGTTCGGCGTGTAGGCTGGAGCTGCTTC
CCATGTTGGGTAACGAGAGTAGCCTGATCC
Inactivating of yjbF
yjbF-R
TGAGTGACCCATTCCGGGGATCCGTCGACC
Inactivating of yjbG
ATTTTCCGGTGAAAACCACTCTCATCAAGGC
yjbG-F
GGCAAAACAGTGTAGGCTGGAGCTG
(Yael Rozen's PhD
thesis)
Inactivating of yjbG
TGCCCAGCGCCAGTAAGCTAAGCAGATGTC
yjbG-R
TTTTTTTCATCATATGAATATCCTC
(Yael Rozen's PhD
thesis)
CCCATTGGTCCGTCGCAGTCGGATTTCGGT
yjbH-F
Inactivating of yjbH
GGCGTAGGATTATTAGTGTAGGCTGGAGCT
GCTTC
yjbH-R
CGCACGGCTGCGTGTCGGGCCAGACGAGA
Inactivating of yjbEFGH
AGAGATCCAACGGTACATTCCGGGGATCCG
and yjbH
TCGACC
Primers for cloning (pDEW201 based)
wza-F
TATACATGGAATTCCGATTAACCCGGCCCA
EcoRI
GATAGACG
wza-R
ATCAGTGGGGTACCCCCCCGCCCTCGCCTT
KpnI
TCAGCG
dfcA-F
TGATCCGAATTCTGAGAGCTGATAGAAACA
GAAGCCACTGGAGAAC
EcoRI
(From insA end)
dfcB-R
CTGAGGGGTACCTCTCTAAAATAAGAGGGC
GCACCTTTAATCCTTG
pgi-F
KpnI
(to dfcB)
TATACATGGAATTCATTTCTCCACCACGCCT
EcoRI
GCCG
RcsABoxIn-F
TATACATGGAATTCGTATTCGGCAATTAATA
EcoRI
CATAGCACG
RcsABoxOut-F
ATACATGGAATTCCGTTGTTATTTCGGAAAA
EcoRI
TACGCAG
yjbE27-R
ATCAGTGCGGTACCTGGAGATTTACCCAAA
KpnI
AACATTTCGG
yjbE76-R
ATCAGTGGGGTACCCGCCCACCTGTACAGG
KpnI
TGCAGC
Primers for relative real time PCR analysis
yjbE-F
TTTGCCATATCTGCGCTTG
yjbE-R
TGGTGGTCCCGGTATTAGAA
yjbF-F
AAGCGACCTGCACTCATTCT
yjbF-R
AGGCCAGCACCACAAATAAC
yjbG-F
TATTGTCGCGTTGCTTTTGA
yjbG-R
CCAGCTCTTCGCTAATCACC
yjbH-F
GCTTGCTACGGCGAAACATA
yjbH-R
GGGAAGAGTTGCACTGAAGC
wzc-F
CCTCGATATTGCAGTGAGCA
wzc-R
CGCTGCTCAGGGTGTAGTTT
wcaD-F
CCCATCACCATCGTCACTTT
wcaD-R
CCACACCATGCCAATAATGA
gmd-F
ATGGCGACCTGAGTGATACC
gmd-R
TTTCTTTTTCCAGACCGAGGA
wcaH-F
AGGAAGACTTTGCCACGGTA
wcaH-R
CTTCCAGCGTTTCGTCTTTC
manC-F
ATGAGCAGCACCGCTTTATT
manC-R
CCGCCAATACCAGCATTAAC
rcsA-F
CGACATTGAAACCGTTGATG
rcsA-R
GCAATTGCCATAAAAACGAT
otsA-F
GGTGAAACAGGGAATGAGGA
otsA-R
CACCAGATCGAGCCGATAAT
katE-F
GCATTCCGGAACGTATTGTT
katE-R
CCACCCTGAACGGTAGAGAA
proX-F
AGCGTACTTTTTGCGACAGC
proX-R
ACTTCGCTGGGTTTGTTGAC
osmY-F
TGCTGGCTGTAATGTTGACC
osmY-R
TCGGTGCTCTTGATGTTGTC
rpmA-F
TAACGGTCGCGATTCAGAAG
rpmA-R
GCAAACAGAGTGTGGTCACG
Primers for primer extension
PrimEx1
GCAAAAATGCCATACAGAAC
PrimEx2
ACGCAGAAGTCGCCGCAAG
PrimEx3
ACTCCCCGCCGAAACCGAC
Primers for in-vitro transcription template generation
T7A1-F
GCTTGGTTATGCCGGTACTG
T7A1-R
ATGCGACTCCTGCATTAGG
pR-F
CCGTGCGTCCTCAAGCTGCTC (Kubori and Shimamoto, 1997)
pR-R
GGGCGTAGAGGATCTGCGCCC (Kubori and Shimamoto, 1997)
yjbE-F
TAACGGAAATCATACCGTGAGG
yjbE-R
AAATGCCATACAGAACTTTTTTCATAAC
DNA amplification: MG1655 genome (extracted with DNeasy Plant Mini Kit
(Qiagen) according to the manufacturer's instructions) was used as a template for PCR
reactions. Each primer was designed to have a calculated annealing temperature of
72ºC (based on the donation of 2ºC by T or A and 4ºC by C or G) to its target site and
a tail containing the restriction enzyme digestion site and 8 mismatched nucleotides at
its far 5' end. PCR was carried out with proofreading Bio-X-Act DNA polymerase in
the presence of 200 nM template and primers using the manufacturer's reagents and
instructions. The following PCR program was used: hot start at 94ºC for 5 min, 26-33
cycles of 94ºC for 1 min, 55-60ºC (annealing) for 1 min and 72ºC for 1 min for every
1 kb of expected product length (elongation). The enzymes were then allowed to clear
from the DNA at 72ºC for 10 min and the reaction was stopped at 4ºC. PCR products
were analyzed in a 1 % agarose gel and either purified from non-specific products or
just desalted using a QIAquick gel extraction kit (Qiagen).
DNA digestion: pDEW609 (yjbF'::luxCDABE) or pDEW229 (lacZ'::luxCDABE)
were used as acceptors for the new DNA fragment after elimination of their original
insert. Four µg of DNA (plasmid and PCR product; determined using ND-1000
spectrophotometer) were digested with 10U restriction enzyme in a final volume of
100 µl containing the appropriate digestion buffer at 37ºC for 3 hours (5U of enzyme
were added in the beginning and 5U after 1.5 hours). The digested vector was then
separated from the original insert by electrophoresis and gel-purification and the
digested PCR product was desalted using QIAquick gel extraction kit (Qiagen).
For
cloning a new promoter, DNA was double digested with EcoRI and KpnI. For
replacing the luxCDABE reporting system with lacZ, DNA was digested with HindIII
which cuts within the luxC gene and right after the luxE gene.
DNA ligation: DNA ligation was carried out with T4 ligase (Roche) according to
manufacturer's instructions in a final volume of 20 µl. The linear vector was treated
with shrimp alkaline phosphatase (Fermentas) and ~200 ng vector were mixed with
insert in a molar ratios of 1:1 to 1:9 (estimated from DNA band intensity in the gel
divided by DNA length). The enzyme was added only after the DNA mixture was
supplemented with the ligation buffer and the appropriate volume of DDW, incubated
for 5 min at 65ºC and then cooled on ice. The reaction tubes were incubated for 4
hours at 16ºC followed by an overnight incubation at 4ºC.
Each ligation reaction was used for a single chemical transformation using AG1688
competent cells as describe above. Positive plasmids were selected by colony PCR
with primers designed to amplified a segment ranging from a location inside the insert
to a location outside the insert (in the vector) and by sequencing. When replacing the
luxCDABE reporting system with lacZ, positive pale blue colonies were isolated from
LB agar plates containing 100 µg/ml ampicillin, 40 g/ml X-gal and 0.5 M NaCl,
included to induce yjbF-driven LacZ activity.
3.8 Monitoring promoter induction: bioluminescence assays
Induction of promoters was routinely monitored by following the luminescence of E.
coli
strains
bearing
pDEW201-derived
plasmids
(Table
2)
harboring
a
promoter::luxCDABE (Photorhabdus luminescens luxCDABE bioluminescence
genes) transcriptional fusions. Exposure of these cells to NaCl was carried out either
in 96-well microtiter plates or in glass Erlenmeyer flasks.
All experiments were carried out in Luria-Bertani (LB broth) medium (5 g l-1 yeast
extract, 10 g l-1 bacto-trypton and 5 g l-1 NaCl). To ensure similar plasmid copy
numbers, the lag phase was eliminated by initiating the experiments with an
exponentially growing culture. A single colony was grown to an OD600=0.3-0.5 on LB
broth supplemented with the appropriate antibiotic (100 g ml-1 ampicillin, 10 g ml-1
tetracycline, 30 g ml-1 chloramphenicol, 50 g ml-1 kanamycin) at 37ºC (except for
the temperature-sensitive mutants CAG22188 (30°C), CAG9333 (25ºC) and their
corresponding wild types), diluted 10-fold in fresh medium and then re-grown to the
desired optical density (unless mentioned otherwise, OD600=0.2).
In strains LM100 and LM1009, production of RpoD is driven by the trp promoter
(Magnusson et al., 2003). The addition of tryptophan (0.1 mg ml-1) along with either
0.2 mM or 0.02 mM indol-acrylic acid (IAA) leads in this strain to either normal or
reduced production of RpoD, respectively. IAA was dissolved in acetone, the final
concentration of which in the assay mixture was always 1%. Before performing
experiments with these strains they were resuspended in a fresh medium containing
the appropriate IAA concentration.
In strains containing plasmid pAR1,
overproduction of RpoH was achieved by adding 0.2% L-arabinose, before
distributing the cells into the wells of a microtiter plate.
For microtiter plate assays, 50 µl of bacterial culture were added to the wells of an
opaque white clear bottom 96-well microtiter plate already containing 50 µl fresh LB
broth supplemented with either twice the required NaCl concentration or 0.09 M NaCl
(basal growth medium concentration). The plate was then sealed with a transparent
cover and incubated at 37ºC for 300 min. Bioluminescence (reported in RLU, the
instrument's arbitrary relative light units) and OD600 were measured at intervals of 15
min, each following a 5 sec shaking. For several hours following exposure to NaCl in
the microtiter plate, OD600 of the NaCl-exposed cells did not change.
When mentioned, the same procedure was performed in 10 ml of medium in 50 ml
Erlenmeyer flasks, incubated for 300 min at 37 ºC with shaking (200 rpm). Aliquots
(100 µl) were withdrawn at 45 min intervals, and bioluminescence and OD600 were
immediately measured in the same luminometer.
In all cases, bioluminescence data were normalized to a uniform cell density by
dividing the measured light intensity (RLU) by the OD600 values measured in the
same well at the same time point.
3.9 Monitoring promoter induction: colorimetric assays
Culture samples (300 µl) were cooled on ice and centrifuged for 4 min (10,000xg,
4ºC). The cells were resuspended in 300 µl phosphate-buffered solution (PBS; 6.85
ml 1 M Na2HPO4, 3.15 ml 1 M NaH2PO4 and 90 ml DDW; pH 7.2) supplemented
with 0.1% triton X-100. The bacterial lysate was mixed with an equal volume of PBS
containing
the
LacZ
substrate
fluorescein-di-β-D-galactopyranoside
(FDG;
Rakhmanova and McDonald, 1997; Fluka) and 100 µl aliquots were transferred in
duplicate into an opaque white 96-well microtiter plate. Fluorescence (excitation: 485
nm, emission: 510 nm) was read for 60 min at 1 min intervals using a microtiter plate
reader (Victor2, Wallac, Finland). LacZ activity, calculated as the rate of fluorescence
signal development (Rahkmanova and McDonald, 1997), is reported as the increase
per minute of the instrument's arbitrary relative fluorescence units (RFU) and
normalized to cell density by dividing by OD600.
3.10 Total RNA extraction
Single colonies of strains MG1655 and QC2410 (rpoS::Tn10) grown (37°C)
overnight on LB broth were diluted 1:100 into fresh LB broth containing 0.09 or 0.7
M NaCl and regrown for 90 and 180 min, respectively, until the wild type strain
reached an OD600=0.3 (Figure 2 A-B). The cultures were then diluted to the density
obtained by the rpoS mutant (OD600=0.15) and 50 ml of each strain were subjected to
total RNA extraction using RNAeasy Midi Kit (Qiagen) and DNase I (Qiagen)
treatment. RNA concentration was determined using ND-1000 spectrophotometer.
3.11 Whole cell DNA-microarray analysis
Twenty ng total RNA from strain MG1655 (WT) and QC2410 (rpoS::Tn10) was
reversed transcribed with poly T18 primer and mRNA expression was assayed using
Affymetrix GeneChip® E. coli Antisense Genome Array (Weizmann Institute of
Science (Rehovot, Israel) MicroArray Unit), according to the manufacturer's
instructions.
A
MG1655
(0.7 M)
0.09 M
OD600
BIOLUMINESCENCE(RLU/OD600)
1
0.1
rpoS::Tn10
(0.7 M)
0.01
0
30
60
90
120
150
180
40000
B
30000
rpoS::Tn10
(0.7 M)
20000
10000
MG1655
(0.7 M)
0.09 M
0
0
30
TIME (min)
60
90
120
150
TIME (min)
Figure 2. Whole genome microarray experiment setup. (A) Growth curves of the different
cultures (see text). Arrows indicate sampling points. (B) yjbF'::luxCDABE activity in the
different cultures. Both parameters were measured simultaneously.
180
3.12 Relative Real Time PCR analysis
Total RNA concentration was measured using a ND-1000 spectrophotometer
(NanoDrop Technologies Inc., Wilmington, DE USA). Five µg of total RNA from
either the wild type (MG1655) or the rpoS mutant (QC2410) were subjected to
reverse transcription carried out with a T18 primer using RevertAid First Strand cDNA
Synthesis Kit (Fermentas). Twenty five ng of cDNA were then mixed with Sybrgreen
reaction mixture (ABI) and assayed with ABI prism 7000 for the relative
quantification of the abundance of each of the following genes, in both strains: yjbE,
yjbF, yjbG, yjbH, wzc, gmd, manC, wcaH, wcaD, and rcsA. Five additional genes that
were similarly amplified were proX, osmY, katE and otsA, RpoS-dependent genes that
served as a negative control and rpmA, as housekeeping gene that served as a
reference. The latter gene yielded average Ct values (n=3) of 17.8 ± 0.7 and 17.5 ± 0.6
in MG1655 and QC2410, respectively. Reaction conditions were as follows, 50 ºC for
2 min, hot start at 95°C for 10 min, and 40 cycles of 95ºC for 15 sec and 60ºC for 1
min. A dissociation test was performed at the end of each run to ensure a single PCR
product in each reaction. Results were obtained with the SDS analysis program (ABI)
using a relative quantification algorithm with automatic Ct calculation. RQ (relative
RNA quantification) was calculated for each experiment repeat as follow:
RQ = 2–Ct
Where
And
Ct = Ct (studied gene) – Ct (rpmA)
Ct = Ct (rpoS::Tn10) – Ct (MG1655)
(RpoS-dependency calculation)
Or
Ct = [averaged Ct (wzc, gmd, manC, wcaH, wcaD)] –
[averaged Ct (yjbE, yjbF, yjbG, yjbH)]
(Relative expression of wca genes in comparison to yjb genes calculation)
3.13 Primer extension
An overnight MG1655 culture was re-grown in LB broth to OD600=0.2 and then
divided into two subcultures of 100 ml each. Total RNA was extracted from both
portions, either at time zero or after the induction of the yjbEFGH operon by 150 min
incubation in the presence of 0.7 M NaCl (37°C, 200 rpm, LB). Total RNA was
isolated using an Ultraspec RNA kit according to the manufacturer's (BIOTECX
Laboratories) instructions. The RNA samples (30 g) were subjected to primer
extension at 42°C for 45 min, using avian myeloblastosis virus reverse transcriptase
(CHIMWEx) and 3 end-labeled prime (see Table 3). The extension products and the
sequencing reaction mixtures primed with PrimEx1 end-labeled primer were
separated on a 6% sequencing gel.
3.14 In-vitro transcription assays
DNA templates were prepared by PCR, using the MG1655 genome as a template for
amplifying the yjbE promoters region and linearized pAR1345 and pPR1AL73 for
amplifying T7A1 and pR promoters respectively (Kubori and Shimamoto, 1997),
using the primers listed in Table 3.
RNA polymerase holoenzyme (RNAP-RpoD; 0.2 µM) was mixed with DNA template
(0.50 pmol yjbE; 0.15 pmol T7A1 and pR) and pre-incubated for 10 min at 37°C in 8
µl of T-buffer (50 mM Tris-HCl (pH 7.9), 100 mM KCl, 10 mM MgCl2, 1 mM DTT
and 150 µg ml−1 casein). Transcription was started by adding substrates: 20 µM [32
P]-GTP or [-32P]-ATP (0.1 µCi µl-1) and 350 µM of each of the other three NTPs.
Single (T7A1 and pR) and multi (yjbE) round transcriptions were allowed to proceed
for 20 and 60 min in the presence or absence of 0.5 mg ml−1 heparin, respectively.
The reaction was quenched with phenol. The transcripts were analyzed by
electrophoresis on a 20% polyacrylamide sequencing gel.
3.15 Bacterial growth - continuous culture
Chemostat cultures were grown in stirred (800 rpm) glass bioreactors (2 liter total
volume, 550 ml washing volume). Temperature was maintained at 37°C by
submerging the bioreactor in a temperature controlled water bath. Aeration (1.4 l/min)
was provided by pumping in filter-sterilized air, and the pH was kept at either 6.8-6.9
(defined minimal medium) or 8.4-8.6 (complex rich medium). Dilution rate (0.1, 0.2,
0.3, 0.5 or 1.0 h-1) was controlled by a peristaltic pump.
Two media were used: a rich complex medium (LB) and a minimal medium (ClimBa,
Ihssen and Egli, 2004). The latter medium contained 12.8 g Na2HPO4 · 2 H2O, 3.0 g
KH2PO4 and 1.77 g (NH4)2SO4 per liter. The following salts and trace elements were
added as a 100-fold concentrated solution after autoclaving (final concentrations are
given per liter): 130 mg MgCl2 · 6 H2O, 80 mg CaCO3, 77 mg FeCl3 · 6 H2O, 11 mg
MnCl2 · 4 H2O, 1.5 mg CuSO4 · 5 H2O, 1.3 mg CoCl2 · 6 H2O, 4 mg ZnO, 1.2 mg
H3BO3, 10 mg NaMoO4 · 2 H2O and 790 mg EDTA Na4 · 2H2O. Glucose (1 g/l) was
used as the sole carbon source. Glucose, salts and trace elements stock solutions were
autoclaved separately and added after cooling.
Ten ml of LB broth were inoculated with a single colony and after 2 h at 37°C were
transferred into the chemostat bioreactor. Samples were withdrawn after steady state
(constant optical density) was reached, following 5-10 volume changes; for dilution
rates of 1, 0.5, 0.3, 0.2 and 0.1 h-1, this occurred 15, 24, 36, 48 or 96 h after
inoculation, respectively. The samples were serially double diluted six times in 96-
well opaque white clear bottom microtiter plates (100 µl final volume) and assayed
for yjbF'::luxCDABE activity as describe above.
3.16 Modifying aeration regime and gas exchange capacity
The effects of aeration and gas exchange rates on gene induction were assayed in both
batch and continuous cultures. In batch mode, different media volumes (50, 100, 150,
200 and 300 ml) were used, in a single volume (300 ml) flask. An overnight culture (3
ml, originating from an overnight colony, grown at 37ºC) was diluted 1:100 into fresh
LB broth, re-grown at 37ºC to an OD600 of 0.8, and then mixed with an equal volume
of LB supplemented with 1.4 M NaCl. For assaying induction of the lacZ gene
promoter, isopropyl -D-1-thiogalactopyranoside (IPTG, 0.5 mM) was also added.
Different volumes of the culture, now containing 0.7 M NaCl, were distributed into
the flasks and incubated with shaking (200 rpm) at 37ºC. Where noted, the media
were aerated with a filtered airflow (200 ml/min), through a glass pipette submerged
to the bottom of the flask. Sample aliquots were withdrawn every 45 min or after 200
min and assayed for luciferase (MG1655, pDEW609/229) or -galactosidase
(MC4100, pMLZ609) activity respectively, as described above.
In chemostat experiments, cultures were allowed to reach steady state in ClimBa under
different aeration regimes (determined by rotor speed and air pumping rate) and then
subjected to osmotic up-shift (0.7 M NaCl) by adding an appropriate volume of
ClimBa medium supplemented with 5 M NaCl without micronutrients and C-source.
Aliquots (1 ml) were withdrawn at 30 min intervals, and bioluminescence and OD600
were immediately measured in the same luminometer. Dissolved oxygen
concentrations were measured using a Yellow Springs Instruments Incorporated
(Yellow Springs, OH, USA) Dissolved Oxygen Meter model 58.
3.17 EPS-related phenotype characterization
Single colonies were spread on LB-agar plates supplemented with 0.09 or 0.5 M NaCl
with or without 150 µg/ml Congo-red (CR; 37, 8) and allowed to grow for 24 h at 37
ºC. Plates were photographed over a black background (- CR) or a light screen (+ CR)
with an Olympus Stylus-770 SW digital camera. A mucoid appearance indicated
overproduction of CA-EPS by the wca operon; red-stained cells indicated presence of
the EPS driven by the yjb operon.
3.18 Light/fluorescence microscopy
Ten µl cells were mounted on microscopic slide, covered with a cover slip and
analyzed with epifluorescence microscope (filter sets 13 and 20, Axiovert 135TV,
Zeiss, Germany).
3.19 Characterization of bacterial pellet volume formed upon centrifugation
To measure small differences in the volume of pellets formed upon centrifugation of
an equal number of cells, a special centrifuge tube was designed consisting of 2
connected chambers (Figure 3): an upper 12 ml space opening into a lower 35 l (0.5
mm wide) capillary. The tube was designed to fit into a 50 ml plastic test-tube (29 X
115 mm) which served as an adaptor for the swing bucket rotor of the centrifuge
(Eppendorf 5810 R). Centrifuged cells sink to the bottom of the upper chamber and
are forced into the capillary so that the height they fill is proportional to the volume
they occupy. This height was measured at a 0.5 mm resolution.
Calibration tests show that the pellet volume correlated linearly with different
volumes (Figure 4 A) or dilutions (Figure 4 B) of the culture (centrifugation force is
maximal, 4000 rpm = 3220 g). The pellet volume was not influenced by
centrifugation force (201 g – 3220 g; Figure 4 C) but centrifugation time below 10
min was not long enough to obtain final results (Figure 4 D).
Load
Sample
Insert
50
mL
50
mL
40
40
30
30
20
20
10
10
Centrifuge
Bacterial
Pellet
Figure 3: A centrifuge tube for measuring small differences in the volume of pellets formed
upon centrifugation. The tube is shown on the left with the 50 ml falcon tube it made to fill in.
Medium is shown in orange and the cells in yellow.
45
30
A
rpoS::Tn10
PELLET HEIGHT (mm)
PELLET HEIGHT (mm)
B
40
25
ǻyjbG rpoS::Tn10
20
15
10
5
35
30
25
20
15
10
5
0
0
2
4
6
8
10
12
0
14
0.25
SAMPLE VOLUME (ML)
0.75
1
30
40
SAMPLE DILUTION
12
40
C
D
10
PELLET HEIGHT (mm)
PELLET HEIGHT (mm)
0.5
8
6
4
2
0
30
20
10
0
1000
2000
3000
4000
0
CENTRIFUGE FORCE (G)
10
20
CENTRIFUGE TIME (MIN)
Figure 4: Calibration tests of the system. rpoS::Tn10 and its descendant yjbG rpoS::Tn10
mutants were grown in LB supplemented with 0.7 M NaCl to OD600=0.3 and subjected to
centrifugation in the tube shown in Figure 3, under different conditions. (A) Different culture
volume; (B) Different culture dilution; (C) Different centrifugation force; (D) Different
centrifugation time.
3.20 Buoyancy assay
Five ml were stained with SYTO-9 nucleic acid stain solution (LIVE/DEAD®
BacLightTM, Molecular Probes, L-13152) and then underlaid with 2 ml 55% Percoll
(Sigma), 25 mM phosphate buffer, pH 6.5) as described by Peleg et al. (2005) in 15
ml plastic test-tubes and centrifuged at 3,220 x g at room temperature for 10 min.
Migration of the culture in the Percoll gradient was digitally photographed above a
UV light table.
3.21 Total carbohydrates determination
Overnight cultures were diluted 1:100 into fresh LB broth supplemented with 0.7 M
NaCl and re-grown to OD600=0.3. Ten ml were then centrifuged (10 min, 4000 rpm)
and washed with 1 ml saline solution to eliminate all traces of residual LB broth.
For total carbohydrates determination, the following analytical protocols were adapted
from Yuval et al. (1998): the sample was centrifuged (10 min, 8000 rpm) and washed
with 100 l 2% Na2SO4. The cells were extracted by adding 400 l DDW and then
800 l chloroform, vigorously mixed and separated by centrifugation (14000 rpm, 10
min). 300 l of the upper (aqueous) phase were mixed with 200 l DDW and with 1
ml anthrone reagent (Sigma) dissolved in 100% H2SO4 (1 mg ml-1). The mixture was
incubated at 90°C for 10 min and the OD630 of the sample was measured. Total
carbohydrates content was calculated from a trehalose (Sigma) calibration curve.
4. RESULTS
4.1 EPSs synthesis in response to osmotic shock
4.1.1 The basic phenomenon: yjbEFGH is induced by elevated osmotic pressure
It was previously demonstrated (Rozen et al., 2001), using a yjbF'::luxCDABE
transcriptional fusion, that yjbEFGH is induced upon a shift from LB broth into
artificial seawater as a consequence of osmotic stress. Osmotic induction of this gene
can be demonstrated by both organic (sucrose; Figure 5 A) and inorganic (NaCl;
Figure 5 B) solutes, but the latter exerts much stronger effects at similar osmolarity;
NaCl was thus subsequently used to induce this promoter. As may be observed from
the kinetics of bioluminescence emission obtained using this reporter fusion,
yjbEFGH induction at the lowest effective NaCl concentration (0.3 M, which
corresponds to 0.6 osmolar) was apparent 20 min after exposure to NaCl, and reached
a maximum after approximately an hour. In the presence of the higher NaCl
concentrations the onset of the induction and its peak were both delayed. The dose
dependency of the response is displayed in Figure 5 C, which also demonstrates the
negative effect a functional rpoS has upon the system.
sucrose (OSMOLAR)
BIOLUMINESCENCE (RLU/OD600)
1400
A
1200
1000
800
0.8
600
0.6
400
0.4
200
0.2
0
0
50
100
150
200
250
300
TIME (min)
NaCl
BIOLUMINESCENCE (RLU/OD600)
14000
(OSMOLAR)
1.4
B
12000
1.2
10000
1
8000
0.8
6000
0.6
4000
0.4
2000
0.2
0
0
50
100
150
200
250
300
MAX.BIOLUMINESCENCE (RLU/OD600)
TIME (min)
50000
C
MG1655
rpoS::Tn10
45000
40000
35000
30000
25000
20000
15000
10000
5000
0
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1
NaCl (M)
Figure 5: Activity of the yjbF'::luxCDABE transcriptional fusion in response to osmotic stress
induced by sucrose (A) and NaCl (B). The dose dependency of the response to NaCl
(maximal luminescence at each salt concentration) is shown in MG1655 and rpoS::Tn10
mutant (C).
4.1.2 Screen for yjb like regulated genes
To identify additional genes that may be co-regulated with yjbEFGH I have screened
for genes that share the two main regulatory features of this operon: induction by
osmotic shock in the wild type strain and overinduction in rpoS::Tn10 mutant.
For this purpose, the MG1655 wild type strain and its rpoS::Tn10 derivative were
both grown in LB supplemented with either 0.09 or 0.7 M NaCl, until the wild type
cultures reached OD600=0.3 (Figure 2 A). All samples were then diluted to the same
optical density and subjected to total RNA extraction followed by whole cell DNAmicroarray analysis. Sampling times were chosen based on the activity of the
yjbF'::luxCDABE transcriptional fusion in each strain in each NaCl dose (Figure 2 B);
After 180 min, it was an order of a magnitude higher in the rpoS::Tn10 mutant than in
the wild type, both grown in LB broth supplemented with 0.7 M NaCl (Figure 2 B).
Among 1138 genes that demonstrated 2-fold overexpression in the wild type upon
growth in LB broth supplemented with 0.7 M NaCl in comparison to growth in basal
LB medium, only 187 also displayed 1.5-fold increase expression in the background
of rpoS::Tn10 in comparison to the wild type (supplementary Table 1; functional
classification is presented in Supplementery Figure 1). This list included, as expected,
3 members of the yjb operon, yjbE, yjbF and yjbH, which displayed osmotic shock
response ratios of 33 (result of a very low background), 3.9 and 2.2 and RpoSresponse ratios of 1.7, 1.6 and 1.7, respectively (Table 4). Also selected by this double
criterion were 13 out of the 21 known genes of the wca operon, displaying an osmotic
shock response ratio of 3.2-27 and RpoS-response ratio of 1.5-2.8 (Table 4). A few
additional RcsC-dependent genes, as defined by Ferrièrs and Clarke (2003), also
behaved similarly, including osmB, rcsA, ugd, ykfE, ygaC, ymgD, yhaL and yhaK. As
already reported by Goller et al. (2006), another functional EPS production operon,
pgaABCD, is also induced by osmotic stress. In our experiment it displayed an
osmotic shock response ratio of 6.1, 5.8, 1.6 and 2.6 respectively but showed no
dependency upon RpoS. The yjb paralogue dfc-pseudo-operon (composed of
gfcABCDE and etk; Peleg et al., 2005) showed neither an induction by osmotic stress
nor an RpoS-dependency.
Table 4: Expression ratios obtained by DNA-microarray analysis of gene-members of various
EPS production related genes following growth in LB supplemented with 0.7 or 0.09 M NaCl,
in rpoS+ and rpoS- backgrounds.
GENE
NaCl RESPONSE RATIO (MG1655)
rpoS::Tn10 /MG1655
yjbE
33.2
1.7
yjbF
3.9
1.6
yjbG
1.9
1.8
yjbH
2.2
1.7
wza
5
2.7
wzb
3.8
1.7
wcaA
3.1
2.1
wcaB
9
2.7
wcaC
8.8
1.9
wcaD
27
2.6
yjb operon
wca operon
4
1.8
gmd
5.1
1.6
wcaH
5.7
2.2
wcaI
2.6
1.7
wzxC
4.3
1.8
wcaK
4
1.5
wcaM
3.2
1.6
rcsA
5.9
1.9
Transcription activator of wca operon
pgaA
6.1
0.8
pga operon
pgaB
5.9
0.9
pgaC
1.7
1.2
pgaD
2.6
1.1
gfcA
1
1.6
gfcB
1.7
1.1
gfcC
1.9
1.5
gfcD
1.1
1.3
gfcE
1.1
1.5
etp
1.7
1.1
osmY
6.3
0.3
wcaG
gfc operon
RpoS-dependent
The apparent similar induction characteristics of yjb and wca (osmotic stress and
RpoS dependency) were confirmed by relative real time PCR analysis (Figure 6),
applying the same sample preparation procedure employed for the DNA-microarray
analysis. Four RpoS-dependent osmotic shock induced genes, proX, osmY, katE and
otsA served here as a negative control. Indeed, as shown in Figure 6, all tested genes
(four members of the yjb operon, five members of the wca operon and their common
regulator rcsA) except for the 38-dependnet genes displayed 38-response ratios
ranging from 4 to 8 confirming the DNA-microarray results.
8
7
6
5
4
3
2
1
0
otsA
katE
osmY
proX
manC
wcaH
gmd
wcaD
wzc
rcsA
yjbH
yjbG
yjbF
-1
yjbE
EXPRESSION RATIO (rpoS::Tn10 /MG1655)
9
Figure 6: Expression ratios (rpoS::Tn10/MG1655), as obtained by relative real time analysis,
of members of the yjb and wca operons, following growth in LB broth supplemented with 0.7
M NaCl. proX, osmY, katE and otsA are RpoS dependent genes that serve here as a control.
4.1.3 The yjb and wca promoters display similar induction patterns
To study the relationship between the EPS-biosynthesis operons in E. coli K12, we
have assembled a set of promoter::luxCDABE transcriptional fusions harboring
promoter-containing segments from three E. coli EPS systems: yjbF'::luxCDABE,
wza':;luxCDABE (Stout, 1996; the first gene of the wca operon) and
gfcB'::luxCDABE (Peleg et al., 2005; a paralogue operon of yjb). As before, the
RpoS-dependent promoter of osmY (osmY'::luxCDABE; Van-Dyk and Rosson, 1998;
Van-Dyk et al., 2001) served as the control. Activities of all transcriptional fusions
were assayed in response to osmotic shock, with an emphasis on their RpoS
dependency. The results are displayed in Figure 7 A-H. As is clearly evident from the
first 4 panels (A-D) of Figure 7, the yjb and wca promoters exhibit an identical pattern
of induction in response to three different NaCl concentrations, both in the wild type
and in the rpoS::Tn10 mutant; furthermore, activation of both promoters is similarly
enhanced by the rpoS::Tn10 mutation and peaked at the same NaCl concentration (0.7
M; Figure 8 A). The only difference observed in the induction of the two promoters is
the intensity, with the wza'::luxCDABE fusion displaying an approximately 5-fold
higher activity. The same was true also for the relative real time PCR analysis (Figure
8 B); the averaged relative expression of the five wca representative genes was 5times higher than the averaged relative expression of the four yjb genes. The dfcpromoter showed no activity at all (Figure 7 E-F); as expected, the osmY promoter
exhibited a strong positive response to osmotic shock, but its activity was strongly
inhibited in the rpoS::Tn10 mutant (Figure 7 G-H).
A
NaCl (M)
16000
0.7 M
14000
0.5 M
12000
0.3 M
0.085 M
10000
BIOLUMINESCENCE (RLU/OD600)
BIOLUMINESCENCE (RLU/OD600)
WT yjbF’::luxCDABE
18000
8000
6000
4000
2000
0
0
50
100
150
200
250
rpoS yjbF’::luxCDABE
35000
B
30000
25000
20000
15000
10000
5000
0
300
0
50
100
70000
WT wza’::luxCDABE
C
60000
50000
40000
30000
20000
10000
0
0
50
100
150
200
250
D
120000
90000
60000
30000
0
0
WT gfcB’::luxCDABE
BIOLUMINESCENCE (RLU/OD600)
BIOLUMINESCENCE (RLU/OD 600)
300
150000
300
E
300
250
200
150
100
50
0
50
100
150
200
250
300
rpoS gfcB’::luxCDABE
400
F
350
300
250
200
150
100
50
0
0
50
100
150
200
250
300
0
50
TIME (min)
WT osmY’::luxCDABE
G
120000
100
150
200
250
300
200
250
300
TIME (min)
BIOLUMINESCENCE (RLU/OD600)
BIOLUMINESCENCE (RLU/OD600)
250
TIME (min)
350
140000
200
rpoS wza’::luxCDABE
180000
TIME (min)
400
150
TIME (min)
BIOLUMINESCENCE (RLU/OD600)
BIOLUMINESCENCE (RLU/OD600)
TIME (min)
100000
80000
60000
40000
20000
0
1600
rpoS osmY’::luxCDABE
H
1400
1200
1000
800
600
400
200
0
0
50
100
150
200
250
300
0
TIME (min)
50
100
150
TIME (min)
Figure 7: Activity of yjbF' (A-B), wza' (C-D), gfcB' (E-F) and osmY' (G-H) - ::luxCDABE
transcriptional fusions in response to various NaCl concentrations in MG1655 (A, C, E and G)
and rpoS::Tn10 mutant (B, D, F and H).
MAX.BIOLUMUNESCENCE (RLU/OD600)
200000
A
MG1655 yjbF'::luxCDABE
180000
rpoS::Tn10 yjbF'::luxCDABE
160000
MG1655 wza'::luxCDABE
140000
rpoS::Tn10 wza'::luxCDABE
120000
100000
80000
60000
40000
20000
0
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
1
NaCl (M)
INDUCTION or EXPRESSION RATIO
(wca/yjb )
8
B
RT-PCR
7
Transcriptional fusion
6
5
4
3
2
1
0
MG1655
rpoS::Tn10
STRAIN
Figure 8: Comparison between the NaCl-dose dependency of yjbF'::luxCDABE and
wza'::luxCDABE, in MG1655 and rpoS::Tn10 mutant (A). The transcriptional fusions activity
ratios as obtained by bioluminescence measurement (wza':;lux/yjbF'::lux) and average
expression ratios as obtained by relative real time PCR analysis (wca genes/yjb genes), in
MG1655 and rpoS::Tn10 mutant, both performed in 0.7 M NaCl (B).
The yjb operon was listed by Ferrièrs and Clarke (2003) among genes controlled by
RcsC. The same authors (Ferrièrs et al., 2007) also pointed out that a sequence highly
similar to the RcsAB Box (Wehland and Bernhard, 2000) is located in the yjbE
promoter region. We confirmed these findings by assessing the osmotic (0.7 M NaCl)
induction and activity of the yjbF'::luxCDABE and wza'::luxCDABE transcriptional
fusions in rcsA, rcsB and rcsC inactive strains (Figure 9). The experiments were
conducted at two temperatures: 27oC and 37oC, as it was previously shown (Sledjeski
and Gottesman, 1996) that the wca operon is more active at the lower temperature. As
before, induction of both promoters exhibited a very similar pattern but a different
intensity. Both were almost completely inhibited in the rcsC and rcsB mutants, and
both displayed higher activity at the lower temperature in the wild type. In the rcsA
mutant, inhibition of activity was temperature dependant: it was significant (74 [wca]
and 49 [yjb]-fold reduction) at 37 ºC and relatively minor (4.2 [wca] and 2.8 [yjb]-fold
reduction) at 26ºC.
MAX.BIOLUMINESCENCE (RLU/OD600)
1000000
27°C wza'::luxCDABE
37°C wza'::luxCDABE
27°C yjbF'::luxCDABE
37°C yjbF'::luxCDABE
100000
10000
1000
100
WT
rcsA
rcsB
rcsC
GENETIC BACKGROUND
Figure 9: Induction of yjbF'::luxCDABE and wza'::luxCDABE transcriptional fusions, in
response to 0.7 M NaCl, in rcsA, rcsB and rcsC mutants and their ancestral wild type, at 27oC
and 37oC.
4.1.4 Overproduction of EPS in the background of RpoS inactive strains
CA-EPS production (mucoid appearance) was unnoticeable in wild type colonies
grown on LB-agar plates supplemented with either 0.09 or 0.5 M NaCl (Figure 10);
however, in 0.5 M NaCl, the rpoS::Tn10 mutant produced moderate mucoidity.
Inactivation of the wca promoter (Pwza rpoS::Tn10; Figure 10) appeared to
somewhat limit growth but clearly abolished mucoidity. The same was true for the yjb
dependent EPS production; unnoticeable in the wild type (Figure 11 A-C) but the
rpoS::Tn10 strain grown on LB-agar plates supplemented with 0.5 M NaCl and
congo-red staining dye was colored in red (Figure 11 D and F). Inactivating the yjb
operon (yjbEFGH rpoS::Tn10; Figure 11 E) suppressed this phenotype and in
addition gave rise to extreme mucoidity. These results demonstrate that the mucoid
appearance stemmed from CA-EPS overproduction by the wca operon and that the
red staining, stemmed from yjb dependent EPS overproduction in the rpoS inactive
strain.
-NaCl
+NaCl
WT
ǻyjbEFGH
WT
ǻyjbEFGH
ǻPwza
rpoS::Tn10
rpoS::Tn10
ǻPwza
rpoS::Tn10
rpoS::Tn10
ǻyjbEFGH
rpoS::Tn10
ǻyjbEFGH
rpoS::Tn10
Figure 10: Colony appearance upon growth on LB broth agar plates supplemented with 0.09
M (left) or 0.5 M (right) NaCl.
Figure 11: Congo red staining of cultures grown on LB broth agar plates supplemented with
0.5 M NaCl.
(A) MG1655; (B) yjbEFGH; (C) Pwza; (D) rpoS::Tn10; (E) yjbEFGH
rpoS::Tn10; (F) Pwza rpoS::Tn10.
4.1.5 Phenotypes in ǻyjb rpoS::Tn10 double mutants
Upon deletion of yjbEFGH, the rpoS::Tn10 strain not only lost the red stain described
above, but also became extremely mucoid (Figures 10 and 11 E) and filamentous
(Figure 12 B). These phenotypes were no longer dependent upon osmotic shock and
were observed also during growth on regular LB agar plates (0.09 M NaCl; Figure 10
left). This observation indicates that the yjb operon has a crucial role in a 38 deficient
strain, but not in a wild type strain, which affects EPS production and cell division.
B
A
10 ȝm
Figure 12: Epifluorescence microscopy of gfp-tagged parental strains (A) and their
yjbEFGH rpoS::Tn10 (B) mutant.
While deletion of either yjbE or yjbH in a rpoS::Tn10 background resulted in similar
phenotypes, inactivation of yjbF resulted only with mucoid phenotype and
inactivation of yjbG with none (Figure 13 A). When this yjbG rpoS::Tn10 double
mutant was subjected to growth in a hyper saline medium (liquid LB broth
supplemented with 0.5 M NaCl), it produced tripled amount of carbohydrates
(determined by colorimetric assay with anthrone reagent; Figure 13 B) in comparison
to the wild type and the yjbG mutant and almost twice than the rpoS::Tn10 strain.
Nevertheless, upon growth on LB agar plates supplemented with 0.5 M NaCl (not
shown) it did not exceed the moderate mucoid appearance of the its ancestral rpoS
mutant and it could not be distinguished from its parental strains by a buoyancy assay
in percoll density gradients (Figure 13 C).
A
ǻyjbE
rpoS::Tn10
ǻyjbF
rpoS::Tn10
ǻyjbH
rpoS::Tn10
9
TOTAL CARBOHYDRATES (mg/5*10 cells)
ǻyjbG
rpoS::Tn10
0.9
0.8
B
0.7
0.6
0.5
0.4
0.3
0.2
0.1
0
MG1655
C
rpoS::Tn10
ǻyjbG
ǻyjbG
rpoS::Tn10
STRAIN
Figure 13: (A) Appearance of yjbE rpoS::Tn10, yjbF rpoS::Tn10, yjbG rpoS::Tn10 and
yjbH rpoS::Tn10 double mutants, upon growth on LB agar plates. (B) Total carbohydrates
content and buoyancy properties of yjbG rpoS::Tn10 mutant and its parental strain grown in
LB broth supplemented with 0.7 M NaCl (C).
4.1.6 A novel method for EPS quantification
The overproduction of EPS in response to osmotic stress in the yjbG rpoS::Tn10
mutant was accompanied with a bulkier and less dense centrifugation pellet. To
quantify this phenomenon we have constructed a special centrifugation tube that
allowed distinguishing between small differences in pellet volume (see Materials and
Methods; Figure 3). In concept, this method was adapted from the capillary
hematocrit assay designed to evaluate red blood cells level in plasma but was
designed to contain a larger volume of medium and hence, to visualize large numbers
of small cells. As shown in Figure 14 A-B, the pellet formed by the yjbG
rpoS::Tn10 mutant was 10-fold higher than that formed by the same amount of wild
type cells (~ 5*107 cells/ml; determined by plating); the rpoS mutant yielded an
interim value; the yjbG mutant and the wild type displayed similar pellets height
although the mutant seemed to be less dense. The phenotype was completely
abolished upon the inactivation of rcsA or wza promoter (Pwza) in the rpoS::Tn10 or
the yjbG rpoS::Tn10 strains (Figure 14 B).
Although the yjbG rpoS::Tn10 mutant does not form mucoid colonies like the other
yjb rpoS::Tn10 strains, it exhibits a related phenotype, expressed only upon
exposure to osmotic stress. The phenotypic differentiation of the yjb gene members
deletions suggests that yjbG deficiency is either dispensable or can be largely
compensated for by another protein.
MG1655
yjbG
yjbG
rpoS::Tn10 rpoS::Tn10
A
7.7*107
Cells/ml
6*107
5.2*107
5.4*107
30
B
LB + NaCl
LB - NaCl
PELLET HEIGHT (mm)
25
20
15
10
5
0
MG1655
ǻyjbG
rpoS::Tn10
ǻyjbG
rpoS::Tn10
rpoS::Tn10
ǻpwza
ǻrcsA
ǻyjbG
ǻyjbG
rpoS::Tn10
ǻrcsA
ǻpwza
rpoS::Tn10
rpoS::Tn10
STRAIN
Figure 14: The pellets formed by the yjbG rpoS::Tn10 mutant and its parental strains grown
in LB broth supplemented with 0.7 M NaCl (A) and quantification of these pellets heights (B)
(LB + NaCl = 0.7 M; LB – NaCl = 0.09 M).
4.1.7 Activity of the yjb and wca promoters is enhanced in the ǻyjbEFGH
rpoS::Tn10 mutant
In order to explain the NaCl-independent mucoid colony appearance exhibited by the
yjbEFGH rpoS::Tn10 mutant, activities of both promoters (yjb and wca) were
assayed in yjb+ and yjb strains in rpoS+ or rpoS- genetic backgrounds. As might be
expected, activities of both promoters were over 10 (wca) and surprisingly 100 (yjb)fold higher in the yjbEFGH rpoS::Tn10 double mutant in comparison to their
rpoS::Tn10 ancestor (Figure 15 A-B, time point -30 min). When the NaCl
concentration was elevated, the high background activities of both promoters were
similarly reduced (Figure 15 A-B, time point 0 min). Activity of both promoters in the
yjbEFGH rpoS::Tn10 mutant was restored during 300 min of incubation but with
different NaCl dose sensitivity in comparison to their ancestral strains: the yjb
promoter was significantly induced above background activity only in 0.3 M NaCl
and was repressed at higher concentrations (Figure 16 A); the wca promoter exhibited
only modest sensitivity to elevated NaCl concentration (Figure 16 B).
These results indicate the yjb operon affects the transcription of EPSs synthesis genes
in a 38 deficient strain in an osmotic stress independent manner. It cannot be
determined if this overinduction is a result of a regulatory role fulfilled by the yjb
encoded proteins or a consequence of the yjb dependent EPS deficiency.
A
100000
10000
1000
rpoS::Tn10
ǻyjbEFGH rpoS::Tn10
100
-60
0
60
120
180
240
1000000
BIOLUMINESCENCE (RLU/OD600)
BIOLUMINESCENCE (RLU/OD600)
1000000
B
100000
10000
1000
rpoS::Tn10
ǻyjbEFGH rpoS::Tn10
100
300
-60
0
60
120
180
240
300
TIME (min)
TIME (min)
Figure 15: Kinetics of bioluminescence emission of rpoS::Tn10 and yjbEFGH rpoS::Tn10
mutants harboring yjbF'::luxCDABE (A) and wza'::luxCDABE (B) transcriptional fusions,
10000000
MAX.BIOLUMINESCENCE (RLU/OD600)
MAX.BIOLUMINESCENCE (RLU/OD600)
before (-30 min) and following exposure to 0.7 M NaCl (indicated by the arrows; 0 min).
A
1000000
100000
10000
1000
1000000
B
100000
10000
MG1655
ǻyjbEFGH
1000
rpoS::Tn10
ǻyjbEFGH rpoS::Tn10
100
100
0
0.2
0.4
0.6
0.8
0
NaCl (M)
0.2
0.4
0.6
NaCl (M)
Figure 16: NaCl dose-dependency of the activity of the yjbF'::luxCDABE (A) and
wza'::luxCDABE (B) transcriptional fusions, in yjbEFGH rpoS::Tn10 mutant and its parental
strains.
0.8
4.1.8 Suppression of the ǻyjbE rpoS::Tn10 mutant mucoid and filamentous
phenotypes
Plasmid pDEW/yjbF (pDEW609; yjbF'::luxCDABE), containing the entire yjbE gene,
was used to suppress the phenotypes exhibited by the yjbE rpoS::Tn10 mutant,
which are identical to those exhibited by the yjbEFGH rpoS::Tn10 mutant.
Introducing pDEW/yjbF into this strain cured the culture from its mucoid colony
appearance (Figure 17 A) and cell filamentation (Figure 17 B). Surprisingly, these
phenotypes were also suppressed by plasmid pDEW/wza, which contains only part of
the wza gene, but not by the original vector (not shown) or the plasmid pDEW/lacZ
(pDEW229; lacZ'::luxCDABE). In accordance with these results, the activity of both
transcriptional fusions, yjbF'::luxCDABE and wza'::luxCDABE decreased to
rpoS::Tn10 levels in the yjbE rpoS::Tn10 mutant (Figure 17 C).
The suppression by wza' cannot be simply a result of a titration of the regulatory
factors from the endogenous wca promoter by the heterologous wca promoter located
on the moderate copy number (~30 cell-1) plasmid pDEW/wza, since it did not
suppress yjbEFGH rpoS::Tn10 mutant phenotypes (not shown). The reason,
presently unknown, should be related to a function encoded on the wza' fragment
cloned in the plasmid.
A
pDEW/yjbF
pDEW/wza
pDEW/lacZ
B
pDEW/lacZ
pDEW/yjbF
pDEW/wza
MAX.BIOLUMINESCENCE (RLU/OD 600)
350000
C
rpoS::Tn10; yjbF'::lux
300000
ǻyjbE rpoS::Tn10; yjbF'::lux
rpoS::Tn10; wza'::lux
250000
ǻyjbE rpoS::Tn10; wza'::lux
200000
150000
100000
50000
0
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
NaCl (M)
Figure 17: Suppression of the mucoid (A) and filamentous (B) phenotypes and of yjbE
rpoS::Tn10 mutant by yjbE and wza'. lacZ' served here as a control. (C) Activity of
yjbF'::luxCDABE and wza'::luxCDABE in rpoS::Tn10 and yjbE rpoS::Tn10 mutants in
response to 0.7 M NaCl.
4.2 Characterization of yjb promote region
4.2.1 Which sigma factor(s) control yjb operon induction?
As was demonstrated already, RpoS represses yjbEFGH expression. This observation
has prompted us to ask for the sigma factor that activates the yjb promoter. In view of
the emerging heterogeneity in sigma factors functionality described in the
Introduction chapter, the influence of each of the seven known E. coli sigma factors
on the induction of yjb promoter in response to osmotic stress was tested.
4.2.2 Identification of the promoter region, promoters and transcription starts
sites
Ferrièrs et al. (2007) have recognized a putative RcsAB Box (Wehland and Bernhard,
2000), 251 bp upstream of the yjbE ATG and it was already shown that the
transcription of yjbEFGH is dependent upon RcsA and RcsB (see section 4.1.3). In
order to locate the promoter region that activates yjbEFGH transcription, the RcsAB
Box was deleted (Figure 18 A) from the original yjbF'::luxCDABE transcriptional
fusion (designated here pDEW/yjbF-1) to generate pDEW/yjbF-2. The NaCl-induced
activity was totally abolished in pDEW/yjbF-2 (Figure 18 B), indicating the existence
of a promoters region upstream of yjbE. To identify the sequences responsible for the
observed promoter activity, total RNA was isolated from strain MG1655 before and
following induction of transcription with 0.7 M NaCl, and the 5` end of the yjbEFGH
mRNA was mapped by primer extension with three specific primers (Figure 19 A-B).
Two transcription start sites were mapped upstream of yjbE start codon with an
almost equal amount of transcripts that have been produced from each and an
additional weak promoter between them. The first transcription start site is located 52
nucleotides (G) upstream of yjbE start codon and the second 18 nucleotides
downstream from the first (T/A).
A
pDEW/yjbF-1
RcsAB
Box
pgi
pDEW/yjbF-2
yjbE
yjbF’
luxCDABE
yjbE
yjbF’
luxCDABE
MAX.BIOLUMINESCENCE (RLU/OD600)
100000
B
NaCl (M)
0.7
0.5
0.3
10000
0.09
1000
100
pDEW/yjbF-1
pDEW/yjbF-2
PLASMID
Figure 18: (A) Organization of the segment cloned in pDEW/yjbF-1 (pDEW609) and its
truncated version cloned in pDEW/yjbF-2. (B) Maximal activity of pDEW/yjbF-1 and
pDEW/yjbF-2 obtained along 300 min in response to various concentrations of NaCl.
A
B
Sequencing
T
G
C
A
Primer
primer position relative to ATG
Extension
29
29 58 123
+ NaCl
- NaCl
174
157
109
92
G
80
TA
63
Figure 19: Promoter(s) analysis upstream of the yjbE gene by primer extension with single
(A; PrimEx1) or three different primers (B; PrimEx1 [left], 2 [middle] and 3 [right]). Arrows
indicate main transcription start sites; Numbers on the right indicate product length.
Sequencing reaction was primed with PrimEx1.
The two putative promoters derived from these transcription start sites are compared
in Table 5 to the proposed recognition sequences of the seven E. coli sigma factors.
The first promoter (P1; +1 = G) has a TATA Box (TATACT) fully identical to the
one recognized by σ38 (Weber et al., 2005) and almost identical to the one recognized
by σ70 (TATAAT; Hawley and McClure, 1983). The second promoter (P2; with +1 =
T or A) has –35 (TTGTGT) and –10 (AATGTT) sequence elements spaced by a 17nucleotides spacer that partially resemble those of σ70 (TTGACA-N(16-19)-TATAAT;
Hawley and McClure, 1983). Both have very little similarity to the proposed
recognition sequences of the other sigma factors.
Table 5: Putative promoters of yjbEFGH in comparison to the proposed consensus
recognition sequences of known sigma factors in E. coli. Purposed promoter elements (–35
and –10) and the mapped transcription start site (+1) are boldfaced and underlined.
SPACER
-35
-10
REFERENCE
(nt)
yjbE
+1
This study
P1
GCCGTAATTCGCATGCTTTAGTTGTGTATACTCGATCCCG
yjbE
+1
This study
P2
AGTTGTGTATACTCGATCCCGCCCGAAATGTTTTTGGGTA
Proposed consensus sequences
RpoD
TTGACA
16-19
(-13) TATAAT (-8)
RpoS
-
-
(-14) TCTATACTTAA (-4)
RpoH
CCTTGAAA
11-12
(-16) CCCCATTT (-9)
Hawley and McClure, 1983
Weber et al., 2005
Cowing et al., 1985
Nonaka et al., 2006
FliA
TAAAGTTT
11
(-15) GCCGATAA (-8)
Helmann and Chamberlin, 1987
Park et al., 2001
RpoE
GGAACTTTT
11
(-13) GGTCAAA (-7)
FecI
AAGGAAAAT
17
(-12) TCCTTT (-7)
TGGCACG
4
(-15) TTGCA (-11)
RpoN
Rhodius et al., 2006
Enz et al., 1995
Barrios et al., 1999
4.2.3 ı70 can drive transcription from yjb P1 in-vitro
Since 38 has been already shown not to activate yjb induction, the relevance of 70 to
yjb induction activation was further investigated by an in-vitro transcription
experiment using linear DNA templates ranging from -228 to +77 according to yjbE
P1 transcription initiation start site (G) and consisting of the RcsAB Box and the two
putative promoters, yjbE P1 and yjbE P2. This template was probed with [-32P]-ATP
or [-32P]-GTP in a 60 min multi-round reaction. As a control, T7A1 and pR
promoters were probed with [-32P]-ATP and [-32P]-GTP respectively, in a single
round reaction. As can be seen in Figure 20, the RNA polymerase holoenzyme drove
in-vitro transcription from yjbE P1 but not from yjbE P2, producing three transcript
group sizes: [-32P]-GTP end-labeled full length (expected to be 77 bases long),
paused (~40 bases long) and abortive transcripts. These data suggest that yjbE
transcription from P2 is 70-independent.
Promoter
89
77
yjbE
Labeled NTP ȖATP ȖGTP
T7A1 ȜpR
ȖATP ȖGTP
Full
Length
products
~40
Pausing
14
Aborative
products
Figure 20: In-vitro transcription from yjbE promoter by RNAP-
70
holoenzyme. T7A1 and pR
promoter served here as a control for the reaction. See text and Materials and methods
chapter for more details.
4.2.4 Regulation of yjbF'::luxCDABE by ı70 in-vivo
There are two major potential routes by which 70 can directly regulate yjbEFGH
transcription: (a) "normal" housekeeping regulation and (b) when growth is inhibited
(as, in our case, by osmotic shock), 70 activates an additional suite of stress response
genes, concomitant to the accumulation of ppGpp (Chang et al., 2002). As the latter
option is expected to occur in a ppGpp dependent manner, the involvement of ppGpp
in yjbEFGH induction needs to be examined before the role of 70 is addressed. To
determine the role of ppGpp in yjbEFGH activation, we have examined
yjbF'::luxCDABE activity in a ppGpp0 strain, mutated in both ppGpp synthase (relA)
and ppGpp synthetase/hydrolase (spoT) genes. As a control, the same experiment was
also performed using a lacZ'::luxCDABE transcriptional fusion; lacZ transcription is
supposed to be ppGpp-independent.
As demonstrated in Figure 21 A, yjbF induction following exposure to 0.3, 0.5 and
0.7 M NaCl was repressed in the mutant by 4, 7 and 9-folds in comparison to the wild
type, respectively. Activity was almost completely restored, however, when a third
mutation, a defective rpoD allele (rpoD40), was added on top of the relA spoT
background. The RpoD40 protein is characterized by a reduced ability to bind RNAP
and therefore, by a reduced ability to compete for its core (Jishage et al., 2002).
Induction of lacZ (with 0.5 mM IPTG) was similar in the wild type and the mutant in
0.09 M NaCl but was repressed in higher NaCl concentration (Figure 21 B) and was
not restored in the relA spoT rpoD40 triple mutant.
From these data it is impossible to determine whether yjb transcription is ppGpp
dependent since the induction of the control gene lacZ was repressed similarly,
probably due to the ppGpp0 strain sensitivity to salt stress. However, this ppGppindependent induction in the presence of a defective 70 (RpoD40), lies in the fact that
due to the lower affinity of the latter to the RNAP, an increase in free RNAP
availability to alternative sigma factors is expected. The enhanced availability should
facilitate the recruitment of free RNAP, and increase the transcription of genes under
MAX.BIOLUMINESCENCE (RLU/OD600)
the control of these alternative factors, including the one that regulates yjbEFGH.
80000
A
MC4100
70000
ǻrelA ǻspoT
60000
ǻrelA ǻspoT rpoD40
50000
40000
30000
20000
10000
0
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
MAX.BIOLUMINESCNCE (RLU/OD600)
NaCl (M)
4500000
B
4000000
MC4100
ǻrelA ǻspoT
3500000
relA spoT rpoD40
3000000
2500000
2000000
1500000
1000000
500000
0
0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
NaCl (M)
Figure 21: Induction of yjbF'::luxCDABE (A) and lacZ'::luxCDABE (B) transcriptional fusions
0
0
in response to various NaCl concentration in wild type, ppGpp and ppGpp rpoD40 strains.
We further investigated the regulation of yjbF induction by 70 in-vivo by monitoring
yjbF'::luxCDABE activity in a strain which transcribes its single copy rpoD gene from
the controllable trp-promoter; this was carried out in both rpoS+ and rpoSbackgrounds. The trp-promoter was activated with the tryptophane analogue indolacrylic acid (IAA). In the course of the present study we have employed two IAA
concentrations as described by Magnusson et al. (2003): 0.2 mM for normal 70
production and 0.02 mM for a reduced production of this sigma factor (low RpoD).
The effect of IAA concentration (= 70 production) and the presence of a functional
RpoS on the maximal activity of yjbF'::luxCDABE was examined in the presence of
either 0.09 M or 0.3 M NaCl. The higher salt concentrations previously used for the
induction of yjbF (0.5 and 0.7 M) were not employed in this case in view of the
higher osmotic sensitivity displayed by the mutants (not shown). In all cases, NaCl
caused a significant induction in bioluminescence. However, compared to the wild
type ("normal RpoD), NaCl-induced activity was 4, 10 and 75-fold higher in the
rpoS::Tn10/normal-RpoD cells, the low-RpoD cells and the rpoS::Tn10/low-RpoD
cells, respectively (Figure 22). Quite clearly, both inactivation of rpoS and reduction
in 70 have led to enhanced expression of the yjb transcriptional reporter. The effect
was obvious even in the presence of 0.09 M NaCl, where only a baseline expression
of yjbEFGH is normally observed, yielding an enhancement of 11, 5 and 65-folds
respectively in comparison to the normal-RpoD cells.
1000000
MAX.BIOLUMINESCENCE (RLU/OD600)
NaCl (M):
0.3
0.09
100000
10000
1000
100
Normal RpoD Normal RpoD
rpoS::Tn10
Low RpoD
Low RpoD
rpoS::Tn10
Figure 22: Activity of the yjbF'::luxCDABE transcriptional fusion under normal (0.2 mM IAA)
+
and low (0.02 mM IAA) induction levels of a single copy of rpoD in rpoS (Ptrp-rpoD; LM100)
and rpoS- (Ptrp-rpoD, rpoS::Tn10; LM1009), in response to 0.3 and 0.09 M NaCl.
4.2.5 Regulation of yjbEFGH promoter by alternative sigma factors
Mutants in either rpoE, rpoN, fliA or fecI did not exhibit a significant difference in
yjbF'::luxCDABE activity in comparison to their wild type strains (Table 6),
suggesting that none of these sigma factors participates in the regulation of yjbEFGH.
Table 6: Effect of sigma factor manipulated strains on yjbF'::luxCDABE activity in response to
osmotic stress.
Sigma
Genetic manipulation and experimental
Effect on yjbF'::luxCDABE
induction
conditions
(manipulated / reference)
RpoD
70
(σ
)
RpoS
38
(σ
)
RpoH
32
(σ
)
RpoE
19
(σ
54
24
28
Normal production of RpoD from trp-promoter
4
in rpoS:Tn10 background; LB 0.3 M NaCl
With or without overproduction of RpoH from
1.06
araB promoter; 0.5 M NaCl
Inactive rpoE; 0.5 M NaCl
0.87
Inactive rpoN; 0.5 M NaCl
1.13
Inactive fliA; 0.5 M NaCl
0.82
Inactive fecI; 0.5 M NaCl
0.89
)
FecI
(σ
promoter; LB 0.3 M NaCl
)
FliA
(σ
10
)
RpoN
(σ
Normal or underproduction of RpoD from trp-
)
The possible dependency of yjbEFGH on the heat shock response sigma factor RpoH
(σ32) was also approached by assessing its induction in an rpoH inactive strain
(CAG9333). The induction of both yjbF and lacZ (a control gene) in this temperature
sensitive mutant were similarly inhibited over 10-fold (data not shown). This, on top
of the slow growth rate of this strain, indicates that its transcriptional capabilities are
generally hindered by its overall poor physiological condition. It was thus impossible
to determine if the decreased induction of yjbEFGH is due to direct regulation of σ32
on its promoter.
To overcome this problem, we have adopted an opposite approach: yjbF'::luxCDABE
activity was assessed under conditions of σ32-overproduction, obtained by an
arabinose-controlled σ32 overproducing vector (pAR1), and compared to a strain with
normal σ32 levels. As demonstrated in Figure 23, the enhanced production of σ32 had
no effect on yjbEFGH expression. At the same time, as expected, the induction of lon,
a heat shock dependent protease, was markedly enhanced in the arabinose-induced
cells.
lon, -arabinose
yjbG, +arabinose
yjbG, -arabinose
1600000
4000
3000
1200000
2000
800000
1000
400000
0
0
50
100
150
200
250
yjbF , BIOLUMINESENCE (RLU/OD600)
lon , BIOLUMINESCENCE(RLU/OD600 )
2000000
lon, +arabinose
0
300
TIME (min)
Figure 23: Activity of the yjbF'::luxCDABE and lon'::luxCDABE transcriptional fusions under
normal (0 % L-arabinose) and high (0.2 % L-arabinose) induction levels of a multi copy rpoH
gene from a plasmid (PBAD::rpoH; MG1655, pAR1), in response to 0.5 M NaCl.
4.2.6 In-vivo characterization of transcription pausing sites in yjb
The two pausing sites that were identified in the in-vitro transcription experiment ~40
bp downstream from yjbE P1 transcription initiation start site (Figure 20) were further
examined in-vivo. Sequence analysis using the mFold software predicts a stem-loop
mRNA secondary structure at the approximate pausing site (Figure 24 A). Two pairs
of plasmids were therefore constructed harboring two different yjbE::luxCDABE
transcriptional fusions (Table 7); one contains the stem-loop (fused at position +76)
and the other does not (fused at position +27).
The transcriptional fusions activity in the wild type strain MG1655 in response to 0.7
M NaCl is shown in Figure 24 B. In both plasmid pairs, the activity of the short fusion
(yjbE27) was ~2-folds higher than the activity of the long fusion (yjbE76). Thus, in
addition to the essentiality of the RcsAB Box sequence to the activation of
transcription that was demonstrated in section 3.2.1, the region upstream of the
RcsAB Box also appears to have a regulatory function. When it was truncated
(RcsABox-yjbE27 or 76) the activity was significantly inhibited. Interestingly,
transcription from yjbE was always weaker than from yjbF (pDEW/yjbF-1); this
result was obtained also from the RT-PCR analysis (Figure 24 C).
Table 7: Plasmids for in-vivo characterization of the pausing site found in-vitro in yjbE 5' UTR.
PLASMID
CLONNED SEGMENT
GENOTYPE
PRESENT FEATURES
pgi end
RcsAB Box
Stem loop at
yjbE 5' UTR
pDEW/yjbE27-1
pgi-yjbE27::lux
+
+
-
pDEW/yjbE76-1
pgi-yjbE76::lux
+
+
+
pDEW/yjbE27-2
RcsABox-yjbE27::lux
-
+
-
pDEW/yjbE76-2
RcsABox-yjbE76::lux
-
+
+
A
+1
+27
GAAAUGUUUUUGGGUAAAUCUCCAUUCAUUCAAUGAAGGGAAAU
+76
MAX.BIOLUMINESCENCE (RLU/OD600)
UGUUAUGAAAAAAGUUCUGUAUGGCAUUUUUG…
16000
- pausing
site
B
14000
12000
10000
8000
+ pausing
site
6000
- pausing
site
4000
2000
+ pausing
site
0
yjbE27-1
yjbE76-1
yjbE27-2
yjbE76-2
INDUCTION or EXPRESSION RATIO
(yjbE/yjbF )
PLASMID (pDEW/)
0.6
C
0.5
0.4
0.3
0.2
0.1
0
RT-PCR
Transcriptional fusion
ASSAY
Figure 24: (A) stem loop predicted by the mFold software at the approximate pausing site.
The complementary nucleotides are underlined; numbers indicate the fusion site to the lux
genes (+27, pDEW/yjbE27-1/2; +76, pDEW/yjbE76-1/2). (B) Induction of the different
yjbE::luxCDABE transcriptional fusion in response to 0.7 M NaCl. (C) yjbE/yjbF induction
(transcriptional fusion) and expression (RT-PCR) ratios, calculated from data obtained
following induction of the yjb operon with 0.7 M NaCl. The transcriptional fusions used here
are yjbF'::luxCDABE (pDEW/yjbF-1) and yjbE27::luxCDABE (pDEW/yjbE27-1).
4.3 Regulation of yjb promoter activity by growth conditions
4.3.1 Induction of the yjb operon occurs in early batch growth
As demonstrated in Figure 25 A, the level of yjb induction in batch cells exposed to
NaCl (0.7 M) at different points along their growth curve was strongly dependent
upon the growth phase. Induction of yjbF'::luxCDABE activity (carried out in a 96well microtiter plate) was maximal when the exposed cells were withdrawn from a
culture at an OD600 of 0.2, and decreased to extinction toward stationary growth phase,
at an OD600 of 1 (growth curve is shown in Figure 25 B). As also shown in Figure 24
A, this was also the pattern of yjb induction exhibited in the rpoS::Tn10 mutant, the
overall activity in which was much stronger. The latter observation is a strong
indication that occurrence of the induction only in the earlier stages of growth was not
a result of negative regulation by RpoS, accumulating as the cells approach stationary
phase. Without the osmotic shock, no induction of the yjb operon was observed either
in the wild type or in the rpoS::Tn10 mutant.
Batch culture cells in complex media experience continuous changes in both density
and specific growth rate (Wanner and Egli, 1990), both of which could be
instrumental in modifying gene regulation and expression (Beav et al., 2006). To
differentiate between the effects of these two parameters on yjb induction, they were
therefore investigated individually and independently in batch and continuous culture.
.
120
A
MG1655, 0.7 M NaCl
30000
rpoS::Tn10, 0.7 M NaCl
100
MG1655, 0.09 M NaCl
25000
80
rpoS::Tn10, 0.09 M NaCl
20000
Plasmid copy number
60
15000
40
10000
PLASMID COPY NUMBER
BIOLUMINESCENCE (RLU/OD600)
35000
20
5000
0
0
0
0.2
0.4
0.6
0.8
1
CULTURE DENSITY (OD600)
OPTICAL DENSITY (600 nm)
1
B
0.1
0.01
0
25
50
75
100
125
150
TIME (min)
Figure 25: (A) Bioluminescence activity of the yjbF'::luxCDABE transcriptional fusion in
response to osmotic shock, in cells withdrawn at different stages of batch culture in LB broth
(B) as assayed in a 96-well microtiter plate. Plasmid copy number per cell is also shown (A).
4.3.2 Induction of the yjb operon is not quorum-sensing dependent
Cell density effects on gene regulation are often mediated by external signal molecules
that affect gene expression when they reach a certain threshold concentration, a
phenomenon commonly referred to as quorum sensing (Fuqua et al., 1994; Ahmer,
2004). In order to determine what limits yjb induction to early stages of growth in
batch culture, first the quorum-sensing mechanism was tested. Two MG1655 batch
cultures were grown to OD600=0.2 (where activity peaks) and OD600=1 (where activity
silenced) and than each was separated to cells and medium by centrifugation. Subcultures of each growth phase were than suspended in each of the mediums and
assayed for yjbF'::luxCDABE activity. Activity of the young and less dense culture in
both media (young and old) was similar and ~40-folds higher than of the old and
more dense culture (Figure 26) indicating that there is no quorum-sensing molecule in
the old medium that represses yjb induction. This experiment was performed only
once.
MAX.BIOLUMINESCENCE (RLU/OD600)
20000
0.2
18000
16000
CULTURE GROWTH PHASE (OD600)
1
14000
12000
10000
8000
6000
4000
2000
0
0.2
1
MEDIUM ORIGIN (CULTURE OD600)
Figure 26: Induction of yjbF'::luxCDABE in wild type cells that were withdrew from OD600=0.2
(young) and 1 (old) and than suspended in each of the media (old and young).
4.3.3 Induction of the yjb operon is highest in minimal media and at low growth
rates
To test yjbF induction under different specific growth rates, strain MG1655 harboring
the yjbF'::luxCDABE transcriptional fusion was grown in a chemostat at different
dilution rates, in either a glucose-limited minimal medium (ClimBa) or in LB broth.
Since several studies reported that rpoS might be inactivated or lost during long-term
incubation at low growth rates (Notley-McRobb et al., 2001; Chen et al., 2004), the
experiment was also conducted in the rpoS::Tn10 mutant.
After a steady state culture density was obtained at each dilution rate, culture aliquots
were removed, diluted to an OD600 of 0.2 (cell density where maximal induction was
observed, see Figure 25 A) and exposed to either 0.5 M or 0.09 M NaCl in 96-well
plates. Bioluminescence of the yjbF'::luxCDABE fusion was then monitored for 200
min. From the results (Table 8), three very clear observations can be made:
(a) In the wild type strain, enhancement of yjbF'::luxCDABE activity at all growth
rates was much stronger in cells grown in a minimal medium than in cells grown in
LB.
(b) In all cases – minimal and complex media alike – activity was always highest in
the slowest growing cells.
(c) The latter phenomenon was not restricted to NaCl-induced cells: significant
induction of yjbF took place even without an osmotic shock.
Activity in the rpoS::Tn10 mutant (assayed in LB only) was, as expected, higher than
that of the wild type, but the basic observations noted above were maintained: activity
was much higher for D=0.1 h-1, both in the presence and absence of NaCl.
Table 8: Effect of dilution rate and culture medium on the activity of yjbF'::luxCDABE in
steady state chemostat culture samples, diluted to an OD600 of 0.2 in a microtiter plate, and
exposed to NaCl at either 0.5 M (induced) or 0.09 M (non-induced). Bioluminescence was
monitored in the microtiter plate for 300 min after exposure to NaCl.
STRAIN
MEDIUM
BIOLUMINESCENCE (RLU/OD600)
Non induced
Induced by NaCl
0.5
168 ± 38
6433 ± 7312
Minimal
0.3
208 ± 12
8892 ± 10725
ClimBa
0.2
490 ± 42
ND
0.1
1294 ± 582
59626 ± 1027
1
293 ± 23
3155 ± 1033
0.5
350 ± 74
1349 ± 946
0.1
4251 ± 817
20329 ± 2666
1
450 ± 130
14525 ± 3190
0.5
750 ± 507
12850 ± 2814
0.1
6006 ± 1121
363181 ± 48241
MG1655
LB
rpoS::Tn10
D (h-1)
LB
4.3.4 Induction of the yjb operon is cell density related
To measure the effect of cell density on yjb activation, samples removed from the LB
chemostats (both wild type and the rpoS::Tn10 mutant) were serially diluted and
assayed for yjbF'::luxCDABE activity as described above. The results (Figure 27)
demonstrate a very clear dependency on cell concentration in the assay plate: at all
conditions tested, activity was considerably higher at the lower cell densities. As
already observed, highest activity was measured at the slower growing cells (dilution
rate 0.1 h-1) both in the wild type and in the rpoS::Tn10 mutant.
500000
MG1655, 0.1 (1/h)
450000
MG1655, 0.5 (1/h)
25000
400000
MG1655, 1 (1/h)
20000
15000
rpoS::Tn10, 0.1 (1/h)
350000
rpoS::Tn10, 0.5 (1/h)
300000
rpoS::Tn10, 1 (1/h)
250000
200000
10000
150000
100000
5000
50000
0
rpoS::Tn10 , BIOLUMINESCENCE (RLU/OD600)
MG1655, BIOLUMINESCENCE (RLU/OD600)
30000
0
0
0.5
1
1.5
2
2.5
OPTICAL DENSITY (OD600)
Figure 27: Effect of cell density and chemostat culture dilution rate on yjbF'::luxCDABE
bioluminescence activity. Aliquots, removed from steady state chemostat LB cultures growing
at different dilution rates, were serially double diluted and their bioluminescence was
monitored in 96-well microtiter plate as described.
4.3.5 Induction of the yjb operon depends upon gas exchange rates
In Figure 28 we compare the induction of yjbF'::luxCDABE exposed to NaCl in the
wells of an almost stationary microtiter plate to that taking place in a continuously
shaking Erlenmeyer flask. In both incubation modes, no activity was observed
without the addition of NaCl; following the osmotic shock, induction was observed in
both cases, but was much stronger in the flasks. As already observed, there was a
strong dependency on cell density, and the general pattern was similar in both
incubation modes, with maximal activity at an OD600 of around 0.2.
Shaking flask 0.7 M NaCl
Shaking flask 0.09 M NaCl
Microtiter plate 0.7 M NaCl
Microtiter plate 0.09 M NaCl
BIOLUMINESCENCE (RLU/OD600)
45000
40000
35000
30000
25000
20000
15000
10000
5000
0
0
0.2
0.4
0.6
0.8
1
CULTURE DENSITY (OD600)
Figure 28: Activity of the yjbF'::luxCDABE transcriptional fusion in response to osmotic shock,
in cells of different stages of LB batch growth as assayed in either a microtiter plate or in a
shaking flask, in 0.09 and 0.7 M NaCl.
The striking difference in the magnitude of the induction in cells of identical density
and specific growth rate, exposed to the same NaCl concentration, may be due to the
difference in aeration rate, high in the shaking flask but very low in the stationary
microtiter plate. It was therefore tested whether the intensity of yjb induction may be
oxygen dependent. To test this hypothesis, we have assayed yjbF'::luxCDABE
induction in a series of identical flasks containing different volumes of the same cell
culture. The culture used in the experiment was batch-grown to an OD600 of 0.8, a cell
density at which yjb induction does not occur in a microtiter plate assay. Total flask
volume was 300 ml, and the media volumes used, 50-300 ml, generated surface to
volume ratios of 1.13-0.03 cm-1. As seen in Figure 29 A, yjbF'::luxCDABE activity
was strongly dependent upon the different surface to volume ratios. It was maximal in
the highest and absent in the lowest.
Since bioluminescence is strongly oxygen dependent, it was important to demonstrate
that the aeration effect was indeed exerted on the activation of yjb and not on the
activity of the luciferase reporter. Two experimental approaches were utilized for this
purpose, in which two additional plasmids were used instead of the one carrying the
yjbF'::luxCDABE fusion. In the first, we have substituted the yjbEFGH promoter with
that of the lacZ gene, and in the second we have made use of the lacZ gene as a
reporter instead of the lux-genes.
When bioluminescence activity of lacZ'::luxCDABE was monitored following
induction of the lacZ promoter by 0.5 mM IPTG (under the same experimental
conditions), induction of lacZ exhibited an opposite pattern in comparison to that of
yjbF (Figure 29 B). While the reason for this pattern is at present unclear, it is obvious
that at the highest culture volumes and lowest area/volume ratios, bioluminescence
was not inhibited. In fact, maximal luminescence exhibited by the lacZ'::luxCDABE
transcriptional fusion was 100-folds higher than the luminescence signal obtained
from the yjbF'::luxCDABE transcriptional fusion. This clearly indicates that the
inhibition of yjbF'::luxCDABE activation at the lower surface/volume ratios was not
due to unavailability of molecular oxygen to the bacterial luciferase. It also signifies
that the decreased yjb driven luminescence at the lower surface/volume ratios is not a
result of a general down-regulation of cellular activity due to lack of oxygen.
To further verify this observation, the oxygen-independent lacZ reporter system was
also used to monitor yjbF activation. The two reporting systems, yjbF'::luxCDABE
and yjbF'::lacZ, were assayed in parallel in the flask system described above, in the
two extreme surface/volume ratios depicted in Figure 29 A. As demonstrated in
Figure 30 A-B, activity of both reporters was strongly inhibited in the low S/V ratio
flask. When an identical flask was aerated, activity returned to a level approximately
75 % of the high S/V flask.
BIOLUMINESCENCE (RLU/OD600)
25000
A
S/V=1.13
S/V=0.48
20000
S/V=0.24
S/V=0.12
15000
S/V=0.03
10000
5000
0
0
50
100
150
200
250
TIME (min)
4000000
25000
B
lacZ'::luxCDABE
yjbF'::luxCDABE
20000
3000000
15000
2500000
2000000
10000
1500000
5000
yjbF', BIOLUMINESCENCE
(RLU/OD600)
lacZ , BIOLUMINESCENCE
(RLU/OD 600)
3500000
1000000
500000
0
0.0
0.2
0.4
0.6
0.8
1.0
1.2
AREA / VOLUME (1/cm)
Figure 29: Effect of gas exchange capacity on yjbF induction. (A) Kinetics of yjbF'::luxCDABE
activity in cultures of different surface area/volume ratios (in cm-1) following induction by NaCl
(0.7 M). (B) Differential effects of surface area/volume ratios on bioluminescence driven by
the yjbF and by the lacZ-promoters induced by in the presence of 0.7 M NaCl and 0.5 mM
IPTG.
BIOLUMINESCESNE (RLU/OD600)
60000
A
50000
40000
30000
20000
10000
0
1.13
0.03
0.03 +
AREATION
AREA / VOLUME (1/cm)
LacZ ACTIVITY (RFU/minXOD600)
200000
B
160000
120000
80000
40000
0
1.13
0.03
0.03 +
AREATION
AREA / VOLUME (1/cm)
Figure 30: Effect of surface area/volume ratio on yjbF induction: comparison of luciferase (A)
and ; -galactosidase (B) reporting systems. One set of low surface/volume ratio flasks was
continuously aerated.
4.3.6 Induction of yjbF is oxygen dependent
To further demonstrate the oxygen dependency of yjb induction, a chemostat
experiment was conducted in which the only manipulated variable was oxygen
concentration. Strain MG1655 harboring the yjbF'::luxCDABE transcriptional fusion
was allowed to reach steady state in ClimBa medium, at a constant specific growth rate
(0.2 h-1) but under three different aeration regimes: (a) aeration (1.4 l min-1) and
mixing at 800 rpm; (b) aeration (1.4 l min-1) and mixing at 200 rpm; (c) mixing at 200
rpm without aeration. Dissolved oxygen concentrations under these conditions were
6.2, 3.9 and 0.1 mg l-1 respectively. Figure 29 displays the kinetics of
bioluminescence following the addition of NaCl (0.7 M). The background
luminescence characteristic of growth at D=0.2 h-1 (compare to Table 8) was initially
inhibited, and re-developed after about 90 min. Activity after 180 min was maximal in
the highest oxygen concentration and absent in the lowest.
2500
BIOLUMINESCENCE (RLU/OD600)
800 rpm + air pumping
200 rpm + air pumping
2000
200 rpm
1500
1000
500
0
-20
0
20
40
60
80
100 120 140 160 180 200
TIME (min)
Figure 31: Osmotic induction (0.7 M NaCl) of yjbF'::luxCDABE in chemostat cultures of a
-1
constant dilution rate (D=0.2 h ) and different dissolved oxygen concentrations. The three
aeration regimes generated stable dissolved oxygen concentrations of 6.1, 3.9 and 0.1 mg O2
-1
l , respectively. NaCl exposure took place in the chemostat by the addition of a concentrated
NaCl solution.
5. DISCUSSION
5.1 What regulon does the yjb operon belong to?
In the few previous reports of DNA-microarray studies that have investigated the
response of E. coli to osmotic shock (Weber and Junk, 2002; Weber et al., 2005),
members of the yjb operon have not been listed among the osmotically regulated
genes. This may be attributed to the lower NaCl doses usually applied (< 0.5 M), the
short exposure duration (< 30 min) or to the relatively low activity of the promoter
that drives its expression. In this study, based on the induction characteristics of the
yjbF'::luxCDABE transcriptional fusion (induction by NaCl and repression by σ38), a
DNA-microarray experiment was performed in order to find genes that behave
similarly. The experimental conditions (NaCl concentration and exposure time) were
extremely drastic and without the data obtained from the transcriptional fusion
characterization that guided this experiment these genes might have been ignored here
as well.
Among the genes that fitted to both criteria stood out genes related to biofilm
formation (bssR, yddV, ttdA, yihR and yeeJ), adhesion (fimbria like proteins) and
above all, EPS synthesis (yjb and wca operons). Surprisingly, only eight additional
RcsC-dependent genes (out of 83, as reported by Ferrières and Clarke (2003)) appear
on this list, suggesting a possible division of those to several smaller regulons.
It was previously shown that activation of the Rcs-pathway increases σ38 translation
in an RcsA-independent manner (Majdalani et al., 2001; Majdalani et al., 2002;
Peterson et al., 2006). Thus, the expression of the RcsC-dependent and σ38-repressed
genes appears to be a subject of conflicting regulatory effects. Although the exact cue
that activates the signal transduction cascade through the Rcs-system is not known,
and since the Rcs-system is clearly regulating genes involved in envelope stress
response (Majdalani and Gottesman, 2005), it can be rationalized that it enhances σ38
translation in order to enhance induction of stress response genes. Indeed, several
Rcs-dependent genes overlap those found in the σ38 regulon (Ferrières and Clarke,
2003; Weber et al., 2005), including osmB (an osmotically and stress inducible
lipoprotein of an unknown function), which is also listed among the eight osmotically
induced, σ38-repressed genes found in the DNA microarray analysis.
In this high throughput screen, most genes of the yjb and wca operons that were
induced in response to osmotic stress exhibited an NaCl response ratio of 2-9 folds
only (Table 4), while the yjbF'/wza' - lux transcriptional fusions yielded an NaCl
response ration of ~50 (yjbF'; Figure 7 A) and ~350 (wza'; Figure 7 C). DNA
microarray analysis is a direct measurement of transcripts abundance, while the use of
transcriptional fusions is indirect. The lux genes have their own ribosome binding
sites, and besides being dependent on the upstream promoter for transcription, all the
downstream applications (e.g. mRNA stability and translation) are expected to be
independent of it and of the native gene it regulates. Thus, luminescence monitoring
does not provide data on post transcriptional regulation on the studied gene. Based on
this it may be suggested that the low expression values obtained by the DNA
microarray analysis could also be a result of post transcriptional regulation of yjb and
wca transcripts. This hypothesis was not examined.
5.2 Comparison between the yjb and wca regulatory regions
From the genes found in the DNA microarray analysis, the wca operon genes were
further studied. The promoter of this operon was cloned and found to be activated and
induced in an almost identical manner to that of yjb. yjb and wca promoters are both
activated by the Rcs-pathway in a RcsA and temperature-dependent manner,
repressed by σ38 (see model in Figure 32) and exhibit identical induction properties
except for the intensity which is much stronger in the latter. This difference cannot be
attributed to the differences of the transcriptional fusions (size of cloned fragment,
fusion site according to the promoter, ect.) since it was obtained also in the RT-PCR
experiment. Differences in induction intensity can stem from one or more out of many
possible DNA sequence properties out of many possible that differentiate yjb and wca
promoter region characteristics.
Pi
bind
Sensor kinase
Response regulator
Auxiliary protein
RcsC
RcsB
RcsA
37
yjb
°C
°C
27
wca
RpoS
RpoD
Compete for
RNA polymerase
Figure 32: A model describing the common regulatory features of yjb and wca operon.
Transcription activation of wca operon by
σ70 has been proposed by Stout (1996) based on
promoter sequence. Transcription activation of yjb operon by
σ70 will be discussed in section
5.7. Arrows represent positive affect on activity/level; blocked lines represent negative effect
on activity/level.
-35
-10
yjbE P2 AGTTGTGT - 17nt - AAATGTT
wza
TCTTGCCT - 16nt - AACACTT
Figure 33: comparison between the proposed -10 and -35 elements of yjbE P2 and wza
promoters. The wza-promoter was identified by Stout (1996).
Stout (1996) has reported that a single promoter is present upstream of the first gene
of the wca operon (wza) while here it was shown that there are at least two promoters
upstream of yjbE. The yjbE P2 -10 and -35 elements are highly similar to those of the
wza promoter (Figure 33). The two primer extension experiments were performed
under very different conditions: transcription of wza was achieved artificially by
overexpression of RcsA, while transcription of yjbE was achieved by osmotic stress.
The same author showed that the 5' UTR of wza contain a JUMPstart sequence (Just
Upstream of Many bacterial Polysaccharide gene clusters; Hobbs and Reeves, 1994),
the role of which is still unknown; such a sequence is not present in yjbE. Ebel and
Trempy (1999) reported that there are two RcsAB boxes upstream of wza while
Ferrières et al. (2007) reported only one upstream of yjbE. In the present study it is
shown that there is a transcription pausing site in the 5' UTR of yjbE that reduces
bioluminescence signal by half; Ferrières et al. (2007) reported that there is another
secondary structure (ERIC, Enterobacterial Repetitive Intergenic Consensus; Wilson
and Sharp, 2006) in the intergenic region between yjbE and yjbF. Any of these
sequence properties, as well as others, may be responsible for the differentiated
induction intensity of the yjb and wca promoters.
5.3 Overproduction of EPSs in an rpoS deficient strain
In accordance with the overexpression of yjb and wca operons, both EPSs (wca
dependent (CA-EPS) and yjb dependent) were also found to be overproduced in the
rpoS mutant.
Inactivation of rpoS appears not to be a rare phenomenon, and has been described as a
natural beneficial mutation (Ferenci, 2007) in glucose-limited E. coli populations
(Notley-McRobb et al., 2001). Mutations in rpoS have also been detected among
environmental and clinical isolates of pathogenic as well as commensal enteric
bacteria (Robbe-Saule et al., 2003; Bhagwat, 2006). The selection pressure on the
rpoS gene is thought to be largely due to competition between the sigma factors σ38
and σ70 for a limiting number of RNA polymerases core subunits (Farewell et al.,
1998). It was shown that the spread of the rpoS inactive allele through the majority of
the population was accompanied with decreased stress tolerance but by an improved
nutrient scavenging ability due to increased expression of σ70-dependent genes
(Ferenci, 2007). While the role of the yjb encoded proteins in EPS production and
cellular physiology remains to be elucidated, it is logical that they have a beneficial
effect under stressed growth conditions. It was reported that the CA-EPS exerts some
stress protection, including against osmotic stress in E. coli O157:H7 (Moa et al.,
2001; Chen et al., 2004; Moa et al., 2006). Hence, it is possible that its overproduction
may compensate for the loss in σ38-dependent general stress response genes protective
activity.
In the course of this study the question of the benefit of EPS overproduction has not
been addressed. It should be explored in the future by assessing the contribution to
stress resistance of the yjb and wca operons to an rpoS mutant in comparison to their
contribution to the wild type.
5.4 The role of yjb genes in rpoS deficient strain
In this study it was shown that overinduction of the yjb operon in an rpoS deficient
strain is also accompanied by the emergence of what seems to be a regulatory role for
yjbE, yjbF, yjbH and, to a much lesser extent, yjbG. In an rpoS deficient strain,
inactivation of yjbE, yjbF or yjbH resulted in NaCl-independent mucoid colony
appearance. The yjbE rpoS::Tn10 and the yjbH rpoS::Tn10 double mutants also
become giant filaments. These observations suggest that the yjb operon, only in an
rpoS mutant, may regulate CA-EPS production, cell division and its own expression.
Based on the yjbE/H rpoS::Tn10 double mutants phenotypes, mucoid and
filamentous, it can be hypothesized that this regulatory action is indirect and is
mediated by Lon protease. RcsA and SulA are both regulated by Lon and lon mutant
is mucoid and filamentous. Hence, the level of Lon or its activity might be a subject
for regulation by YjbE and YjbH when σ38 is inactive (see model in Figure 34). The
observations that the yjbF rpoS::Tn10 mutant is only mucoid and that yjbG
rpoS::Tn10 mutant is neither mucoid nor filamentous, raise the possibility that this
regulatory affect is differentiated based on the integrity of the YjbEFGH system. In
this model, YjbE and YjbH are essential for the system integrity and their inactivation
results in phenotypes identical to those of the yjbEFGH rpoS::Tn10 mutant. YjbF
inactivation affects the function of the system in a way that enhances CA-EPS
synthesis but doesn't repress cell division, and YjbG is the most dispensable
component of the system. Based on sequence analysis, YjbG contains a domain
(Domain of Unknown Function 1017 (DUF1017)) that is present also in its
paralogous protein GfcC; although gfcC is not expected to be expressed in E. coli
MG1655, it may be hypothesized that if it is present, even at very low levels, it can
compensate for YjbG function.
The truncated wza' gene, cloned in pDEW/wza was able to compensate for the
absence of yjbE in the yjbE rpoS::Tn10 mutant. This result is disturbing since the
endogenic wza gene, like the whole wca operon, is supposed to be intact and
functional in this strain; it can be suggested that since this plasmid is present in ~30
copies in each cell, it is probably able to drive overexpression of Wza' that may
compensate somehow for the loss of YjbE.
RcsCDB
Phosphorelay system
yjb
Cell division
wca
FtsZ
YjbE/H
?
SulA
Lon
Regulation
of EPS
production
RcsA
RpoD
Figure 34: A model describing the proposed modulation of EPS production and cell division
by yjbE and yjbH through Lon in
σ38 deficient strain. Arrows represent positive affect on
activity/level; blocked line represents negative affect on activity/level.
5.5 A novel method for EPS production detection and comparison
EPS overproduction in the yjbG rpoS::Tn10 was revealed and demonstrated
following the establishment of a simple methodology for centrifuge pellet size
quantification. Bacterial cultures can be distinguished based on differences in their
extracellular properties which reflected by the volume they occupy when compressed.
The yjbG rpoS::Tn10 mutant, unlike other yjb rpoS::Tn10 strains, did not form
mucoid colonies but slightly overproduced CA-EPS when subjected to osmotic stress.
While a buoyancy assay was insufficient to indicate differences in EPS content
between this mutant and its parental strains or following growth in iso-osmotic and
hyperosmotic conditions, the differences in bacterial pellet volumes were very
pronounced. We also applied this method to other EPS production systems, putting to
the test an EPEC O127:H6 wild type strain and its non-capsulated etk::kanR mutant
(Peleg et al., 2005); the wild type strain exhibited doubled (2.3 ± 0.1) pellet volume in
comparison to the non-capsulated mutant.
This simple and fast procedure was utilized here to track changes in EPS production
in E. coli but it can be also used for initial screen of environmental and clinical
isolates. The observed differences can reflect not only differences in EPS content, but
also in cell size, cell shape and extracellular structures like fimbriae and flagella.
5.6 The yjbEFGH promoter region
Two major promoters were identified between the RcsAB Box and the yjbE start
codon. Both promoter sequences mostly resemble the σ70 or σ38 recognition
consensus sequence. Truncation of the whole DNA segment upstream of the
promoters (including the RcsAB Box) abolished promoter activity completely.
Ferrières et al. (2007) reported differentiated expression of yjbE and the downstream
yjb gene members due to stem loop structure in the intergenic region between yjbE
and yjbF. In their experiment the yjb genes were induced artificially by
overexpression of DjlA which stimulates RcsC phosphorylation of RcsB (Kelley and
Georgopoulos1997); under these conditions, the expression levels of yjbF, yjbG and
yjbH were several magnitudes lower than the expression of yjbE. Using different
induction conditions (NaCl induced osmotic stress) the bioluminescence signal
obtained from the yjbF'::luxCDABE transcriptional fusion was always greater than the
signal obtained from any of the yjbE'::luxCDABE transcriptional fusions. In addition,
the expression ratio of yjbE/yjbF obtained by the relative RT-PCR analysis was very
similar. These results suggest that when the yjb operon is induced by osmotic stress,
this stem loop, a termination signal, is being ignored by RNAP. The other stem loop
structure that was described in the 5' UTR of yjbE and was shown to reduce
transcription activity in-vitro and in-vivo, might be a transcription barrier that acts to
modulate the fraction of RNAPs that produce full length yjb transcripts. While it
seems to be active during osmotic stress, it might be resolved in other conditions, e.g.
during biofilm formation.
5.6 Regulation of yjb operon by sigma factors
The simple rationale almost universally employed to characterize regulons and map
regulation networks is based on activity inhibition in the absence of the regulator and
activity enhancement when it is in excess. The use of this approach to determine
which sigma factor protein controls yjbEFGH induction in response to osmotic shock,
led to the intuitive conclusion that none of the seven known sigma factors of E. coli
positively regulates yjbEFGH promoters in-vivo. We have shown that the absence of
σ24, σ54, σ19 or σ28 did not affect yjbEFGH induction, and neither did overproduction
of σ32. We also showed that underproduction of σ70, like inactivation of σ38, increased
the induction of yjbEFGH. Furthermore, this operon was not mentioned in any of the
comprehensive gene lists shown to be regulated by σ38 (Weber et al., 2005), σ32 (Zhao
et al., 2005) or σ19 (Dartigalongue et al., 2001) under any conditions.
σ70 level manipulation is problematic since the effect is universal. It is the most
dominant sigma factor in the cell, and cannot be completely inactivated. Hence,
overinduction of the yjb operon in the low RpoD culture can be attributed to many
different mechanisms and does not rule out the basic assumption that like 90% of E.
coli genes, yjbEFGH is probably also transcribed by the RNAP-σ70 holoenzyme. Our
in-vitro data indicate that σ70 can drive productive transcription from yjbE first
promoter (P1). Also supporting this notion is the high similarity of yjbE P1 TATA
Box to the consensus sequences recognized by σ70. Based on these evidences it can be
hypothesized that σ70 is likely to be the sigma factor protein that activates this
promoter regardless of the in-vivo results.
The second promoter (P2) that yielded no transcripts in the in-vitro experiment might
be σ70-independent. Indeed, it has been reported that several genes are activated by
more than a single sigma factor protein (Raina et al., 1995; Weber et al., 2005) and in
fact, σ38 and σ70 have been shown to activate the same promoters in-vitro (Gaal et al.,
2001; Hengge-Aronis, 2002). The dramatic overinduction in the Ptrp-rpoD rpoS::Tn10
strain rules out the possibility that σ38 is involved in yjb transcription; the fact that
there was no significant change in the overall activity of the transcriptional fusion in
any of the other sigma factor manipulated strains, may be a result of (1) wider
redundancy than thought, between sigma factors (yjb transcription can be carried out
by several factors) or (2) the existence of a novel sigma factor(s) that has not
discovered yet in E. coli genome; bacteria species posses a variable number of sigma
factor encoding genes, from as little as one in Mycoplasma species to over 60 in
Streptomyces coelicolor (Mittenhuber, 2002; Gruber and Gross, 2003), so although E.
coli genome has been extensively studied and annotated, the concept of a new sigma
factor may be worthy of additional scrutiny.
5.8 Regulation of yjbEFGH induction by oxygen and specific growth rate
Regulation of gene expression is a complex multidimensional process, and the activity
of a single gene may be induced, modulated and silenced to different extents by
numerous parameters. The yjb operon is an excellent example of this complexity. By
isolating each of three variables, growth rate, culture density and oxygen
concentration, and assessing their effects independently of the others, it was
demonstrated here that all the three influence the induction of this operon. Low
culture density and high oxygen availability affected the osmotically induced activity
of this operon, while low growth rates acted as an independent inducer
Actual modulation signals in gene regulation may stem from either changes in the
external environment or from internal physiological consequences of such changes.
Induction of the yjbEFGH operon is dependent upon the Rcs-phosphorelay system.
Since the phosphorylation cascade is initiated at the cytoplasmic membrane, the
activation signal is likely to be an external one. Osmotic induction by NaCl fits this
model. In this study, however, I have demonstrated that the induction of yjbEFGH
also responds to an internal signal, low specific growth rate; even without an apparent
external signal, activity in the non-induced control was much stronger in slowgrowing cells.
Osmotic stress on batch-grown cells temporarily arrests growth for the time period
required for maintenance and adaptation, before resuming proliferation. This growth
arrest may be viewed as very similar to growth at a low dilution rate in a chemostat,
explaining the induction of the yjb operon under both sets of conditions.
As shown in this study, yjbEFGH induction also depends upon the availability of
oxygen. I have demonstrated that this dependency, apparent both in batch and
continuous growth, is not a reflection of the general wellbeing of the culture, nor is it
an artifact stemming from the use of luciferase as a reporter. At this time it is not clear
whether the effect is mediated by a direct O2 interaction of the type exerted on
regulatory proteins such as Fnr, ArcAB or OxyR (Repoila et al., 2003; Alexeeva et
al.,2003; Shalel-Levanon et al., 2005; Zheng and Storz, 1999) or whether it is a more
general effect caused by a decrease in respiration and its implications. It has not been
reported whether the induction of the Rcs-phosphorelay system that controls the yjb
operon is oxygen dependent; an indirect indication that it may be, is provided by
Colón-González et al. (1999) who have shown that biofilms, regulated by the Rcssignaling pathway (Ferrier's and Clark, 2003) are not formed by E. coli K-12 during
anaerobic growth. Ryu and Beuchat (2004) have similarly shown that the production
of an EPS chemically and structurally similar to CA-EPS (Junkins and Doyle, 1992)
by enterohemorrhagic E. coli O157:H7 under aerobic growth is dramatically reduced
in a modified atmosphere containing only 1% oxygen.
A third parameter implicated in the regulation of yjb induction is cell density: activity
was highest at low culture densities and minimal at higher ones. This observation
explains why yjb induction was higher in the early stages of growth in batch culture.
The oxygen dependence data provide a simple interpretation of this phenomenon,
since lower cell densities allow for higher oxygen concentrations to be maintained in
the culture. Quorum-sensing mechanism, has been ruled out.
The yjb operon has been shown to be induced during biofilm formation (Ferrières and
Clark, 2003) and to be involved in the synthesis of an EPS different from CA-EPS
(Ferrières et al., 2007), but its physiological role in cellular metabolism and biofilm
construction has not yet been established. Based upon these findings it may be
hypothesized that whatever that role is, it would be apparent only in the early stages
of biofilm life, when cell densities are still low and oxygen concentrations are still
high.
5.9 EPS production in response to osmotic stress
Three different EPSs has been shown to be produced as part of E. coli response to
osmotic stress: poly--1,6-N-acetyl-D-glucosamine (PGA; Wang et al., 2004), CAEPS (Sledjeski and Gottesman, 1996) and yjb dependent EPS. All of these were also
linked to different extents of essentiality to biofilm formation. PGA has been shown
to be essential for biofilm formation ; CA-EPS has been shown only to be essential
for 3-dimensional architecture (Danese et al., 2000); and the yjb dependent EPS has
been yet shown not to be essential for biofilm formation although the genes are
induced during growth on solid surface (Ferrièrs and Clarke, 2003). While yjb and
wca operon are clearly co-regulated, the pga-operon belongs to a distinct NaClinduced regulon.
The question that remained unclear is why these EPS production systems are being
activated during salt stress?
It can be suggested that EPSs might have some protective effect against osmotic
shock, an idea that was shown to be true for the CA-EPS (Chen et al., 2004). Anyway,
genes which are induced by a certain stressor might be involved in protection from a
different stressor- a pattern of gene activation refers to cross regulation and protection
(Gunasekera et al., 2008). A third possibility might be the "will" of the bacterium to
go sessile. It prepares to exchange its planktonic lifestyle with biofilm. Biofilm is an
assemblage of bacterial cell attached to a surface and enclosed in adhesive EPSs. The
biofilm mode of growth undoubtedly improves survival of the organism and it is
probably the most resistant bacterial appearance which is hard to treat even with
antimicrobial agents. Goller et al. (2006) demonstrated that monovalent cation
concentrations can be an important regulatory cue for biofilm formation in E. coli,
supporting this idea but no further study has ever been done on the benefit derived
from this lifestyle during salt/osmotic stress.
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7. SUPPLEMENTARY INFORMATION
Supplementary Table 1: Microarray analysis results (section 4.1.2); 187 genes that fit both
38
criteria: induced by NaCl and repressed by .
GENE
RELATED FUNCTION
NaCl
RESPONSE
RATIO (WT)
NaCl
RESPONSE
RATIO (rpoS)
RpoS
DEPENDENCY
pflE
Predicted, Anaerobic respiration
7.3
7.6
5.5
ydeQ
Adhesion, predicted fimbria-like
protein
2.6
13.0
4.4
bssR
Biofilm formation, regulator
7.4
2.1
4.4
yibI
Hypothetical protein (NCBI
record discontinued)
7.3
31.5
4.1
gspK
Putative secretion/transport
protein
2.6
5.8
3.5
G7692
Phantom gene
5.2
3.4
3.0
abgT
Folic acid metabolism, paminobenzoyl-glutamate
transporter
6.8
2.9
2.9
ylbE-2
Predicted protein
3.0
10.2
2.9
ompL
Predicted outer membrane
protein
2.3
10.7
2.9
rutD
Nitrogen metabolism, pyrimidine
degradation
3.2
4.9
2.8
wza
CA-EPS synthesis, translocation
through the outer membrane
5.1
9.9
2.8
thiF
Biosynthesis of thiamin
2.4
8.9
2.8
wcaB
CA-EPS EPS synthesis,
predicted acyl transferase
9.1
103.0
2.7
yidK
Predicted sodium dependent
metabolites transporters
3.1
6.6
2.7
wcaD
CA-EPS synthesis, predicted
polymerase
27.3
174.0
2.6
gspJ
Putative transport protein,
pseudo-gene
2.5
43.6
2.5
allD
Nitrogen metabolism,
ureidoglycolate dehydrogenase
9.2
9.2
2.4
ypjB
Predicted protein
8.5
11.4
2.3
2.1
16.2
2.3
2.2
23.1
2.2
2.2
2.1
2.2
frlB
rhaR
ycjS
Carbohydrates catabolism,
fructoselysine 6-phosphate
deglycase
Carbohydrates catabolism, Lrhamnose catabolism,
transcription regulator
Carbohydrates catabolism,
predicted D-galactose 1dehydrogenase
ttdR
Predicted, DNA-binding
transcriptional regulator
10.3
2.6
2.2
wcaH
(gmm)
CA-EPS synthesis, GDPmannose mannosyl hydrolase
5.7
14.4
2.2
metF
Methionine biosynthesis,
methyltetrahydrofolate reductase
2.4
4.6
2.2
cynS
Cyanate catabolism, cyanate
hydratase
9.6
3.1
2.1
fsaB
(talc)
Relevance unknown, fructose 6phosphate aldolase
6.1
14.9
2.1
frwD
Carbohydrates catabolism,
fructose phosphotransferase
5.3
21.4
2.1
wcaA
CA-EPS synthesis, predicted
glycosyl transferase
3.1
9.6
2.1
yjgN
Function unknown, conserved
inner membrane protein
2.7
5.5
2.1
yibD
Predicted glycosyl transferase
2.9
4.4
2.0
wzc
CA-EPS synthesis, tyrosine
kinase
4.4
18.1
2.0
acrE
(envC)
Multi-drug efflux system,
lipoprotein
8.2
17.9
2.0
yidP
Predicted transcription regulator
2.6
6.8
2.0
hyfB
Anaerobic respiration,
hydrogenase
5.2
13.7
2.0
xylB
Carbohydrates catabolism,
xylulokinase
8.6
15.9
2.0
yadK
Adhesion, predicted fimbria-like
protein
3.6
2.9
2.0
wecH
LPS biosynthesis, enterobacterial
common antigen biosynthesis, Oacetyltransferase
8.3
9.7
2.0
ybfD
Putative DNA ligase
5.1
5.5
2.0
ydeR
Adhesion, predicted fimbria-like
protein
2.3
5.1
2.0
yrhA
Conserved protein, pseudo gene
3.9
15.9
2.0
argF
Arginine biosynthesis, ornithine
transcarbamylase
3.8
8.1
2.0
pflD
Anaerobic respiration, formate
acetyltransferase
6.0
20.1
2.0
thiH
Subunit of thiamine biosynthesis
complex
2.4
4.4
1.9
hcp
Nitrogen metabolism,
hydroxylamine reductase
4.1
4.7
1.9
bglH
Carbohydrates specific outer
membrane porin, pseudo gene
2.6
17.5
1.9
phnF
Phosphorus metabolism,
transcription regulator
12.8
10.2
1.9
yjjQ
Predicted transcription regulator
5.6
4.6
1.9
wcaC
CA-EPS synthesis, glycosyl
transferase
8.8
15.4
1.9
ycfT
Putative transport protein
4.1
8.5
1.9
rcsA
CA-EPS synthesis, transcription
regulator
5.9
12.9
1.9
ydfT
Phage related function, predicted
antiterminator
2.1
3.2
1.9
ygeK
Predicted transcription regulator
2.3
4.1
1.9
agaS
Carbohydrates catabolism,
putative tagatose-6-phosphate
aldose/ketose isomerase
5.4
12.3
1.9
ycfV
Adhesion, predicted fimbria-like
protein
3.6
6.5
1.8
yceJ
Predicted cytochrome b562
subunit
12.2
11.0
1.8
frvB
Phosphoenolpyrovate-dependent
sugar uptake system
2.2
3.1
1.8
mdtN
Predicted inner membrane
transporter protein
10.4
14.2
1.8
phnH
Phosphorus metabolism, carbonphosphorus lyase
3.2
2.9
1.8
ycbU
Adhesion, predicted fimbria-like
protein
8.5
17.7
1.8
frlC
Carbohydrates catabolism,
fructoselysine 3-epimerase
2.5
7.9
1.8
yncG
Hypothetical glutathione Stransferase-like protein
4.2
10.4
1.8
osmB
Osmotic stress response,
lipoprotein of unknown function
3.3
6.3
1.8
copA
Metals homeostasis, phosphatetype ATPase cation transporter
3.1
3.8
1.8
yjcH
Conserved inner membrane
protein
3.2
11.2
1.8
ybhI
Carbohydrates uptake,
tricarboxylate transporter
5.9
18.5
1.8
ykgC
Predicted FAD/NAD(P) binding
protein
2.0
3.7
1.8
wzxC
CA-EPS synthesis,
polysaccharide transporter
4.3
7.4
1.8
lldP
Carbohydrates catabolism,
lactate transporter
4.6
25.5
1.8
prpD
Carbohydrates catabolism, 2methylcitrate dehydratase
3.7
2.4
1.8
yqhH
Predicted outer membrane
protein
3.4
5.0
1.8
Fcl (wcaG)
CA-EPS synthesis, GDP fucose
synthase
4.0
6.2
1.8
bglF
Carbohydrates catabolism, glycoside permease
phosphoenolpyruvate-dependent
2.5
6.5
1.8
hidH
Conserved hypothetical protein
2.5
3.5
1.8
prpR
Carbohydrates catabolism,
transcription regulator of
propionate catabolism genes
4.9
5.5
1.8
alpA
Lon deficient suppressor,
transcription regulator
6.8
20.2
1.8
gadB
Acid resistance, glutamate
decarboxylase
4.1
6.9
1.8
hcaA1
Carbohydrates catabolism,
phenylpropionate dioxygenase
3.7
4.9
1.8
yiaB
Conserved inner membrane
protein
4.1
9.9
1.8
pag (crcA)
Lipopolysaccharide biosynthesis,
palmitoyl transferase for Lipid A
7.6
9.9
1.8
citG
Translation, triphosphoribosyldephospho-CoA synthase
5.5
8.3
1.8
yahH
Predicted protein
3.2
7.5
1.8
yaaX
Predicted protein
2.0
4.5
1.7
G7388
Phantom gene
18.5
12.5
1.7
yddK
Predicted protein
34.0
14.8
1.7
yiaA
Conserved inner membrane
protein
2.8
12.0
1.7
ydiN
Predicted amino acids major
facilitator superfamily transporter
3.6
5.8
1.7
tfaD
Phage related function, predicted
tail fiber assembly protein
24.6
52.1
1.7
yijF
Conserved protein
2.2
3.6
1.7
ygiZ
Conserved inner membrane
protein
3.0
7.7
1.7
yohH
Conserved protein
2.7
14.7
1.7
yjbE
EPS synthesis and biofilm
formation, predicted protein
33.2
45.3
1.7
ygaD
Conserved protein
9.5
3.1
1.7
yhaI
Putative cytochrome
2.9
18.4
1.7
yphE
Predicted ATP-dependent sugar
transporter
4.3
5.1
1.7
yafZ
Phage related function,
conserved protein
2.3
3.3
1.7
insD
Transposable element (IS2),
function unknown
2.8
3.3
1.7
bcsC
Cellulose biosynthesis, oxydase
2.3
6.2
1.7
ypdF
Glycopeptides catabolism,
aminopeptidase
6.5
6.4
1.7
nanT
Sialic acid catabolism ,sialic acid
MSF transporter
4.8
12.3
1.7
yjbH
EPS synthesis and biofilm
formation, predicted porin
2.2
3.3
1.7
wcaI
CA-EPS synthesis, predicted
glycosyl transferase
2.6
4.3
1.7
wzb
CA-EPS synthesis, tyrosine
phosphatase
3.9
5.3
1.7
ygeY
Predicted peptidase
5.3
10.7
1.7
ydeT
Predicted protein
6.2
3.1
1.7
ygaR
Predicted protein
6.6
23.8
1.7
yafW
Predicted enolase
3.2
6.8
1.7
leuO
Leucine biosynthesis,
transcription regulator
2.3
3.2
1.7
yicE
Purine catabolism,
proton/xanthine antiporter
2.0
4.2
1.6
fliR
Flagella synthesis, membrane
component of the flagella export
apparatus
2.3
4.7
1.6
gmd
CA-EPS synthesis, GDPmannose 4,6-dehydratase
5.1
7.2
1.6
ydeZ
Conserved inner membrane
heme binding protein
2.1
2.8
1.6
ydiM
Predicted proton-driven
metabolite uptake protein
6.0
4.7
1.6
yeiC
Predicted kinase
3.6
12.8
1.6
hyaA
Anaerobic respiration, subunit of
hydrogenase I
4.3
8.1
1.6
yiaK
Carbohydrates catabolism, 2,3diketo-L-gulonate reductase
3.7
22.3
1.6
yihP
Carbohydrates uptake, predicted
member of the GPH family of
galactose-pentose-hexuronide
3.7
23.6
1.6
yoaI
Predicted protein
3.0
2.0
1.6
dcuD
Predicted anaerobic expressed
transport protein
2.4
20.7
1.6
phnE
Pseudo-gene,
phosphonate/organophosphate
ester transporter
2.4
17.1
1.6
yagX
Predicted aromatic compound
dioxygenase
2.2
5.0
1.6
wzzE
LPS biosynthesis, enterobacterial
common antigen polysaccharide
chain length modulation protein
2.8
8.1
1.6
tqsA
Quorum sensing, transporter of
autoinducer 2
2.1
2.3
1.6
ybcY
Phage related function, predicted
SAM-dependent
methyltransferase
2.0
8.0
1.6
yibI
Predicted inner membrane
protein
3.1
16.6
1.6
yjbF
EPS synthesis and biofilm
formation, predicted lipoprotein
3.9
5.1
1.6
yiaN
Component of YiaMNO
transporter
3.0
3.9
1.6
wcaM
CA-EPS synthesis, hypothetical
protein
3.2
5.7
1.6
dgoK
Carbohydrates catabolism, 2dehydro-3-deoxygalactonokinase
4.8
3.4
1.6
afuC
(fbpC)
Component of a predicted ferric
transporter
3.5
6.0
1.6
yddV
Biofilm formation, 3',5'-cyclic
diguanylic acid metabolism,
diguanylate cyclase
3.9
23.5
1.6
yihQ
-glycosidase
2.2
3.4
1.6
yjbI
Conserved protein
2.1
3.9
1.6
arpB
Predicted protein
5.1
2.8
1.6
ybcW
Phage related function, predicted
protein
2.0
2.2
1.6
atoB
Carbohydrates catabolism, fatty
acid oxidation, acetyl-CoA
acetyltransferase
2.5
5.0
1.6
ybcK
Phage related function, predicted
recombinase
10.5
4.2
1.6
ttdA
Biofilm formation, L-tartrate
dehydratase enzyme
11.8
16.8
1.6
kbaY
(agaY)
Carbohydrates catabolism,
tagatose-1,6-bisphosphate
aldolase 1
5.9
19.0
1.6
yjjB
Conserved inner membrane
protein
3.8
5.8
1.6
ygjI
Predicted amino acid
transporters
2.5
6.7
1.6
yeiN
Conserved protein
2.1
2.3
1.6
yfjJ
Phage related function, predicted
protein
2.9
3.5
1.6
yiaL
Putative lipase
3.6
7.0
1.6
gabP
Carbohydrates catabolism, 4aminobutyrate transporter
2.0
18.3
1.6
yeaU
Carbohydrates catabolism, Dmalate dehydrogenase
5.6
13.0
1.6
ydfK
Phage related function,
conserved protein
2.2
4.2
1.6
yihR
Biofilm formation, predicted
aldose 1-epimerase
13.4
27.2
1.6
rffT
(wecF)
LPS biosynthesis, 4-acetoamido
4,6-D-dideoxy-D-galactose
transferase
3.6
7.4
1.6
yfdL
Phage related function,
conserved protein
2.8
4.0
1.6
ymgD
Predicted protein
3.7
5.3
1.6
yiaW
Conserved inner membrane
protein
2.9
5.8
1.6
ygcL
Predicted protein
2.8
4.5
1.5
ypjA
Adhesion, adehsin-like
autotransporter
3.7
2.6
1.5
thiD
Thiamin biosynthesis,
phospho/hydroxymethylpirimidine
kinase
3.8
7.8
1.5
yfjS
Phage related function, inner
membrane lipoprotein
4.2
10.1
1.5
actP
Acetate/glycolate permease
3.3
3.5
1.5
yiaO
Component of YiaMNO
transporter
2.6
4.2
1.5
xapA
Nucleotide metabolism,
xanthosine phosphorylase
4.2
8.4
1.5
rhtB
Homoserine lactone transporter
2.4
3.3
1.5
ygaQ
Predicted protein
10.5
17.0
1.5
yjcF
Conserved protein
7.2
4.6
1.5
eutA
Amine catabolism, activation
factor for ethanolamine ammonia
lyase
6.6
3.0
1.5
pphA
Protein missfolding stress,
phosphatase
7.1
6.7
1.5
wcaK
CA-EPS synthesis, predicted
pyruvyl transferase
4.0
6.5
1.5
yceQ
Predicted protein
2.5
2.6
1.5
yeeJ
Biofilm formation, adhesin
5.7
8.6
1.5
hyaF
Anaerobic respiration, nickel
incorporation into hydrogenase
isoenzyme I
4.8
8.0
1.5
ivy
Inhibitor of C-type lysozyme
4.6
6.1
1.5
yiaV
Predicted component of drag
transporter
2.2
5.3
1.5
yibH
Putative membrane protein
2.3
2.6
1.5
hyfE
Anaerobic respiration,
hydrogenase-IV
2.1
4.2
1.5
yfbL
Predicted peptidase
3.1
3.7
1.5
gspG
Putative secretion protein
2.4
3.4
1.5
agaC
Predicted component of Nacetylgalactoseamine PEP
phosphotrasferase system
8.2
12.6
1.5
yehC
Putative chaperone
15.9
4.0
1.5
appA
Phosphorous metabolism,
subunit of acid phosphatase
2.5
3.7
1.5
phnI
Phosphorous metabolism
(phosphonate)
2.5
3.3
1.5
pinH
Predicted invertase
6.4
4.7
1.5
hcaD
Carbohydrates catabolism,
ferredoxin NAD+ reductase
2.1
4.2
1.5
ydcU
Predicted membrane component
of spermidine/putrescine
transporter
2.1
2.9
1.5
yaiX
Predicted acyl transferase
7.2
14.6
1.5
yiaT
Putative outer membrane protein
2.9
17.0
1.5
yjhD
Phage related function, predicted
protein
2.4
2.6
1.5
ugd
CA-EPS synthesis, UDP-glucose
6-dehydrogenase
3.1
4.7
1.5
ygfS
Predicted oxidoreductase
2.7
8.8
1.5
atoD
Carbohydrates catabolism, fatty
acid oxidation, acetoacetyl-CoAtransferase
9.4
9.5
1.5
agaB
Predicted component of Nacetylgalactoseamine PEP
phosphotrasferase system
15.8
17.3
1.5
EPS synthesis
Adhesion and biofilm formation
Other membrane derived proteins (not EPS and biofilm related)
C-source catabolism
Nitrogen and phosphorus metabolism
Anaerobic respiration
Macromolecules biosynthesis
Stress related
Phage related functions
Other transcription regulators (not EPS and biofilm related)
Other functions
Supplementay Figure 1: Functional classification (annotation) of the 187 genes that fit both
38
criteria: induced by NaCl and repressed by .
Escherichia coli
!
Escherichia Na+ " Na+
coli
%EPSs$#
& $ yjbF E. coli %yjbF'::luxCDABE
" yjbEFGH yjbF(38) RpoS &
EPS
yjbF '
() 38 ' (colanic acid; CA-EPS)
EPSwca wca RT-PCR
wza’::luxCDABE
*
RcsB RcsC %$ 38 %$ RcsA yjb&, -&wca&+
38 wca yjb rpoS (RNAP)'% factors$
RNAP yjb
yjbF
E. coli 38 %E. coli (.+,- &$ 38
38 38
38 70 38
wca yjb + EPSs rpoS wca EPS yjb &yjbEFGH yjbEFGH& EPS&
%CA-EPS $ ! rpoS
+ " %yjb & wca$ ! NaCl ( (
yjbF’::luxCDABE rpoS yjbE
%yjbE$
$ wza’::luxCDABE !%lacZ’::luxCDABE
yjb 38 70*E. coli /!
%alternative$ (house keeping) ' '
" yjbF’::luxCDABE "!yjbF38 70" in-vivo yjbF
in-vitro (primer extension analysis) %$
RNAP-70yjb&
70 70 yjbF RNAP-
70 !70
in-vivo yjb
yjb 0
yjbF’::luxCDABE LB (=0.1 1/h) yjb (>0.2 1/h) - yjb
! 38 yjbF
yjbF yjb wca yjb 38 38
EPSs
38
Yjb !38
+38