REGULATION OF CQSA AND CQSS IN VIBRIO CHOLERAE

Zachary Neal Donnell

A DISSERTATION

PRESENTED TO THE FACULTY

OF PRINCETON UNIVERSITY

IN CANDIDACY FOR THE DEGREE

OF DOCTOR OF PHILOSOPHY

RECOMMENDED FOR ACCEPTANCE

BY THE DEPARTMENT OF

MOLECULAR BIOLOGY

[Advisor: Bonnie L. Bassler]

March 2015

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© Copyright by Zach Donnell, 2015. All rights reserved.

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TABLE OF CONTENTS Page number Abstract v Acknowledgements vi Chapter 1: An introduction to quorum sensing 1 Overview of quorum sensing 2

Quorum sensing in Gram-negative bacteria 3

Quorum sensing in Gram-positive bacteria 4

Quorum sensing in Vibrios 5

Feedback in V. cholerae quorum sensing 7

Chapter 2: Regulation of QS synthases and receptors in V. cholerae 16 Introduction 17 Results 17 Discussion 20 Materials and Methods 27 Chapter 3: Identification of regulators of CqsA and CqsS in V. cholerae 34 Introduction 35 Results 35 Discussion 38 Materials and Methods 47 Chapter 4: cqsA and cqsS regulation is post-transcriptional 57 Introduction 58 Results 58

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Discussion 62 Materials and Methods 71 Chapter 5: Examination of cqsA growth phase regulation 81 Introduction 82 Results 82 Discussion 86 Materials and Methods 93 References 100

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Abstract Quorum sensing (QS) is a cell-to-cell communication mechanism used by

bacteria to determine their population density and to regulate genes involved in group

behaviors. The human pathogen Vibrio cholerae contains two distinct QS systems that

converge into a shared phosphorelay pathway. The cholera-specific pathway is

composed of the autoinducer CAI-1, which is produced by the synthase CqsA and

recognized by the membrane-bound histidine kinase CqsS. The universal QS pathway is

composed of the autoinducer AI-2, which is produced by the autoinducer synthase LuxS

and binds to the receptor LuxPQ. Here, we investigate regulation of the V. cholerae

autoiducer synthases and receptors. We discovered that the LuxS/PQ system exhibits

no regulation under conditions examined, whereas the CqsA/S system contains an auto-

regulatory positive feedback loop. We characterized feedback in the CqsA/S by

identifying the Qrr sRNAs as the source of feedback, and that they act via indirect

repression of CqsA/S through an unknown intermediate. Additionally, we showed that

regulation of cqsA and cqsS occurs at the post-transcriptional level. We further

investigated growth phase regulation of cqsA, and discovered DctD as a potential

regulator of the Qrr sRNAs.

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Acknowledgements

I would like to thank my wife Jiajia Ren and the Donnell family for emotional support, my

advisor Bonnie Bassler and the Bassler lab for scientific support, and the National

Science Foundation and the National Institutes of Health for funding support.

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CHAPTER 1:

AN INTRODUCTION TO QUORUM SENSING

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Overview of quorum sensing

Quorum sensing (QS) is a cell-to-cell communication mechanism in which

bacteria produce, release, and detect chemical messages to determine their population

density (Ng and Bassler, 2009). Information regarding population size drives bacterial

group behaviors because it allows bacteria to act as individuals or as collectives. Many

behaviors bacteria engage in require group participation to be effective, and consistent

with this notion, QS regulates processes including virulence, biofilm formation, and

bioluminescence (Ng and Bassler, 2009; Waters and Bassler, 2005).

The specific chemical messages bacteria use are called autoinducers (Nealson

and Hastings, 1979). Autoinducers range in structure from small molecules to

oligopeptides, and generally serve as signals specific to a particular species.

Autoinducer concentration serves as a proxy for the bacterial population size, and

detection of autoinducers controls the transition from individual to group behaviors. A

group of bacteria are in a low cell density (LCD) state when autoinducer concentration is

low and the bacteria exhibit individualistic behaviors (e.g. prioritizing cell growth over

production of bioluminescence). A group of bacteria are in a high cell density (HCD)

state when autoinducer concentrations have reached sufficient thresholds to induce

changes in gene expression that favor group behaviors.

Bacterial QS systems fall into two broad categories: Gram-negative QS systems

and Gram-positive QS systems. Gram-negative QS systems generally use small

signaling molecules and cytoplasmic or membrane-bound receptors, whereas Gram-

positive QS systems generally use peptides as the messages coupled with membrane-

bound receptors. In addition to these two primary categories, the QS systems of many

Vibrios contain features of both Gram-negative and Gram-positive QS systems,

specifically small signaling molecules coupled with membrane-bound receptors.

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Quorum sensing in Gram-negative bacteria

The first QS system discovered was found in the marine bacterium Vibrio fischeri,

a symbiont of the bobtail squid Euprymna scolopes (Nealson et al., 1970). Initial studies

of this bacterium concerned the exclusive high cell density production of

bioluminescence via the luciferase operon (Figure 1.2). Two proteins, LuxI and LuxR,

were found to be responsible for controlling when the luciferase operon in V. fischeri was

expressed (Engebrecht and Silverman, 1984). LuxI, the autoinducer synthase, catalyzes

the production of the autoinducer N-3-(oxo-hexanoyl)-homoserine lactone (More et al.,

1996). The autoinducer acts as the ligand that activates LuxR, the master regulator of

the V. fischeri QS circuit (Engebrecht et al., 1983; Engebrecht and Silverman, 1984), and

serves to stabilize the intrinsically unstable protein. In addition to activating the genes

necessary for luciferase production, LuxR also activates the transcription of luxI, which

generates increased autoinducer. This positive feedback loop accelerates V. fischeri’s

transition from the LCD to HCD state (Nealson and Hastings, 1979).

Subsequent to the discovery of LuxI-R QS in V. fischeri, numerous other bacterial

species were shown to contain similar QS systems. For example, the QS circuit of the

opportunistic pathogen Pseudomonas aeruginosa contains two connected LuxI-R-type

QS systems (Passador et al., 1993; Pearson et al., 1994; Pearson et al., 1995). These

systems, LasI-R and RhlI-R, act in series to activate and repress the expression of

hundreds of genes in the P. aeruginosa genome, including genes necessary for toxin

production (Figure 1.3). In this case, the LasI-R system behaves like the LuxI-R system

in V. fischeri, except in addition to activating the LasI autoinducer synthase, LasR also

activates the RhlI synthase, thereby initiating a second LuxI-R type QS system (Pesci et

al., 1997). In addition to the LuxI-R type QS systems, P. aeruginosa also produces a

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third chemical signal, 2-heptyl-3-hydroxy-4-quinolone (PQS), which further links the two

canonical QS circuits (Pesci et al., 1999).

Quorum sensing in Gram-positive bacteria

Gram-positive bacteria employ oligopeptides as their autoinducers. Oligopeptides

are transcribed and translated from genes and then are processed post-translationally,

often though circularization of the peptide backbone. Gram-positive bacteria use

membrane-bound histidine-kinases to detect autoinducer concentrations. Unlike

homoserine lactone autoinducers that diffuse through the lipid bilayer, oligopeptide

autoinducers must be actively transported across the membrane via membrane-bound

transporters.

The most thoroughly studied Gram-positive QS system is that of Staphylococcus

aureus (Havarstein et al., 1995; Ji et al., 1995). The primary components of the S.

aureus QS system are encoded in the agrBDCA operon (Figure 1.4). AgrB is a

membrane-bound protein responsible for processing the autoinducer precursor AgrD by

truncation and circularization into an autoinducing peptide (AIP), and then transporting

the signal oligopeptide across the lipid bilayer. When the extracellular AIP concentration

reaches a critical threshold, it binds to the receptor AgrC. AgrC and AgrA act as a two-

component system, in which AgrC is the membrane-bound histidine-kinase that passes

phosphate to the response regulator, AgrA. AgrA activates the transcription of the

divergent promoters P2 and P3.

In addition to the species-specific autoinducer signals, a unique autoinducer has

also been demonstrated to confer cell density information between bacterial species in

both Gram-negative and Gram-positive bacteria. Autoinducer 2 (AI-2) is a collection of

molecules that share the precursor DPD, 4,5-dihydroxy-2,3-pentanedione (Schauder et

 5  

al., 2001). DPD rapidly converts between alternative configurations, and some

configurations incorporate boron into the structure, as seen in V. harveyi AI-2 (Chen et

al., 2002). AI-2 is an extremely important aspect of QS because it reveals that QS-

communication can occur across species, and the population size of one bacterial

species can affect the gene expression of another.

Quorum sensing in Vibrios

The V. harveyi QS system is a composite of the canonical Gram-positive and

Gram-negative QS systems (Ng and Bassler, 2009; Waters and Bassler, 2005). The V.

harveyi system uses small-molecule autoinducers common to Gram-negative QS

systems, but these small molecules are detected by membrane bound receptors

common to Gram-positive QS systems. The V. harveyi system also uses three parallel

QS pathways that converge through a shared phosphorelay pathway to regulate

downstream gene expression.

The first autoinducer synthase in the system, LuxM (Bassler et al., 1993; Bassler

et al., 1994a), catalyzes the production of V. harveyi autoinducer 1 (HAI-1; N-(3-

hydroxybutyryl)-homoserine lactone) (Cao and Meighen, 1989). This autoinducer is

detected by the membrane-bound receptor LuxN (Freeman et al., 2000). The second

autoinducer synthase, CqsA (Miller et al., 2002), catalyzes the production of the so-

called cholera autoinducer 1 (CAI-1; (S)-3-hydroxytridecan-4-one) (Higgins et al., 2007),

and this autoinducer is detected by the membrane-bound receptor CqsS (Miller et al.

2002). The third autoinducer synthase in this system is LuxS, which catalyzes the

production of autoinducer 2 (AI-2; (2S,4S)-2-methyl-2,3,3,4-tetrahydroxytetrahydrofuran

borate), and this autoinducer is detected by the membrane-bound receptor complex

LuxPQ (Neiditch et al., 2005; Neiditch et al., 2006). At LCD, LuxN, CqsS, and LuxPQ act

 6  

as kinases and funnel phosphate to the phosphorelay protein LuxU, which serves as the

point in the QS circuit where HAI-1, CAI-1, and AI-2 information converges (Henke and

Bassler, 2004; Miller et al., 2002). The phosphorelay protein LuxU passes phosphate to

the response regulator LuxO (Bassler et al., 1994b; Freeman and Bassler, 1999), which

binds DNA and activates the transcription of genes encoding five small RNAs (sRNAs)

termed quorum regulatory RNAs 1-5 (Qrr1-5) (Lenz et al., 2004). Qrr1-5 repress

translation of the QS master regulator LuxR by binding to and promoting degradation of

luxR mRNA (Tu and Bassler, 2007). Additionally, Qrr1-5 activate the expression of the

QS master regulator AphA by binding to the aphA transcript and inhibiting the formation

of an endogenous stem-loop that blocks AphA translation (Rutherford et al., 2011). Thus,

at LCD, AphA is produced and LuxR is repressed.

At high cell density, when autoinducers have accumulated, the autoinducers HAI-

1, CAI-1, and AI-2 bind their cognate receptors LuxN, CqsS, and LuxPQ, respectively. In

the bound state, these receptors act as phosphotases and reverse the flow of phosphate

through the phosphorelay. When LuxO is unphosphorylated, it does not activate the

transcription of qrr1-5. In the absence of Qrr1-5, luxR mRNA is translated, and LuxR

activates and represses hundreds of genes in the V. harveyi genome. Additionally, at

HCD when no Qrr sRNAs are present, aphA mRNA cannot be translated, and so no

AphA is made.

The Vibrio cholerae QS circuit is similar to that of V. harveyi, although there are a

few key differences. First, V. cholerae only possesses two parallel QS circuits (CqsA-

CqsS and LuxS-LuxPQ) as opposed to the three found in V. harveyi (Miller et al., 2002).

Second, V. cholerae contains only four Qrr sRNAs (Qrr1-4) unlike the five Qrr sRNAs in

V. harveyi. Third, Qrr1-4 in V. cholerae act redundantly to repress expression of hapR,

 7  

whereas Qrr1-5 in V. harveyi act additively to repress expression luxR. HapR and LuxR

are homologs.

Feedback in V. cholerae quorum sensing

Feedback plays a critical role in the robustness of information processing

pathways in V. cholerae. In addition to the core circuit of the V. cholerae QS system

described above, numerous feedback loops operate to optimize signal transmission

through the system. Here, the most important components of feedback in the V. cholerae

QS system are described.

Auto-repression occurs when a transcriptional regulator inhibits transcription of its

own RNA (Becskei and Serrano, 2000). Such feedback loops serve dual roles. First, the

feedback provides an upper bound on the production of the regulator. When regulator

levels reach a threshold determined by the strength of promoter binding, the regulators

bind their own promoter and occlude RNA polymerase from binding. Therefore, these

regulators turn off their own production only when a sufficient amount is already present

in a cell, reducing the dynamic range (Nevozhay et al., 2009). Second, auto-repression

dampens the noise between cells, meaning that the variation in regulator number

between cells is reduced (Paulsson, 2004). Ultimately, through the combination of these

two features, auto-repression loops serve to finely tune the concentration of the regulator

within and between cells.

In the V. cholerae QS system, three transcription factors are regulated via auto-

repression: LuxO, HapR, and AphA. LuxO, the activator of the Qrr sRNA genes, is an

NtrC-type activator, which bind DNA regardless of their phosphorylation state and

activate transcription though an ATP-driven mechanism. Although transcription factors in

this class typically act solely as activators, binding of the luxO promoter by LuxO

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represses transcription and serves to repress luxO expression because binding does not

depend on phosphorylation (Tu et al., 2010). AphA and HapR, the master regulators of

the LCD and HCD responses, respectively, are capable of activating and repressing

promoters (Rutherford et al., 2011; Svenningsen et al., 2009). Both AphA and HapR bind

their own promoters and finely tune their expression through auto-repression.

Positive feedback also exists within the V. cholerae QS circuit. HapR activates

the expression of the qrr1-4 promoters indirectly through an unknown mechanism

(Svenningsen et al., 2009), and this architecture is important for the following reasons.

First, because Qrr1-4 are responsible for degrading the hapR mRNA, HapR activation of

qrr1-4 can serve to provide another check on maximum HapR levels. Furthermore, this

circuit architecture inherently accelerates V. cholerae’s transition from the HCD to LCD

state by increasing Qrr1-4 levels upon entry into an environment lacking autoinducers.

Negative feedback occurs within the circuit as well. First, similar to HapR

activation of qrr1-4, AphA represses the expression of these same sRNA genes. AphA is

expressed most highly at LCD, when Qrr1-4 are also most highly expressed. AphA

repression of qrr1-4 serves as a check on Qrr1-4 levels. If too much Qrr1-4 are produced

in a cell, AphA levels are increased and AphA repression of qrr1-4 is enhanced, which

then lowers Qrr1-4 concentration and AphA levels. Finally, AphA and HapR have also

been shown to repress the expression of one another (Rutherford et al., 2011). This

feature enhances the differences in gene expression between LCD and HCD states. At

LCD, AphA is expressed and represses HapR. When cells begin the transition from LCD

to HCD, HapR is produced and this boosts repression of AphA. Therefore, the system

set-up promotes commitment in master regulator expression.

Finally, as mentioned above, positive feedback between QS downstream

regulators and autoinducer production (autoinduction) has been shown to be a nearly

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universal feature of QS systems in bacteria (Ng and Bassler, 2009; Shadel and Baldwin,

1991). However, no such mechanism has been demonstrated in V. cholerae. The work

in the following chapters focuses on defining the mechanism by which V. cholerae forms

positive feedback loops between QS regulators and the production of CAI-1.

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A

B

C

D

E

Figure 1.1 Quorum-sensing autoinducers

A. N-3-(oxo-hexanoyl)-HSL (V. fischeri), 3-oxo-C12-HSL (P. aeruginosa), C4-HSL (P.

aeruginosa), and N-(3-hydroxybutyryl)-HSL (V. harveyi). B. 2-heptyl-3-hydroxy-4-

quinolone (P. aeruginosa). C. Autoinducing peptide I (S. aureus). D. 4,5-dihydroxy-2,3-

pentanedione (DPD) (universal autoinducer) and (2S,4S)-2-methyl-2,3,3,4-

tetrahydroxytetrahydrofuran borate (AI-2 in Vibrios). E. (Z)-3-aminoundec-2-en-4-one (V.

harveyi) and (S)-3-hydroxytridecan-4-one (V. cholerae).

 11  

A.

B.

Figure 1.2 Quorum sensing in V. fischeri

The V. fischeri LuxI-R QS system at A. LCD and B. HCD. “AHL” represents N-3-(oxo-

hexanoyl)-homoserine lactone and the double horizontal bars represent the inner and

outer bacterial membranes.

 12  

A.

B.

Figure 1.3 Quorum sensing in P. aeruginosa

The P. aeruginosa LasI-R/RhlI-R QS systems at A. HCD for the LasI-R system and B.

HCD for the RhlI-R system. The double horizontal bars represent the inner and outer

bacterial membranes.

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A.

B.

Figure 1.4 Quorum sensing in S. aureus

The S. auerus QS system is represented at A. LCD and B. HCD. The horizontal bar

represents the bacterial membrane, and the “P” attached to AgrC and AgrA indicates

when these proteins are in the phosphorylated states. AgrD is a precursor for the

autoinducing peptide (AIP), which is processed and transported across the membrane

by AgrB. AgrC is a histidine kinase that acts as the receptor in a two component system

with the response regulator and transcription factor AgrA.

 14  

A.

B.

Figure 1.5 Quorum Sensing in V. harveyi

The QS system of V. harveyi at A. LCD and B. HCD. The double horizontal bars

represent the inner and outer bacterial membranes, “CAI-1” is (Z)-3-aminoundec-2-en-4-

one, AI-1 is N-(3-hydroxybutyryl)-homoserine lactone, and AI-2 is (2S,4S)-2-methyl-

2,3,3,4-tetrahydroxytetrahydrofuran borate, and the “P” attached to proteins indicates

when they are phosphorylated.

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A.

B.

Figure 1.6 Quorum Sensing in V. cholerae

The QS system of V. cholerae at A. LCD and B. HCD. The double horizontal bars

represent the inner and outer bacterial membranes, the “P” attached to proteins

indicates when they are phosphorylated, “CAI-1” is (S)-3-hydroxytridecan-4-one, and AI-

2 is (2S,4S)-2-methyl-2,3,3,4-tetrahydroxytetrahydrofuran borate.

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CHAPTER 2:

REGULATION OF QS SYNTHASES AND RECEPTORS IN V. CHOLERAE

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Introduction

Quorum sensing is a cell-to-cell communication process used by bacteria to

determine their population density and regulate community-dependent gene expression.

The human pathogen V. cholerae contains two parallel QS circuits that converge onto a

shared phosphorelay system. The autoinducer receptors, CqsS and LuxPQ, and their

cognate autoinducer synthases, CqsA and LuxS, control the V. cholerae QS response.

Although positive feedback from a master regulator to autoinducer production is a nearly

universal feature of bacterial QS circuits, regulation of the autoinducer synthase and

receptor genes in V. cholerae has not been explored. Here, we examined expression of

these the V. cholerae QS autoinducer synthase and receptor genes at the RNA and

protein levels in multiple QS-locked mutant backgrounds during the transition from

exponential to stationary phase. We demonstrate that positive feedback exists in the V.

cholerae QS system, but is confined only to the CqsA/CqsS QS circuit. The LuxS/LuxPQ

does not exhibit regulation under the conditions tested, suggesting a biased-feedback

circuit topology in the V. cholerae QS system.

Results

Positive feedback loops between a QS circuit master regulator and its partner

autoinducer production gene are a nearly ubiquitous feature of bacterial QS systems.

However, the existence of such a feature has not been investigated in V. cholerae. In

order to determine if the genes encoding the QS receptors or QS synthases of the V.

cholerae QS circuit are auto-regulated, we measured mRNA and protein levels from

these genes in QS mutant backgrounds locked in differing QS states. We also measured

gene expression over the course of growth, from exponential to stationary phase, to

determine if growth phase plays a role in QS gene expression.

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LuxS and LuxQ are not regulated by QS or growth phase

Previous work has shown that the LuxS AI-2 synthase and the LuxPQ AI-2

receptor are present in diverse bacterial species. Nonetheless, despite being a crucial

component of the V. cholerae QS pathway, autoregulation of these genes has not yet

been investigated. To examine if feedback in the QS pathway regulates these genes,

luxS-FLAG and luxQ-FLAG C-terminal translational fusions were generated and the

genes cloned onto plasmids. These fusions are driven by their endogenous promoters,

enabling measurement of RNA via qPCR and protein via Western blotting. The plasmids

were introduced into wild-type and QS-locked strains of V. cholerae containing the ΔvpsL

mutation to minimize biofilm formation that could interfere with optical density

measurements. The low cell density (LCD) locked V. cholerae strain contains the

phosphomimetic luxO-D47E allele that is constitutively active and continuously activates

the promoters of qrr1-4. In this configuration, Qrr1-4 levels are high, and consequently

hapR RNA is degraded, so no HapR master regulator is produced. The high cell density

(HCD) locked strain contains the ΔluxO allele, and consequently the qrr1-4 promoters

are not activated. In this configuration, because qrr1-4 are not transcribed, HapR

production is constitutive. Because the growth phase of the bacteria can also affect

expression of genes, samples were collected at multiple points throughout the LCD to

HCD transition.

Cultures were back-diluted into shaking flasks and samples were processed for

RNA and protein assessment. We first observed that there is no difference in luxS

mRNA expression among the different strain backgrounds tested, suggesting that luxS

mRNA is not QS-regulated (Figure 2.1A). Additionally, luxS mRNA levels were

consistent among all growth phases tested, suggesting that there is also no growth-

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phase regulation. LuxS protein levels were also consistent between wild-type and QS-

locked strain backgrounds (Figure 2.1B), further suggesting no QS regulation of LuxS.

Similarly, LuxS protein levels were identical regardless of growth phase.

In addition to the synthase LuxS, we also investigated QS and growth control of

LuxQ using mRNA and protein measurements in analogous experiments and strains.

Trends in luxQ RNA levels were identical to those for luxS (Figure 2.2A), indicating that

luxQ mRNA is not regulated by the V. cholerae QS circuit or by growth phase.

Additionally, LuxQ protein levels were also consistent between strain backgrounds and

between growth phases (Figure 2.2B).

CqsA and CqsS are QS and growth phase regulated

Autoregulation onto CqsA and CqsS from the QS circuit has not been

investigated. To test for regulation of the CqsA synthase and CqsS receptor, CqsA-

FLAG and CqsS-FLAG translational fusions were constructed on plasmids and

introduced into wild-type and QS-locked strains of V. cholerae. Like the LuxS-FLAG and

LuxQ-FLAG fusions mentioned earlier, the CqsA-FLAG and CqsS-FLAG fusions were

driven by their endogenous promoters.

Cultures were back-diluted into shaking flasks, and samples were isolated

throughout growth. cqsA mRNA expression increased during the transition from

exponential to stationary phase, suggesting that cqsA is growth-phase regulated (Figure

2.3A). Additionally, at the same ODs, cqsA mRNA levels differed between wild-type and

QS-locked strain backgrounds. Specifically, LCD locked strains exhibited lower than

wild-type levels of cqsA mRNA throughout the growth curve, whereas HCD locked

strains showed higher than wild-type levels of cqsA mRNA, also throughout the growth

curve. CqsA protein (Figure 2.3B) tracked with cqsA mRNA levels. Protein levels

 20  

increased in all cases from exponential to stationary phase. LCD and HCD locked strains

showed lower and higher production, respectively, compared to wild-type.

We also determined that cqsS mRNA and CqsS protein levels exhibited patterns

similar of those found for cqsA. cqsS mRNA increased in all strains during the transition

from exponential to stationary phase (Figure 2.4A). cqsS mRNA levels were higher in

HCD locked strains than in wild-type, and LCD locked strains possessed lower cqsS

mRNA than wild-type. Likewise, CqsS protein levels (Figure 2.4B) were identical to those

found for cqsS mRNA.

Discussion

The CqsA-CqsS and LuxS-LuxPQ QS systems in V. cholerae act in parallel to

regulate downstream genes responsible for individual and group behaviors (Miller et al.,

2002). Despite a clear understanding of the general topology of the circuit, specific

questions regarding feedback in these systems have not been investigated. Positive

feedback onto autoinducer production is a nearly universal feature of QS systems. This

feedback loop presumably exists to hasten the transition from the LCD to HCD state by

ensuring that bacterial populations are flooded with autoinducer after an initial increase

in autoinducer concentration occurs. Interestingly, we have shown that the architecture

of the V. cholerae QS system appears to contain biased-feedback in the parallel circuit.

We first demonstrated that the LuxS-LuxPQ QS system is neither QS nor growth

phase regulated. This feature of the QS circuit is striking because it is counter to the

ubiquitous feedback setup found in other QS systems. There are a number of potential

reasons why the V. cholerae LuxS-LuxPQ QS system does not contain feedback. First,

since LuxS is tied directly to SAM metabolism (Schauder et al., 2001), and thus, central

metabolism, positive feedback in this case may disrupt key concentrations of metabolites,

 21  

and thus needs to be avoided. A second possibility is that the LuxS-LuxPQ system lacks

positive feedback because the system cannot tolerate it. The LuxS-LuxPQ system

detects AI-2, a class of interconvertible molecules shown to be a generic bacterial QS

signal. In this case, V. cholerae may integrate bacteria-wide population density

information into it’s QS circuit, and thus amplification of AI-2 would skew its recognition

of other species in the local community.

We also demonstrated that the CqsA-CqsS system is both QS and growth phase

regulated. Unlike the LuxS-LuxPQ system, the CqsA-CqsS system follows the positive

feedback convention found in the architecture of most other QS circuits. We showed that

both mRNA and protein levels of these genes differ between locked LCD and HCD

strains, suggesting it is the mRNA that is regulated. The mRNA could be regulated at the

transcriptional or post-transcriptional level, and this will be investigated in future work.

Presumably, this feedback setup functions to rapidly push V. cholerae into a HCD state

after initial CAI-1 concentration thresholds are reached. In addition to QS feedback, both

cqsA and cqsS are regulated by growth phase. Neither gene is expressed in the wild-

type at exponential phase, but both are fully expressed at stationary phase. One

explanation for this may be that V. cholerae promotes individual behaviors if nutrient

conditions are such that rapid growth is possible, even in instances when the population

size is large.

The exact mechanism for CAI-1 feedback has not been defined at this point and

merits further investigation. The feedback could potentially operate similarly to LuxI-LuxR

systems, in which HapR could activate cqsA in order to produce more CAI-1.

Alternatively, the feedback could resemble the LuxMN positive feedback system in V.

harveyi, in which Qrr sRNAs negatively regulate luxMN expression at LCD, creating a

positive feedback loop as Qrr levels decrease when entry into HCD occurs. Future work

 22  

will focus on further delineating the specific mechanism behind feedback in the CqsA-

CqsS QS system.

 23  

A. B. Figure 2.1 LuxS is not regulated by growth-phase or QS

(A) luxS mRNA was measured by qRT-PCR and (B) LuxS protein was measured by

Western blotting in V. cholerae from a luxS-FLAG translational fusion in the ΔvpsL,

ΔvpsL luxO-D47E (locked LCD), and the ΔvpsL ΔluxO (locked HCD) strains. Samples

were isolated from the same flask at different points during growth.

 24  

A. B. Figure 2.2 LuxQ is not regulated by growth-phase or QS (A) luxQ mRNA was measured by qRT-PCR and (B) LuxQ protein was measured by

Western blotting in V. cholerae from a luxQ-FLAG translational fusion in the ΔvpsL,

ΔvpsL luxO-D47E (locked LCD), and ΔvpsL ΔluxO (locked HCD) strains. Samples were

isolated from the same flask at different points during growth.

 25  

A. B. Figure 2.3 CqsA is growth-phase and QS regulated (A) cqsA mRNA was measured by qRT-PCR and (B) CqsA protein was measured by

Western blotting in V. cholerae from a cqsA-FLAG translational fusion in the ΔvpsL,

ΔvpsL luxO-D47E (locked LCD), and ΔvpsL ΔluxO (locked HCD) strains. Samples were

isolated from the same flask at different points during growth.

 26  

A. B. Figure 2.4 CqsS is growth-phase and QS regulated

(A) cqsS mRNA was measured by qRT-PCR and (B) CqsS protein was measured by

Western blotting in V. cholerae from a cqsS-FLAG translational fusion in the ΔvpsL,

ΔvpsL luxO-D47E (locked LCD), and ΔvpsL ΔluxO (locked HCD) strains. Samples were

isolated from the same flask at different points during growth.

 27  

Materials and Methods

Bacterial strains and media

V. cholerae El Tor C6706str2 and isogenic mutant strains and E. coli S17-1λpir

(de Lorenzo and Timmis, 1994) strains were grown at 30°C in Luria-Bertani (LB) medium.

Liquid cultures were grown in flasks or test tubes with shaking for aeration. Strains used

in this study are noted in Table 2.1. Antibiotics were used at the following concentrations:

chloramphenicol 10 µg/mL, polymyxin B 50 U/mL, and streptomycin 5mg/mL. Plasmids

were electroporated into electrocompetent E. coli S17-1λpir by the MicroPulser (Bio-

Rad).

DNA manipulations and plasmids

Plasmid construction was performed by standard methods. Polymerase chain

reactions (PCR) were performed using the iProof DNA polymerase (Bio-Rad), and

restriction digestions and ligations were performed using restriction endonucleases and

T4 DNA ligase (New England Biolabs).

Plasmids pZND86, pZND87, pZND88, and pZND95 were constructed by a

similar strategy. All plasmids contain synthase or receptor promoters and ORFs PCR-

amplified from the V. cholerae El Tor C6706str2 genome. The reverse primer in each

case encodes the C-terminal FLAG fusion, and the T1 stem loop of the E. coli rrnB rho-

independent terminator (Table 2.2). Specifically, plasmid pZND86 contains 507 base

pairs upstream of the cqsS translational start site and the cqsS ORF. Plasmid pZND87

contains 500 base pairs upstream of the luxS translational start site and the luxS ORF.

Plasmid pZND88 contains 488 base pairs upstream of the luxP translational start site

and the luxP ORF and luxQ ORF. Plasmid pZND95 contains 522 base pairs upstream of

 28  

the cqsA translational start site and the cqsA ORF. PCR amplicons for pZND87, pZND88,

and pZND95 were digested with SalI and BamHI restriction endonucleases and ligated

into pJS1194 digested with the same endonucleases. The pZND86 PCR amplicon was

digested with NotI and KpnI restriction endonucleases and ligated into pJS1194 digested

with the same endonucleases. Plasmids were transformed into E. coli S17-1λpir and

mated to V. cholerae strains as described (Skorupski and Taylor, 1996).

RNA and protein isolation

Overnight cultures were back-diluted 1:1000 into fresh LB and shaken in flasks.

OD measurements were conducted periodically, and RNA and protein samples were

isolated simultaneously at OD = 0.2, 0.6, 1.0, and 1.4. For RNA samples, 5 OD units of

culture were added to 20% RNA stop solution (95% ethanol, 5% phenol), mixed by

inversion, and frozen in liquid nitrogen. Samples were stored at -80°C until processing

the following day. For protein samples, 1 OD unit of culture was immediately centrifuged

at 16,000xg for 1 minute, the supernatant was removed, and pellets were stored at

-80°C until further processing.

To isolate RNA from the samples, samples were thawed at room temperature

and centrifuged at 5000x g for 10 minutes at 4°C. Supernatants were discarded and

pellets were processed to extract RNA following the Trizol method (Tu and Bassler,

2007). To prepare protein from pellets, pellets were dissolved in Bug Buster (Millipore)

and incubated at room temperature for 20 minutes.

 29  

qRT-PCR

Following total RNA isolation, cDNA was generated from 1µg RNA using

Superscript III reverse transcriptase (Invitrogen) following the manufacturer’s guidelines.

Quantitative real time PCR (qRT-PCR) analyses were conducted in quadruplicate from

each cDNA sample using Sybr green PCR master mix (Applied Biosystems) according

to Tu & Bassler, 2007. Measurements were performed on the ABI Prism Sequence

Detection System using the ΔΔCt method. Samples were normalized to an internal rpsL

control. All primers in this analysis can be found in Table 2.2.

Western blots

Samples prepared in BugBuster solution were mixed with 0.5% SDS. In

experiments testing for CqsA-FLAG or LuxS-FLAG protein levels, the BugBuster+SDS

mix was boiled for 3 minutes before being run on an SDS-PAGE gel. In experiments

testing for CqsS-FLAG or LuxQ-FLAG protein levels, 1% Triton-X was added to help

dissolve membrane-bound dimers, and the BugBuster+SDS mix was incubated at 37°C

for 1 hour before being applied to a SDS-PAGE gel. 10µl of each sample was

electrophoresed at 150V for 2 hours on 15-well 4-15% gradient SDS-PAGE gels (Bio-

Rad). Experiments testing for CqsA-FLAG were wet blotted onto nitrocellulose

membranes, and experiments testing for CqsS-FLAG, LuxS-FLAG, and LuxQ-FLAG

were wet blotted onto PVDF membranes pre-soaked in methanol. Gels were blotted for

1 hour at 100V at 4°C. After blotting, membranes were cut in half to allow the RNA

polymerase β’ subunit control and FLAG epitopes to be probed independently. After

blotting, membranes were blocked in TBST+5% dry milk for 1 hour with shaking at room

temperature. Subsequently, α-FLAG-HRP antibody (Sigma-Aldrich) was diluted 1:5000

 30  

in TBST and incubated against the FLAG membrane for 2 hours. Simultaneously, α-RNA

polymerase β’ subunit antibody (Abcam) was diluted 1:100,000 in TBST and incubated

against the RNA polymerase β’ subunit membrane for 1 hour. The membranes testing

for α-RNA polymerase β’ subunit were rinsed with TBST and incubated with α-mouse-

HRP for 1 hour. After this incubation, both membranes containing the FLAG and α-RNA

polymerase β’ subunit were rinsed with TBST and incubated with Amersham EL Prime

Western Blotting Detection Reagent according to the manufacturer’s guidelines.

Membranes treated with the chemiluminescent reagents were visualized via film

exposure for 5 minutes.

 31  

Plasmid name Description Source pJS1194 Empty vector J. Schaffer, unpublished pZND86 pJS1194 with cqsS-FLAG This study pZND87 pJS1194 with luxS-FLAG This study pZND88 pJS1194 with luxPQ-FLAG This study pZND95 pJS1194 with cqsA-FLAG This study

Table 2.1 Plasmids used in this study

 32  

Table 2.2 Primers used in this study

Primer name Sequence Use 401 AAA AGC GGC CGC CGC TAT TTA CTC AAA CGT AAA GAG G pZND86 (insert) 404 AAA AGC GGC CGC GCA TGC AAA AAG ACC CTT CAT AAA T pZND86 (vector) 405 AAA AGT CGA CGG CGT AAA GTG GTA CTG GAA GGT C pZND87 (insert)

406 AAA AGG ATC CAA AAC GAA AGG CCC AGT CTT TCG ACT GAG CCT TTC GTT TTA CTT GTC GTC ATC GTC TTT GTA GTC GTG AAC CTT CAG CTC ATT GAG CA

pZND87 (insert)

407 AAA AGT CGA CCT TTA TCG CCG CGG TGG ATC TTG pZND88 (insert)

408 AAA AGG ATC CAA AAC GAA AGG CCC AGT CTT TCG ACT GAG CCT TTC GTT TTA CTT GTC GTC ATC GTC TTT GTA GTC ATT TAA GCC AGC GTT TTT TTG GCC

pZND88 (insert)

412 AAA AGG ATC CAA AAC GAA AGG CCC AGT CTT TCG ACT GAG CCT TTC GTT TTA CTT GTC GTC ATC GTC TTT GTA GTC ACG AAA ATA AAA ATC ACC GTA GTT GAC CG

pZND95 (insert)

419 AAA AGG TAC CAA AAC GAA AGG CCC AGT CTT TCG ACT GAG CCT TTC GTT TTA CTT GTC GTC ATC GTC TTT GTA GTC CAC CCA AGC TGC CAC TTT ATT TAG C

pZND86 (insert)

420 AAA AGG TAC CGG ATC CGG TGA TTG ATT GAG CAA GC pZND86 (vector) 422 CGT CAT CGT CTT TGT AGT C FLAG qRT-PCR 424 TCG CTT ATC ACT CAA TAG TG cqsA qRT-PCR 471 CAA AAC CTT GGC TCT GGT AC cqsS qRT-PCR 472 CGG AAT ATG TGT TAG TGA AGC luxQ qRT-PCR 473 GAT TGC GAA AAA CGT GAT TGC luxS qRT-PCR

MB51 AAA AGT CGA CTT GCG CAG CCC GAC CCG ATT C pZND95 (insert)

VC597 GGT GTT CGC TAC CAC ACA GTT rpsL forward qRT-PCR

VC598 AAG ACT TAG GAC GCT TCA CAC C rpsL reverse qRT-PCR

 33  

Strain name Organism Genotype Plasmid Source

CW2034 V. cholerae C6706str2 ΔvpsL None (Waters et al., 2008)

CW2035 V. cholerae C6706str2 ΔvpsL luxO-D47E None (Waters et al., 2008)

CW2037 V. cholerae C6706str2 ΔvpsL ΔluxO None (Waters et al., 2008)

ZDC424 V. cholerae C6706str2 ΔvpsL pZND87 This study ZDC426 V. cholerae C6706str2 ΔvpsL luxO-D47E pZND87 This study ZDC430 V. cholerae C6706str2 ΔvpsL ΔluxO pZND87 This study ZDC432 V. cholerae C6706str2 ΔvpsL pZND88 This study ZDC434 V. cholerae C6706str2 ΔvpsL luxO-D47E pZND88 This study ZDC438 V. cholerae C6706str2 ΔvpsL ΔluxO pZND88 This study ZDC450 V. cholerae C6706str2 ΔvpsL pZND86 This study ZDC454 V. cholerae C6706str2 ΔvpsL luxO-D47E pZND86 This study ZDC462 V. cholerae C6706str2 ΔvpsL ΔluxO pZND86 This study ZDC473 V. cholerae C6706str2 ΔvpsL pZND95 This study ZDC475 V. cholerae C6706str2 ΔvpsL luxO-D47E pZND95 This study ZDC479 V. cholerae C6706str2 ΔvpsL ΔluxO pZND95 This study

Table 2.3 Bacterial strains used in this study

 34  

CHAPTER 3:

IDENTIFICATION OF REGULATORS OF CQSA AND CQSS IN V. CHOLERAE

 35  

Introduction

Quorum sensing is a mechanism of cell-to-cell communication used by bacteria

to determine their population size and regulate the expression of genes involved with

group behaviors. In the Vibrio cholerae QS system, the autoinducer synthases CqsA and

LuxS catalyze the production of autoinducers CAI-1 and AI-2 which bind the autoinducer

receptors CqsS and LuxPQ. These parallel QS circuits integrate through a shared two-

component phosphorelay system. The CqsA/CqsS QS system is regulated in a manner

consistent with a positive feedback loop originating from a QS regulator. Since multiple

regulators exist in the V. cholerae QS system, we used genetic methods to identify the

regulator responsible for positive feedback onto the CqsA/CqsS system. We show that

the Qrr1-4 sRNAs are responsible for regulating CqsA, and that this regulation occurs

indirectly through an unidentified intermediate.

Results

cqsA and cqsS regulated by the Qrr sRNAs

The preceding chapter showed that cqsA and cqsS mRNA and CqsA and CqsS

protein production are different between wild-type V. cholerae and LCD and HCD-locked

mutant strains. We aimed to define the components involved in this regulation. First, we

considered known QS regulators as candidates, and we conducted epistasis

experiments. A tetracycline inducible qrr4 gene on a plasmid was introduced into four V.

cholerae mutants harboring combinations of single, double, and triple deletions of the

three known QS regulators. cqsA and cqsS mRNA were measured in the ΔluxO single

mutant, the ΔluxO ΔhapR and ΔluxO ΔaphA double mutants, and the ΔluxO ΔaphA

ΔhapR triple mutant with and without induction of qrr4 expression. cqsA expression

 36  

decreased following induction of Qrr4 in all four strains, suggesting that cqsA regulation

is independent of AphA and HapR, and thus is Qrr4-dependent (Figure 3.1). cqsS

expression modestly decreased in all strains following Qrr4 induction, however, the fold

change was not as substantial as that for cqsA (Figure 3.2).

To further verify that cqsA and cqsS are Qrr-regulated, we measured cqsA and

cqsS mRNA levels in additional QS-mutant strains. (Jian-Ping Cong, unpublished data).

cqsA and cqsS mRNA levels were measured in luxO-D47E vs. luxO-D47E Δqrr1-4

strains to verify that regulation of cqsA and cqsS is a result of Qrr regulation and not

direct LuxO regulation. cqsA and cqsS expression was higher in the luxO-D47E Δqrr1-4

strain than in the luxO-D47E strain. Thus, the Qrr sRNAs are required for cqsA and cqsS

regulation. All strains lacking HapR showed no change, verifying that cqsA and cqsS are

not regulated by HapR (not shown). Going forward, we are using cqsA as a proxy for

cqsS because cqsA regulation by Qrr4 is more dramatic than that for cqsS.

Qrr4 indirectly regulates cqsA and cqsS

The Qrr sRNAs function by binding to complementary target mRNA sequences,

generally near translational start sites and catalyzing degradation or sequestration of the

mRNAs. Because Qrr4 repressed the expression of cqsA and cqsS mRNA, we

attempted to identify sequences in or near these genes complementary to Qrr4. We first

used 5’ RACE and RNASeq to define the start and end of the cqsA and cqsS transcripts

to determine what sequences could be available for base pairing. We found that the

5’UTR of cqsA is 41bp long, and the 5’UTR of cqsS is 294bp long. We used the

bioinformatic tool RNAHybrid and visual inspection to scan for complementary

sequences between cqsA/S and Qrr4, but no complementary sequences were present.

 37  

To test for direct Qrr4 regulation of the cqsA transcript, we constructed a two-

plasmid system in E. coli, which does not contain V. cholerae QS components. One

plasmid carried tetracycline-inducible qrr4 and the second plasmid harbored an IPTG-

inducible cqsA-mKate2 translational fusion. Our rationale was that if regulation was

direct, Qrr4 would be sufficient to control expression of cqsA in E. coli. cqsA expression

did not change with increasing Qrr4 induction, indicating that cqsA is not directly

regulated by Qrr4 (Figure 3.3).

To further verify that Qrr regulation of cqsA is indirect, we conducted a Qrr4-pulse

experiment in vivo in V. cholerae to measure the timing for Qrr4 to repress cqsA

expression. Our rationale was that posttranscriptional regulation occurs rapidly, within

two minutes. Thus, if cqsA is regulated indirectly, the timescale for regulation would be

longer than what is typical for direct regulation. We pulse-induced Qrr4 in the ΔluxO

strain and measured cqsA mRNA by Northern blotting. 2-fold repression of cqsA

occurred only after 30 minutes of Qrr4 induction, suggesting that Qrr4 likely acts through

an intermediate to regulate cqsA (Figure 3.4).

A candidate screen for cqsA regulators

We reasoned that if Qrr4 acts indirectly to control cqsA, it must act through an

intermediate. Previously, a screen was conducted in V. cholerae to identify genes

regulated by Qrr4. Specifically, Qrr4 was pulse-induced, and V. cholerae gene

expression was measured by microarray analysis (Yi Shao, Table 3.1). 18 genes

identified in this microarray are candidates for regulatory intermediates acting between

Qrr4 and cqsA. If so, our expectation was that a deletion in an intermediate acting

between Qrr4 and cqsA would be incapable of transmitting information about Qrr

concentration to cqsA/S. We hypothesized that such a mutant would show a cqsA HCD

 38  

phenotype because the mutant could not repress cqsA. To test for cqsA regulation by

these 18 candidates, a plasmid containing cqsA-FLAG driven by its endogenous

promoter was introduced into V. cholerae strains each containing a deletion in one of the

candidate genes regulated by Qrr4. CqsA-FLAG expression in these strains was

compared to that in V. cholerae wild-type, locked LCD, and locked HCD strains, and six

strains exhibited cqsA expression similar to the locked HCD strain (Figure 3.5A). Three

of these strains contained deletions in hypothetical genes (VCA0935, VC1280, and

VC1323), two contained deletions in ABC transporter genes (VC1327 and VC0704), and

one contained a deletion in a galactokinase (VC1595). Because their phenotypes were

similar to wild-type, the other 12 strains were not further examined. We next introduced a

plasmid containing tetracycline-inducible qrr4 into the six HCD phenotype strains to

attempt to complement the apparent cqsA expression defects (Figure 3.5B). Our

rationale was that if the strain with a deletion in the putative intermediate remained

capable of regulation of cqsA via qrr4 induction, then the candidate gene could not, in

fact, be the correct intermediate. All strains showed decreased cqsA expression

following qrr4 induction, suggesting that Qrr4 was still capable of regulating cqsA. Thus,

none of the genes from the microarray appear to be the intermediate factor that links

Qrr4 to cqsA.

Discussion

The genes encoding the V. cholerae autoinducer synthase and receptor, cqsA

and cqsS, are activated during the transition from LCD to HCD. The preceding chapter

has demonstrated that this change in expression is controlled by QS. However, the

specific mechanism was not defined. We examined four QS genes (hapR, aphA, luxO,

and qrr4) for potential roles in cqsA and cqsS regulation. There are multiple models that

 39  

could explain how cqsA and cqsS are QS regulated. For example, cqsA/S could be

activated during the transition from LCD to HCD by HapR, which increases in

concentration from LCD to HCD. Alternatively, cqsA/S could be repressed by AphA or

Qrr1-4 at LCD, and the amount of repression would decrease during the transition from

LCD to HCD as these regulators decrease in concentration. We devised genetic

epistasis experiments to determine which components of the QS circuit influence cqsA/S

regulation. These experiments showed that the QS-based regulation of cqsA/S

originates from the Qrr sRNAs, and regulation can be controlled by exogenous

production of Qrr4. Even in mutants in which the two QS master regulators (HapR and

AphA) were deleted, cqsA/S expression was modified by addition of Qrr4.

Qrr1 and Qrr5 were previously shown to repress luxMN expression in the related

species V. harveyi by directly binding and catalyzing degradation of the mRNA (Teng et

al., 2011). We were surprised to find that although cqsA/S (a parallel synthase/receptor

pair) are also Qrr-regulated, the regulation in this case is not direct. No apparent Qrr-

binding sites could be found in either cqsA or cqsS, and we showed that a cqsA-mKate2

translational fusion is not regulated by Qrr4 in E. coli. We also showed that cqsA

repression by Qrr4 pulse-induction in V. cholerae occurs only after 32 minutes, even

though direct regulation should occur in only a few minutes. Although the V. harveyi and

V. cholerae QS circuits are highly similar, many differences have arisen and this finding

is yet another. A similar situation exists with respect to LuxR/HapR positive feedback

onto the Qrr sRNAs. LuxR directly binds the qrr1-5 promoters and activates expression

in V. harveyi, whereas HapR indirectly activates qrr1-4 though an unknown intermediate

in V. cholerae (Svenningsen et al., 2009). These results indicate that V. cholerae uses

more steps between autoregulatory loops in the QS circuit than does V. harveyi.

Possibly, such a regulatory architecture is critical for proper timing of regulation, or it

 40  

could provide additional points through which additional information from other regulatory

pathways can be integrated.

We devised a model that accounts for the indirect Qrr regulation of cqsA. We

propose two possibilities (Figure 3.6). In one scheme, Qrr1-4 activate a regulator ‘Y’ that

represses cqsA. Alternatively, Qrr1-4 represses a regulator ‘X’ that activates cqsA. In

both cases, induction of Qrr sRNAs ultimately results in repression of cqsA. Our

analyses of candidates for X and Y showed that deletions in six Qrr4-regulated genes

displayed locked cqsA HCD phenotypes similar to that of the ΔluxO strain, but in these

candidate mutants cqsA remained regulated by exogenous production of Qrr4. It is

possible that these genes may be involved in altering endogenous Qrr levels. It is also

possible that there are multiple steps between the Qrr sRNAs and cqsA, and therefore

this screen only accounts for the first step in this chain of regulation. Finally, it is also

possible that the true intermediate that acts between Qrr4 and cqsA did not change

expression strongly enough to be identified through the microarray screen, and therefore

was not included in the list of 18 candidates. In order to better understand the

mechanism that underlies Qrr4 regulation of cqsA, the future studies will examine how

cqsA is regulated using cqsA fusion constructs.

 41  

A.

B.

Figure 3.1 cqsA and cqsS are regulated by Qrr4

A. cqsA and B. cqsS mRNAs were measured by qRT-PCR in V. cholerae QS deletion

mutants. Blue bars show no qrr4 induction. 25ng/mL aTc was added to induce qrr4

expression from a tetracycline-inducible promoter on a plasmid (yellow bars). Samples

were isolated at OD600 = 1.0. Four replicates were measured in each sample; bar height

represents the mean and error bars represent standard error of the mean (SEM).

 42  

A.

B.

Figure 3.2 cqsA and cqsS are regulated by the Qrr sRNAs

A. cqsA and B. cqsS mRNAs were measured using the Quantgene plex reagent system

in V. cholerae QS mutants. Three independent measurements were taken and mean

and SEM are shown.

 43  

Figure 3.3 Qrr4 does not directly regulate cqsA

CqsA-mKate2 production was measured by FACS. Anhydrous tetracycline (aTc) was

added to LB at concentrations of 0 ng/mL, 10 ng/mL, 50 ng/mL, and 100ng/mL to induce

qrr4 expression. IPTG was added at concentrations of 0uM, 10uM, 100uM and 500uM to

induce cqsA-mKate2 expression.

 44  

Figure 3.4 Qrr4 pulse-induction regulation of cqsA

cqsA and Qrr4 mRNAs were measured by Northern blotting. A culture of V. cholerae

harboring Ptet-qrr4 was divided into aliquots at OD600 = 1.0, and 50ng/mL aTc was

added to one aliquot to induce qrr4 expression. Samples from both aliquots were

removed after 0, 2, 4, 8, 16, and 32 minutes, and RNA was isolated.

 45  

A.

B.

Figure 3.5 Screen for cqsA regulators

A. cqsA-FLAG expression was measured by Western Blotting in strains containing

deletions of 18 genes regulated by Qrr4. All samples were isolated at OD600 =1.0. B.

cqsA-FLAG expression was measured by Western Blotting in strains exhibiting locked

HCD cqsA phenotypes. “-“ indicates no Qrr4 induction by aTc and “+” indicates Qrr4

induction by 50ng/mL aTc following back-dilution from overnight cultures.

 46  

Figure 3.6 Model for Qrr regulation of cqsA in V. cholerae

The QS circuit of V. cholerae is shown. At LCD, CqsS acts as a kinase, and the

downstream phosphorelay activates the expression of the Qrr sRNAs. At HCD, CqsS

acts as a phosphatase, and the Qrr sRNAs are not activated. Qrr1-4 could feedback

onto CqsA via an intermediate “X” or “Y.”

 47  

Materials and Methods

Bacterial strains and media

V. cholerae El Tor C6706str2 and isogenic mutant strains were grown at 30°C

and E. coli S17-1λpir (de Lorenzo and Timmis, 1994) strains were grown at 37°C in

Luria-Bertani (LB) medium. Liquid cultures were grown in flasks or test tubes with

shaking for aeration. Strains used in this study are noted in Table 3.3. Antibiotics were

used at the following concentrations: chloramphenicol 10 µg/mL, ampicillin 100ug/mL.

Plasmids were electroporated into electrocompetent E. coli S17-1λpir by the MicroPulser

(Bio-Rad).

DNA manipulations and plasmids

Plasmid construction was performed by standard methods. Polymerase chain

reactions (PCR) were performed using the iProof DNA polymerase (Bio-Rad), and

restriction digestions and ligations were performed using restriction endonucleases and

T4 DNA ligase (New England Biolabs). Plasmid pZND97 was constructed by PCR-

amplifying cqsA 41bp upstream of the ORF from V. cholerae C6706str2 gDNA and the

mKate2 ORF from pZND79. The two amplicons were combined via splicing by overlap

extension (SOEing) PCR. The vector for pZND97 was created by PCR-amplifying the

backbone of pZE12. The amplicons were digested with BamHI and semi-blunt ligated

together. Plasmid pZND104 was created by amplifying the Ptet-qrr4 region on pZA-qrr4

and subcloning onto the pASK75 vector. Specifically, 54bp upstream of the qrr4

transcriptional start site though the qrr4 rho-independent terminator were amplified. The

amplicon and pASK75 were digested with AvrII and KpnI-HF and ligated together.

Plasmid pZND122 was created by amplifying the 493bp transfer origin oriT of pEVS143

 48  

and subcloning this region onto pZND104. Specifically, the oriT region was PCR-

amplified from pEVS143 and the vector backbone was amplified from pZND104, and the

amplicons were digested with KpnI-HF and XbaI-HF and ligated together. The aphA

deletion plasmid pZND46 was constructed by PCR-amplifying regions directly upstream

(769bp) and downstream (800bp) of the aphA ORF and fusing these amplicons together

by SOEing PCR. This region was subcloned onto pKAS32 by digesting the insert and

vector with NotI and EcoRI restriction endonucleases (NEB) and ligating together. All

plasmids were transformed into E. coli S17-1λpir, and V. cholerae-harbored plasmids

were mated to V. cholerae strains as described (Skorupski and Taylor, 1996).

qRT-PCR

Overnight cultures were back-diluted 1:1000 into fresh LB and shaken in flasks at

30°C, and OD measurements were conducted periodically. Cell cultures were isolated at

OD=1.0, and 5 OD units of culture were added to 20% RNA stop solution (95% ethanol,

5% phenol), mixed by inversion, and frozen in liquid nitrogen. Samples were stored at -

80°C until processing the following day. To isolate RNA from the samples, samples were

thawed at room temperature and centrifuged at 5000x g for 10 minutes at 4°C.

Supernatants were discarded and pellets were processed to extract RNA following the

Trizol method (Tu and Bassler, 2007).

Following total RNA isolation, cDNA was generated from 1µg RNA using

Superscript III reverse transcriptase (Invitrogen) following the manufacturer’s guidelines.

Quantitative real time PCR (qRT-PCR) analyses were conducted in quadruplicate from

each cDNA sample using Sybr green PCR master mix (Applied Biosystems) according

to Tu & Bassler, 2007. Measurements were performed on the ABI Prism Sequence

 49  

Detection System using the ΔΔCt method. Samples were normalized to an internal rpsL

control. All primers in this analysis can be found in Table 3.2.

Northern Blots

cqsA riboprobes were synthesized by first PCR-amplifying V. cholerae El Tor

C6706str2 gDNA template with primers in Table 3.2, and then performing T7 in vitro

transcription on the PCR amplicon (Ambion) with 32P-α-UTP. qrr4 oligoprobes were

synthesized by end-labeling primers in Table 3.2 with 32P-γ-ATP using T4 PNK (New

England Biolabs). Riboprobes and oligoprobes were purified by Illustra Microspin

columns according to the manufacturer’s guidelines (GE Healthcare). For Northern blot

experiments, an overnight culture was back-diluted 1:1000 into fresh LB and shaken in a

flask at 30°C, was grown to OD=1.0, and subsequently split into two flasks each

containing half of the volume of the original. 50ng/mL aTc was added to one flask, and

50ul of ethanol was added to the other. 5 OD units of culture were isolated from each

flask at 2, 4, 8, 16, and 32 minutes after reaching OD=1.0. RNA was Trizol extracted as

above. 10ug of total RNA was resolved on 6% polyacrylamide (PAA, 7M urea) at 300V

for 2 hours. RNA was transferred to Amersham Hybond-XL nylon membranes

(Amersham Biosciences, Piscataway, NJ) for 1 hour at 50V and at 4°C. For hybridization,

membranes were incubated overnight at 70°C for cqsA riboprobes and at 42°C for qrr4

oligoprobes in 15mL Rapid-hyb buffer (GE Healthcare, Piscataway, NJ). Following

overnight incubation, membranes were washed three times in SSC buffer + 0.1%SDS

(5X, 1X, and 0.5X at 42°C for oligoprobes; 2X, 1X, and 0.5X at 70°C for riboprobes).

Blots were imaged for 20 hours unless otherwise noted on phophorimager screens and

scanned on a Typhoon 9410 (GE Healthcare).

 50  

Direct RNA measurements

RNA amounts were measured using the Quantigene plex 2.0 reagent system

(Affymetrix, Santa Clara, CA) as described (Tu et al., 2010). Overnight cultures were

back-diluted 1:1000 into fresh LB and shaken in flasks at 30°C, and OD measurements

were conducted periodically. Cell cultures were isolated at OD=1.0, and RNA was

processed following the manufacturer’s guidelines. Fluorescent beads targeting cqsA,

cqsS were normalized to beads targeting hfq.

Western blots

Overnight cultures were back-diluted 1:1000 into fresh LB and shaken in flasks at

30°C, and OD measurements were conducted periodically. Cell cultures were isolated at

OD=1.0, and 1 OD unit of culture was immediately centrifuged at 16,000xg for 1 minute,

the supernatant was removed, and pellets were stored at -80°C until further processing.

To prepare protein from pellets, pellets were dissolved in Bug Buster (Millipore) and

incubated at room temperature for 20 minutes. Samples prepared in Bug Buster solution

were mixed with 0.5% SDS. The Bug Buster+SDS mix was boiled for 3 minutes before

being run on an SDS-PAGE gel. 10µl of each sample was electrophoresed at 150V for 2

hours on 15-well 4-15% gradient SDS-PAGE gels (Bio-Rad). Protein was wet blotted

onto nitrocellulose membranes for 1 hour at 100V at 4°C. After blotting, membranes

were cut in half to allow the RNA polymerase β’ subunit control and FLAG epitopes to be

probed independently. After blotting, membranes were blocked in TBST+5% dry milk for

1 hour with shaking at room temperature. Subsequently, α-FLAG-HRP antibody (Sigma-

Aldrich) was diluted 1:5000 in TBST and incubated against the FLAG membrane for 2

hours. Simultaneously, α-RNA polymerase β’ subunit antibody (Abcam) was diluted

 51  

1:100,000 in TBST and incubated against the RNA polymerase β’ subunit membrane for

1 hour. The membranes testing for α-RNA polymerase β’ subunit were rinsed with TBST

and incubated with α-mouse-HRP for 1 hour. After this incubation, both membranes

containing the FLAG and α-RNA polymerase β’ subunit were rinsed with TBST and

incubated with Amersham EL Prime Western Blotting Detection Reagent according to

the manufacturer’s guidelines. Membranes treated with the chemiluminescent reagents

were visualized via film exposure for 5 minutes.

mKate2 reporter assay

E. coli strains were grown aerobically overnight at 37°C in LB medium. Cultures

were back-diluted 1:1000 into M9 + 0.5% glucose and grown to OD=0.1. Upon back-

dilution, anhydrous tetracycline (aTc, Clontech) or isopropyl β-D-1-thiogalactopyranoside

(IPTG) was added to cultures at concentrations indicated in Figure 3.3. mKate2

fluorescence was measured using flow cytometry (BD Biosciences FACSAria).

 52  

Table 3.1 Plasmids used in this study

Plasmid name Description Source pASK75 Empty vector (Skerra, 1994) pEVS143 Empty vector (Dunn et al., 2006)

pKAS32 Empty vector (Skorupski and Taylor, 1996)

pZA-qrr4 Ptet-qrr4 L. Feng, unpublished pZE12 Empty vector (Levine et al., 2007) pZND104 pASK75 with Ptet-qrr4 This study pZND122 pASK75 with Ptet-qrr4 and oriT This study pZND86 pJS1194 with cqsS-FLAG This study pZND95 pJS1194 with cqsA-FLAG This study pZND97 pZE12 with Ptac-cqsA-mKate2 This study pZND46 pKAS32-ΔaphA This study

 53  

Primer name Sequence Use 263 AAA AGC GGC CGC TGC GGG TGA AGC GAT CCA AAT T pZND46 (insert)

264 GTT TGG CTT GGC CCT CTA TCT AAA CGG TTG TTG TGC T pZND46 (insert) 265 GTT TAG ATA GAG GGC CAA GCC AAA CCT GTC GAT GTA pZND46 (insert) 266 AAA AGA ATT CCT GGC CAA CCG TTT GAA CAC TG pZND46 (insert) 411 /5Phos/GTC AGC TGG CGT TAA ATT TTT TAT AAC TAG pZND97 (insert) 429 AAA AGG ATC CTG CGG CGA GCG GTA TCA GCT CA pZND97 (vector) 430 GTG CTC AGT ATC TTG TTA TCC GC pZND97 (vector)

433 AAA AGG ATC CAA AAC GAA AGG CCC AGT CTT TCG ACT GAG CCT TTC GTT TTA TCT GTG CCC CAG TTT GCT AGG G pZND97 (insert)

485 GGC CTG AGA CCA GAA TTC GAG CT pZND104 (vector)

486 AAA AAC CTA GGA GGA ATT AAT CAT CTG GCC ATT CGA TGG pZND104 (vector)

497 AAA AAC CTA GGT CCC TAT CAG TGA TAG AGA TTG ACA TCC CTA T pZND104 (insert)

498 AAA AAG GTA CCA AGA TGC TAT GGC GAA TGT GGT GAA TA pZND104 (insert)

515 AAA AAA GGT ACC GAT CCG GTG ATT GAT TGA GCA AGC TTT pZND122 (insert)

516 AAA AAA TCT AGA AGC ACC GCC AGG TGC GAA TAA G pZND122 (insert)

517 AAA AAA TCT AGA CTG GAA CAA CAC TCA ACC CTA TCT CGG pZND122 (vector)

518 GCT TAT TAA CCA CCG AAC TGC GGG T pZND122 (vector)

539 CCA GCC CAA TAC GAA TGT TT cqsA riboprobe amplicon

540 GTT TTT TTA ATA CGA CTC ACT ATA GGG AGG CAA TGA TCC CAG GAC CAT GAC G

cqsA riboprobe amplicon

KPO-0063 CGT CTA TAA GTG TGA ACA ATG GTG qrr4 oligoprobe

Table 3.2 Primers used in this study

 54  

Strain name Organism Genotype Plasmid Source

ZDE302 E. coli S17-1λpir Wild type pZND97 This study ZDE339 E. coli BW-RI Wild type pZND97 This study ZDE347 E. coli BW-RI Wild type pZND97, pZA-qrr4 This study ZDC550 V. cholerae C6706str2 ΔvpsL ΔluxO pZND95, pZND122 This study ZDC691 V. cholerae C6706str2 ΔvpsL ΔluxO ΔhapR pZND95, pZND122 This study ZDC697 V. cholerae C6706str2 ΔvpsL ΔluxO ΔaphA pZND95, pZND122 This study

ZDC693 V. cholerae C6706str2 ΔvpsL ΔluxO ΔhapR ΔaphA pZND95, pZND122 This study

ZDC546 V. cholerae C6706str2 ΔvpsL ΔluxO pZND86, pZND122 This study ZDC687 V. cholerae C6706str2 ΔvpsL ΔluxO ΔhapR pZND86, pZND122 This study ZDC695 V. cholerae C6706str2 ΔvpsL ΔluxO ΔaphA pZND86, pZND122 This study

ZDC689 V. cholerae C6706str2 ΔvpsL ΔluxO ΔhapR ΔaphA pZND86, pZND122 This study

ZDC473 V. cholerae C6706str2 ΔvpsL pZND95 This study ZDC475 V. cholerae C6706str2 ΔvpsL luxO-D47E pZND95 This study ZDC479 V. cholerae C6706str2 ΔvpsL ΔluxO pZND95 This study ZDC581 V. cholerae C6706str2 ΔVC0583 pZND95 This study ZDC582 V. cholerae C6706str2 ΔVC0704 pZND95 This study ZDC583 V. cholerae C6706str2 ΔVC0811 pZND95 This study ZDC584 V. cholerae C6706str2 ΔVC0975 pZND95 This study ZDC585 V. cholerae C6706str2 ΔVC0976 pZND95 This study ZDC586 V. cholerae C6706str2 ΔVC1188 pZND95 This study ZDC587 V. cholerae C6706str2 ΔVC1280 pZND95 This study ZDC588 V. cholerae C6706str2 ΔVC1323 pZND95 This study ZDC589 V. cholerae C6706str2 ΔVC1325 pZND95 This study ZDC590 V. cholerae C6706str2 ΔVC1327 pZND95 This study ZDC591 V. cholerae C6706str2 ΔVC1328 pZND95 This study ZDC592 V. cholerae C6706str2 ΔVC1361 pZND95 This study ZDC593 V. cholerae C6706str2 ΔVC1590 pZND95 This study ZDC594 V. cholerae C6706str2 ΔVC1594 pZND95 This study ZDC595 V. cholerae C6706str2 ΔVC1595 pZND95 This study ZDC596 V. cholerae C6706str2 ΔVC1596 pZND95 This study ZDC597 V. cholerae C6706str2 ΔVC2647 pZND95 This study ZDC598 V. cholerae C6706str2 ΔVCA0935 pZND95 This study ZDC651 V. cholerae C6706str2 ΔVC0704 pZND122, pZND95 This study ZDC653 V. cholerae C6706str2 ΔVC1280 pZND122, pZND95 This study ZDC655 V. cholerae C6706str2 ΔVC1323 pZND122, pZND95 This study ZDC657 V. cholerae C6706str2 ΔVC1595 pZND122, pZND95 This study ZDC659 V. cholerae C6706str2 ΔVCA0935 pZND122, pZND95 This study ZDC661 V. cholerae C6706str2 ΔVC1327 pZND122, pZND95 This study ZDC570 V. cholerae C6706str2 ΔvpsL ΔluxO pZND122 This study

CW2034 V. cholerae C6706str2 ΔvpsL None (Waters et al., 2008)

CW2035 V. cholerae C6706str2 ΔvpsL luxO-D47E None (Waters et al., 2008)

 55  

CW2036 V. cholerae C6706str2 ΔvpsL ΔhapR None (Waters et al., 2008)

CW2037 V. cholerae C6706str2 ΔvpsL ΔluxO None (Waters et al., 2008)

MM1162 E. coli S17-1λpir Wild type pKAS32-ΔhapR (Miller et al. 2002)

ZDE92 E. coli S17-1λpir Wild type pZND46 This study

WN778 V. cholerae C6706str2 luxO-D47E None WL Ng, unpublished

BH2126 V. cholerae C6706str2 luxO-D47E Δqrr1-4 None (Hammer and Bassler, 2007)

WN780 V. cholerae C6706str2 Δqrr1-4 None WL Ng, unpublished

SLS501 V. cholerae C6706str2 Δqrr1-4 ΔhapR None (Svenningsen et al., 2008)

Table 3.3 Bacterial strains used in this study

 56  

VC0583 hemagglutinin/protease regulatory protein, authentic frameshift Regulatory functions

VC0704

spermidine/putrescine ABC transporter, periplasmic spermidine/putrescine-binding protein, putative

Transport and binding proteins

VC0811 hypothetical protein Unknown VC0975 conserved hypothetical protein Hypothetical proteins VC0976 conserved hypothetical protein Hypothetical proteins VC1188 malate oxidoreductase Energy metabolism VC1280 hypothetical protein Unknown VC1323 hypothetical protein Unknown

VC1325 galactoside ABC transporter, periplasmic D-galactose/D-glucose-binding protein

Transport and binding proteins

VC1327 galactoside ABC transporter, ATP-binding protein Transport and binding proteins

VC1328 galactoside ABC transporter, permease protein Transport and binding proteins

VC1361 amino acid ABC transporter, permease protein Transport and binding proteins

VC1590 acetolactate synthase, catabolic Amino acid biosynthesis VC1594 aldose 1-epimerase Energy metabolism VC1595 galactokinase Energy metabolism VC1596 galactose-1-phosphate uridylyltransferase Energy metabolism VC2647 conserved hypothetical protein Hypothetical proteins VCA0935 hypothetical protein Unknown

Table 3.3 Genes studied in the microarray screen

 57  

CHAPTER 4:

CQSA AND CQSS REGULATION IS POST-TRANSCRIPTIONAL

 58  

Introduction

Quorum sensing is a cell-to-cell communication mechanism used by bacteria to

determine their population density and regulate gene expression accordingly. Vibrio

cholerae is a marine bacterium and human pathogen that contains two QS pathways

which converge into a shared phosphorelay to control the QS response. One of these

QS systems, the CqsA/S system, contains a positive feedback loop controlled by indirect

repression from the Qrr sRNAs onto CqsA/S. Here, we investigate CqsA/S repression

and show that the mechanism responsible for CqsA/S repression is post-transcriptional.

Additionally, we identify regions within the cqsA gene that could serve as potential sites

for post-transcriptional regulation.

Results

cqsA and cqsS are post-transcriptionally regulated

The Qrr sRNAs indirectly regulate the level of cqsA and cqsS RNAs in V.

cholerae. Here, we further investigate cqsA and cqsS gene expression by constructing

gene fusions to dissect the specific mechanism of regulation. We first aimed to

determine if the cqsA and cqsS mRNAs are regulated at the transcriptional or post-

transcriptional level. We constructed plasmids containing IPTG-inducible cqsA-FLAG

and cqsS-FLAG by replacing their endogenous promoters with the E. coli lacZYA

promoter (Figure 5.1). We transformed these plasmids into three V. cholerae strains that

differed in Qrr sRNA expression: a ΔvpsL strain, a ΔvpsL luxO-D47E strain, and a ΔvpsL

ΔluxO strain harboring a plasmid carrying Ptet-qrr4. Addition of aTc to the ΔvpsL ΔluxO

strain harboring Ptet-qrr4 causes qrr4 induction, allowing this strain to exhibit either a

locked LCD (induced) or locked HCD (uninduced) phenotype. cqsA-FLAG or cqsS-FLAG

 59  

expression was induced with IPTG in these strains, and CqsA-FLAG or CqsS-FLAG was

measured at three time points during growth from exponential to stationary phase. The

results show that both cqsA and cqsS expression are Qrr regulated even when driven by

the lac promoter, suggesting that regulation is post-transcriptional (Figures 4.2 and 4.3).

CqsA-FLAG and CqsS-FLAG levels are similar between ΔvpsL and ΔvpsL ΔluxO strains

when qrr4 is not induced. Conversely, CqsA-FLAG and CqsS-FLAG levels exhibit low

expression in the ΔvpsL luxO-D47E and ΔvpsL ΔluxO strains when qrr4 is induced.

To further investigate if cqsA regulation is post-transcriptional, we constructed

plasmids containing promoter fusions and transcriptional fusions linking the cqsA

promoter and 5’UTR with the gfp open reading frame (Figure 4.4). These plasmids were

introduced into a V. cholerae ΔvpsL ΔluxO strain harboring a plasmid with Ptet-qrr4. The

results show that gfp expression does not change when qrr4 is induced, suggesting that

Qrr4 does not regulate the cqsA promoter or 5’UTR. Again, this is consistent with post-

transcriptional regulation.

Screens for a cqsA regulator

We attempted to screen for regulators of cqsA and cqsS mRNA, however,

numerous problems arose. Fluorescent protein signals from V. cholerae are significantly

less bright compared to species such as V. harveyi and E. coli (Knut Drescher,

unpublished data). Although fluorescent constructs have been used successfully for

screening for V. cholerae mutants, such as Pqrr4-gfp, these constructs are generally

transcriptional fusions that are driven by strong promoters. Thus, all attempts to

construct cqsA and cqsS promoter, transcriptional, and translational fusions to

fluorescent proteins in V. cholerae failed to yield signal on FACS and in 96-well plate

 60  

readers, despite using the brightest available fluorescent proteins. Therefore, screens for

cqsA or cqsS expression using fluorescent proteins is not possible in V. cholerae.

Screening in E. coli relies on induced cqsA expression because cqsA is not expressed

from its endogenous promoter in E. coli. Additionally, the dynamic range for cqsA

expression in V. cholerae is dramatically lower when cqsA is not driven by its

endogenous promoter, which limits the usefulness of a Plac-cqsA-gfp construct in both E.

coli and V. cholerae when a large range of expression levels is needed to identify

mutants exhibiting altered expression.

For these reasons, we devised a screen that takes into account the above

experimental constraints. We constructed a plasmid containing an IPTG-inducible Plac-

cqsA-gfp fusion, and transformed it into E. coli harboring aTc-inducible V. cholerae

genomic fragments. Our rationale was that if a V. cholerae genomic fragment contained

a gene that regulates cqsA post-transcriptionally, it would either increase or decrease

GFP expression, depending on whether it activated or repressed cqsA. Despite repeated

attempts, we consistently found that the GFP expression among preliminary hits was

heterogeneous and not repeatable.

Because direct screening for cqsA regulation proved difficult, we additionally

conducted a candidate screen to attempt to identify a cqsA regulator based on the

following criteria. First, we reasoned that cqsA is likely regulated post-transcriptionally,

and second, we demonstrated that cqsA expression is decreased in an Δhfq mutant

(data not shown), suggesting a sRNA as a candidate regulator. Third, if only one

intermediate exists between the Qrr sRNAs and cqsA, the sRNA intermediate would

need to exhibit binding specificity to both qrr4 and cqsA. Fourth, if the QS control of cqsA

levels is linked to the growth phase regulation of cqsA (see chapter 2), we would expect

to identify an intermediate whose own expression changes during the transition from

 61  

exponential to stationary phase growth. Fifth, research from other groups has suggested

that CRP may indirectly regulate cqsA expression post-transcriptionally (Liang et al.,

2008), and we therefore narrowed our search to sRNAs that are regulated by glucose.

We identified two sRNA regulators that fit all of these criteria. Spot42 and VSsrna24 are

both sRNAs that regulate mRNAs post-transcriptionally, appear to be QS- and CRP-

controlled, and differ in concentration during the transition from exponential to stationary

phase growth. We investigated if these RNAs could control cqsA by expressing aTc-

inducible promoter fusions to Spot42 and VSsRNA24 and measuring cqsA mRNA levels

in V. cholerae QS mutants (data not shown). Our results show that neither Spot42 nor

VSsRNA24 affect cqsA RNA expression levels.

Determining the site of cqsA regulation.

Because we were not able to identify a gene responsible for regulating cqsA, we

attempted to identify a location on the cqsA RNA that was necessary and sufficient for

Qrr-based regulation. Our rationale was that if we could identify a sequence in cqsA

necessary and sufficient for regulation, would could use BLAST analysis of the V.

cholerae genome to identify complementary putative sRNA regulator sequences. We

first constructed IPTG-inducible gene fusions that replaced the cqsA and cqsS 5’UTRs

with the E. coli cmR (chloramphenicol resistance gene) 5’UTR to determine if the cqsA/S

5’UTR, a common target for sRNA regulation, is recognized for regulation. These

constructs failed to yield any signal when examined by Western blotting and Northern

blotting (data not shown). We also constructed IPTG-inducible gene fusions that

replaced the cqsA and cqsS ORF with the gfp ORF. Again, these fusions did not yield

consistent gfp expression by Northern and Western blotting (data not shown). We next

took a more nuanced approach to determine the site of Qrr-based regulation. We

 62  

constructed plasmids containing sequential truncations of the N- and C-termini of cqsA in

increments of 150bp to determine which region is necessary for Qrr regulation of cqsA.

These plasmids were introduced into a V. cholerae ΔvpsL ΔluxO ΔcqsA strain harboring

a plasmid with Ptet-qrr4 (Figures 4.5 and 4.6). Our rationale was that if a mutant failed to

show cqsA repression by Qrr4, then the deleted region is necessary for cqsA regulation.

All C-terminal truncation mutants showed substantially decreased basal expression

when compared to wild-type expression, but none showed substantial differences in

cqsA levels when qrr4 was induced. The N-terminal truncation mutant series showed a

different pattern of expression than for the C-terminal truncation mutants. The 150bp N-

terminal truncation showed no cqsA expression, even when Qrr4 expression was not

induced. The only mutant that was not regulated by Qrr4 was the 900bp N-terminal

truncation (Figure 4.6).

Discussion

The autoinducer-receptor pair cqsA and cqsS are regulated at the mRNA level by

QS through an indirect mechanism. We proposed that this regulation could occur

through a transcriptional method via regulation of the production of the mRNAs, or could

occur post-transcriptionally via the process of degrading existing mRNAs. We examined

this question by constructing gene fusions that separated the promoters and the open

reading frames of these genes and studying them independently. We found that both

cqsA and cqsS are regulated post-transcriptionally. This mode of feedback is contrary to

most known QS systems, which use transcriptional regulation to impose positive

feedback in the QS system. However, this mode of feedback is similar in architecture to

the luxMN system in V. harveyi that uses direct Qrr regulation of the luxMN transcript to

generate positive feedback. The cqsA/S post-transcriptional regulatory mechanism could

 63  

be either an RNA binding protein or an RNA regulator. sRNA-sRNA interactions are not

widely reported in bacteria, but have been observed often in eukaryotes in the form of

miRNA sponges (Ebert et al., 2007).

There are many possible reasons why V. cholerae uses post-transcriptional

regulation over transcriptional for cqsA/S. First, post-transcriptional regulation may show

faster dynamics than transcriptional regulation. Although the timing for Qrr feedback onto

cqsA is similar to what is expected for transcriptional regulation, the intermediate

between the Qrr sRNAs and cqsA/S may receive additional inputs that can confer a

rapid response to changing environmental conditions. Second, the V. cholerae QS

response is opposite from most pathogens in that virulence factor and biofilm production

are activated at LCD. By activating cqsA/S through a post-transcriptional mechanism,

these genes are constantly being transcribed and therefore their products are primed to

rapidly accumulate in concentration in the event that the Qrr sRNAs are depleted. Third,

since sRNAs play a large role in the QS responses of V. cholerae and are responsible

for regulating multiple QS genes, V. cholerae may have evolved use of sRNAs as the

preferred post-transcriptional regulation mechanism as a residual feature of V.

cholerae’s evolutionary history.

We also examined the cqsA transcript to determine which regions of the

transcript are necessary for the post-transcriptional repression. We discovered though

gene fusions and truncations of cqsA that the transcript is sensitive to modification.

Deletions of, or fusions to, the 3’ end of the transcript dramatically lower the basal level

of cqsA expression. Because of these observations, we hypothesize that that the 3’ end

of the cqsA transcript may be necessary for stability of the transcript. One possibility is

that this region of the transcript is somehow tied to regulation, for instance, this region

 64  

may be necessary for stabilizing the transcript, and modifications to the secondary

structure from a regulator serve to prime the transcript for degradation by exonucleases.

 65  

A.

B.

Figure 4.1 cqsA and cqsS fusion constructs

A. cqsA and B. cqsS fusions were constructed on the pEVS143 plasmid backbone.

Endogenous promoters contained ~500bp of sequence upstream of the ORF. The lac

promoters consisted of tac promoters from pEVS143 modified to the lac sequence. All

constructs contained the first rho-independent terminator from the rrnB gene of E. coli to

stop transcription.

 66  

Figure 4.2 Post-transcriptional regulation of cqsA

CqsA-FLAG expression was measured by Western blotting. V. cholerae mutants

harboring IPTG-inducible cqsA-FLAG fusions were induced with 100uM IPTG and grown

in shaking flasks. 50ng/mL aTc was added to one flask to induce Qrr4 production.

Aliquots were taken at OD600 = 0.2, 0.8, and 1.4 and processed for Western blotting.

 67  

Figure 4.3 Post-transcriptional regulation of cqsS

CqsS-FLAG expression was measured by Western blotting. V. cholerae mutants

harboring IPTG-inducible cqsS-FLAG fusions were induced with 100uM IPTG and grown

in shaking flasks. 50ng/mL aTc was added to one flask to induce Qrr4 production.

Aliquots were taken at OD600 = 0.2, 0.8, and 1.4 and processed for Western blotting.

 68  

Figure 4.4 cqsA promoter is not regulated by Qrr4

cqsA expression was measured by Northern blotting. A ΔvpsL ΔluxO ΔcqsA V. cholerae

strain harboring a plasmid with Ptet-qrr4 and a gfp-FLAG gene controlled by the cqsA

promoter was induced with 100uM IPTG and grown in shaking flasks. 50ng/mL aTc was

added to flasks to induce Qrr4 production. Aliquots were taken at OD600 = 1.0 and

processed for Northern blotting.

 69  

Figure 4.5 C-terminal cqsA truncations

cqsA expression was measured by Northern blotting. Full-length cqsA and cqsA C-

terminal truncations were expressed from the endogenous promoter on plasmids in a V.

cholerae ΔvpsL ΔluxO ΔcqsA strain harboring a plasmid with Ptet-qrr4. All samples were

measured at OD=1.0. “+” indicates Qrr4 induction with with 50ng/mL aTc, “-“ indicates no

Qrr4 induction. “Δ” indicates the bases of the cqsA ORF that are deleted.

 70  

Figure 4.6 N-terminal cqsA truncations

cqsA expression was measured by Northern blotting. Full-length cqsA and cqsA N-

terminal truncations were expressed from the endogenous promoter on plasmids in a V.

cholerae ΔvpsL ΔluxO ΔcqsA strain harboring a plasmid with Ptet-qrr4. All samples were

measured at OD=1.0. “+” indicates Qrr4 induction with with 50ng/mL aTc, “-“ indicates no

Qrr4 induction. “Δ” indicates the bases of the cqsA ORF that are deleted.

 71  

Materials and Methods

Bacterial strains and media

V. cholerae El Tor C6706str2 and isogenic mutant strains were grown at 30°C

and E. coli S17-1λpir (de Lorenzo and Timmis, 1994) strains were grown at 37°C in

Luria-Bertani (LB) medium. Liquid cultures were grown in flasks or test tubes with

shaking for aeration. Strains used in this study are noted in Table 4.3. Antibiotics were

used at the following concentrations: chloramphenicol 10 µg/mL, ampicillin 100ug/mL,

kanamycin 100ug/mL. Plasmids were electroporated into electrocompetent E. coli S17-

1λpir by the MicroPulser (Bio-Rad).

DNA manipulations and plasmids

Plasmid construction was performed by standard methods. Polymerase chain

reactions (PCR) were performed using the iProof DNA polymerase (Bio-Rad), and

restriction digestions and ligations were performed using restriction endonucleases (New

England Biolabs) and T4 DNA ligase (NEB). Phosphorylation of unphosphorylated DNA

ends was accomplished by T4 polynucleotide kinase (NEB) according to the

manufacturer’s guidelines.

Plasmid pZND109 was constructed via semi-blunt cloning by PCR-amplifying

cqsA from V. cholerae gDNA and PCR-amplifying the pEVS143 backbone vector,

digestion of amplicons with KpnI-HF, phosphorylation of DNA ends, and ligation.

pZND116 was created by PCR-amplifying the cmR resistance gene from pJS1194 and

subcloning and replacing the kanR gene this gene on pZND109. PCR amplicons were

digested with SalI-HF and AvrII and ligated together. Plasmid pZND111 was generated

by semi-blunt cloning via PCR-amplifying gfp from pEVS143 as the insert, and PCR-

amplifying a different region of pEVS143 as the vector backbone. The relevant cqsA

 72  

fragments were included in the PCR primers. Amplicons were digested with KpnI-HF,

phosphorylated, and ligated. pZND117 was constructed identically as pZND116 except

pZND111 was used in place of pZND109. Plasmid pZND120 was constructed via semi-

blunt cloning by PCR-amplifying cqsS from V. cholerae gDNA and amplifying the

pZND116 backbone as the cloning vector. Amplicons were digested with KpnI-HF,

phosphorylated, and ligated together. Plasmid pZND121 was generated via semi-blunt

cloning by PCR amplifying the gfp ORF from pEVS143 and the cqsS 5’UTR from V.

cholerae gDNA. These amplicons were combined using SOEing PCR to create the

cloning insert, and the cloning vector was generated by PCR amplifying the pZND111

backbone. The amplicons were restriction digested with KpnI-HF, phosphorylated, and

ligated together. Plasmid pZND129 was generated by PCR-amplifying 522bp of the cqsA

promoter from V. cholerae gDNA and superfolderGFP from pNUT173. These amplicons

were combined using SOEing PCR. The vector was PCR-amplified from the pZND116

plasmid backbone. The PCR fragments were restriction digested with EcoRI-HF and

KpnI-HF and subsequently ligated. Plasmid pZND130 was constructed in an identical

manner as pZND129, except the pZND130 insert primers contained sequences for a

superfolderGFP 5’UTR rather than a cqsA 5’UTR.

Plasmid pZND45 was constructed to delete a 75bp segment of the V. cholerae

genome comprising the RNA Polymerase binding region for cqsA. The 800bp regions

upstream and downstream of “TAA AAA TAA TTT TTC CCA GAT GAC AGA CGT TTT

TAA TCA CGC AAT ATA TCA CTC GTC AGC TGG CGT TAA ATT TTT” were PCR-

amplified and combined using SOEing PCR. The PCR amplicon and pKAS32 were

restriction digested with NotI-HF and EcoRI-HF and ligated together.  

Plasmids pZND145--pZND154 and pZND157 were all constructed using a similar

methodology. Primers were used to PCR-amplify the backbone of pZND95, but were

 73  

placed to allow for truncations of the cqsA ORF in multiples of 150bp. Either the forward

or reverse primer was engineered to contain a 5’ phosphate to allow for blunt-end

ligation immediately after PCR amplification. All plasmids were transformed into E. coli

S17-1λpir, and V. cholerae-harbored plasmids were mated to V. cholerae strains as

described (Skorupski and Taylor, 1996).

Northern Blots

cqsA riboprobes were synthesized by first PCR-amplifying V. cholerae El Tor

C6706str2 gDNA template with primers in Table 4.2, and then performing T7 in vitro

transcription on the PCR amplicon (Ambion) with 32P-α-UTP. qrr4, spot42, and VSsrna24

oligoprobes were synthesized by end-labeling primers in Table 4.2 with 32P-γ-ATP using

T4 PNK (New England Biolabs). Riboprobes and oligoprobes were purified by Illustra

Microspin columns according to the manufacturer’s guidelines (GE Healthcare). For

Northern blot experiments, overnight cultures were back-diluted 1:1000 into fresh LB and

shaken in test tubes at 30°C, grown to OD=1.0. 5 OD units of culture were added to 20%

RNA stop solution (95% ethanol, 5% phenol), mixed by inversion, and frozen in liquid

nitrogen. Samples were stored at -80°C until processing the following day. To isolate

RNA from the samples, samples were thawed at room temperature and centrifuged at

5000x g for 10 minutes at 4°C. Supernatants were discarded and pellets were processed

to extract RNA following the Trizol method (Tu and Bassler, 2007). 10ug of total RNA

was resolved on 6% polyacrylamide (PAA, 7M urea) at 300V for 2 hours. RNA was

transferred to Amersham Hybond-XL nylon membranes (Amersham Biosciences,

Piscataway, NJ) for 1 hour at 50V and at 4°C. For hybridization, membranes were

incubated overnight at 70°C for cqsA riboprobes and at 42°C for qrr4 oligoprobes in

 74  

15mL Rapid-hyb buffer (GE Healthcare, Piscataway, NJ). Following overnight incubation,

membranes were washed three times in SSC buffer + 0.1%SDS (5X, 1X, and 0.5X at

42°C for oligoprobes; 2X, 1X, and 0.5X at 70°C for riboprobes). Blots were imaged for 20

hours unless otherwise noted on phosphorimager screens and scanned on a Typhoon

9410 (GE Healthcare).

Western blots

Overnight cultures were back-diluted 1:1000 into fresh LB and shaken in flasks at

30°C, and OD measurements were conducted periodically. Cell cultures were isolated at

OD = 0.2, 0.8, and 1.4, and 1 OD unit of culture was immediately centrifuged at

16,000xg for 1 minute, the supernatant was removed, and pellets were stored at -80°C

until further processing. To prepare protein from pellets, pellets were dissolved in Bug

Buster (Millipore) and incubated at room temperature for 20 minutes. Samples prepared

in Bug Buster solution were mixed with 0.5% SDS. The Bug Buster+SDS mix was boiled

for 3 minutes before being run on an SDS-PAGE gel. 10µl of each sample was

electrophoresed at 150V for 2 hours on 15-well 4-15% gradient SDS-PAGE gels (Bio-

Rad). Experiments testing for CqsA-FLAG were wet blotted onto nitrocellulose

membranes, and experiments testing for CqsS-FLAG were wet blotted onto PVDF

membranes pre-soaked in methanol. After blotting, membranes were cut in half to allow

the RNA polymerase β’ subunit control and FLAG epitopes to be probed independently.

After blotting, membranes were blocked in TBST+5% dry milk for 1 hour with shaking at

room temperature. Subsequently, α-FLAG-HRP antibody (Sigma-Aldrich) was diluted

1:5000 in TBST and incubated against the FLAG membrane for 2 hours. Simultaneously,

α-RNA polymerase β’ subunit antibody (Abcam) was diluted 1:100,000 in TBST and

incubated against the RNA polymerase β’ subunit membrane for 1 hour. The

 75  

membranes testing for α-RNA polymerase β’ subunit were rinsed with TBST and

incubated with α-mouse-HRP for 1 hour. After this incubation, both membranes

containing the FLAG and α-RNA polymerase β’ subunit were rinsed with TBST and

incubated with Amersham EL Prime Western Blotting Detection Reagent according to

the manufacturer’s guidelines. Membranes treated with the chemiluminescent reagents

were visualized via film exposure for 5 minutes.

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Plasmid name Description Source pEVS143 Empty vector (Dunn et al., 2006) pJS1194 Empty vector J. Schaffer, unpublished pKAS32 Empty vector (Skorupski and Taylor, 1996) pZND109 pEVS143 with Plac-cqsA-FLAG This study pZND111 pEVS143 with Plac-cqsA5'UTR-gfp-FLAG This study pZND116 pEVS143 with Plac-cqsA-FLAG This study pZND117 pEVS143 with Plac-cqsA5'UTR-gfp-FLAG This study pZND120 pEVS143 with Plac-cqsS-FLAG This study pZND121 pEVS143 with Plac-cqsS5'UTR-gfp-FLAG This study pZND122 pASK75 with Ptet-qrr4 and oriT This study pZND123 pEVS143 with Plac-CmR5'UTR-cqsA-FLAG This study pZND124 pEVS143 with Plac-CmR5'UTR-cqsS-FLAG This study pZND129 pEVS143 with PcqsA-cqsA5'UTR-gfp-3xFLAG This study pZND130 pEVS143 with PcqsA-gfp5'UTR-gfp-3xFLAG This study pZND143 pEVS143 with Ptac-spot42 K. Papenfort, unpublished pZND144 pEVS143 with Ptac-VSsrna24 K. Papenfort, unpublished pZND145 pZND95 cqsA C-terminal truncation (150bp) This study pZND146 pZND95 cqsA C-terminal truncation (300bp) This study pZND147 pZND95 cqsA C-terminal truncation (450bp) This study pZND148 pZND95 cqsA C-terminal truncation (600bp) This study pZND149 pZND95 cqsA C-terminal truncation (750bp) This study pZND150 pZND95 cqsA C-terminal truncation (1050bp) This study pZND151 pZND95 cqsA N-terminal truncation (150bp) This study pZND152 pZND95 cqsA N-terminal truncation (300bp) This study pZND153 pZND95 cqsA N-terminal truncation (450bp) This study pZND154 pZND95 cqsA N-terminal truncation (600bp) This study pZND157 pZND95 cqsA N-terminal truncation (900bp) This study pZND45 pKAS32-ΔPcqsA This study pZND95 pJS1194 with cqsA-FLAG This study

Table 4.1 Plasmids used in this study

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Primer name Sequence Use 468 GTC AGC TGG CGT TAA ATT TTT TAT AAC TAG G pZND109 (insert)

463

AAA AAG GTA CCA AAA CGA AAG GCC CAG TCT TTC GAC TGA GCC TTT CGT TTT ACT TGT CGT CAT CGT CTT TGT AGT CAC GAA AAT AAA AAT CAC CGT AGT TGA CCG pZND109 (insert)

469 AAA AAG GTA CCG ATC CGG TGA TTG ATT GAG CAA GC

pZND109, pZND111, pZND121 (vector)

464

GTG CTC AAC ATA TTG TTA TCC GCT CAC AAT GTA AAT TGT TAT CCG CTC ACA ACA GCT CAT TTC AGA ATA TTT GCC AGA ACC

pZND109, pZND111, pZND120, pZND121 (vector)

523 AAA AAA GTC GAC GAA GAT GCG TGA TCT GAT CCT TCA ACT C pZND116 (insert)

524 AAA AAA CCT AGG CGT TGT GTC TCA AAA TCT CTG ATG TTA CAT TGC pZND116 (insert)

525 AAA AAA CCT AGG ACT CGC TAC GCT CGG TCG TTC GAC T pZND116 (vector) 526 AAA AAA GTC GAC ATC ACG CAT CTT CCC GAC AAC GCA GA pZND116 (vector)

466 GTC AGC TGG CGT TAA ATT TTT TAT AAC TAG GAT ATA TTG CGA TGG CTA GCA AAG GAG AAG AAC TCT TC pZND111 (insert)

467

AAA AAG GTA CCA AAA CGA AAG GCC CAG TCT TTC GAC TGA GCC TTT CGT TTT ACT TGT CGT CAT CGT CTT TGT AGT CGT TGT ACA GTT CAT CCA TGC CAT GTG TAA TC

pZND111, pZND121 (insert)

501 GAT GAA GGT TTT GGC AGT TTG GAT CCG pZND120 (insert) 519 GTG CTC AAC ATA TTG TTA TCC GCT CAC A pZND120 (vector)

474

AAA AAG GTA CCA AAA CGA AAG GCC CAG TCT TTC GAC TGA GCC TTT CGT TTT ACT TGT CGT CAT CGT CTT TGT AGT CCA CCC AAG CTG CCA CTT TAT TTA GC pZND120 (insert)

522 GAT GAA GGT TTT GGC AGT TTG GAT CCG G pZND121 (insert)

502 TCC TTT GCT AGC CAT GCT CAC TAT CAC TAC CGT TGC ATT CTC pZND121 (insert)

503 GTA GTG ATA GTG AGC ATG GCT AGC AAA GGA GAA GAA CTC TTC pZND121 (insert)

527 TTT TGA ATT CTT GCG CAG CCC GAC CCG ATT CT pZND129, pZND130 (insert)

531 TTC TCC TTT GCT CAT CGC AAT ATA TCC TAG TTA TAA AAA ATT TAA CGC CA pZND129 (insert)

532 CTA GGA TAT ATT GCG ATG AGC AAA GGA GAA GAA CTT TTC ACT GG pZND129 (insert)

533

AAA AGG TAC CAA AAC GAA AGG CCC AGT CTT TCG ACT GAG CCT TTC GTT TTA CTA TTT ATC GTC ATC TTT GTA GTC GAT ATC ATG ATC TTT ATA ATC ACC GTC ATG GTC TTT GTA GTC TTT GTA GAG CTC ATC CAT GCC ATG TGT AAT C

pZND129, pZND130 (insert)

529 TTT TGG TAC CGA TCC GGT GAT TGA TTG AGC pZND129, pZND130 (vector)

530 TTT TGA ATT CCA GAA CCG TTA TGA TGT CGG CGC pZND129, pZND130 (vector)

534 ATC TCC TTA ACT AGG GAG TGA TAT ATT GCG TGA TTA AAA ACG TCT GTC pZND130 (insert)

535 CGC AAT ATA TCA CTC CCT AGT TAA GGA GAT ATA CAT ATG AGC AAA GGA pZND130 (insert)

271 AAA AGC GGC CGC CTA TGC GAC TAT CTC GCG ATT C pZND45 (insert)

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272 CAA TAT ATC CTA GTT ATA TTA GCA ACT TAG AAT AAT TAA TAA TTC TCG ATG pZND45 (insert)

273 TT ATT CTA AGT TGC TAA TAT AAC TAG GAT ATA TTG CGA TGA pZND45 (insert)

274 AAA AGA ATT CGG AAC GCA GCG ATT CAC TTC AT pZND45 (insert)

576 GTG TTT GGC TCA GTA TTC TGC CGC cqsA 3' region riboprobe

577 GTT TTT TTA ATA CGA CTC ACT ATA GGG AGG ACG AAA ATA AAA ATC ACC GTA GTT GAC CGC

cqsA 3' region riboprobe

567 GTC AGC TGG CGT TAA ATT TTT TAT AAC TAG G cqsA 5' region riboprobe

568 GTT TTT TTA ATA CGA CTC ACT ATA GGG AGG CTT GTT TAC CCA ATA CAA GGT GTT TAC CG

cqsA 5' region riboprobe

541 CTG TCC GTG GAG AGG GTG AA superfolderGFP riboprobe

542 GTT TTT TTA ATA CGA CTC ACT ATA GGG AGG ATC CGG ATA ACG GGA AAA GC

superfolderGFP riboprobe

539 CCA GCC CAA TAC GAA TGT TT cqsA riboprobe

540 GTT TTT TTA ATA CGA CTC ACT ATA GGG AGG CAA TGA TCC CAG GAC CAT GAC G cqsA riboprobe

Table 4.2 Primers used in this study

 79  

Strain name Organism Genotype Plasmid Source

CW2034 V. cholerae C6706str2 ΔvpsL None Waters et. al, 2008

CW2035 V. cholerae C6706str2 ΔvpsL luxO-D47E None Waters et. al, 2008

CW2037 V. cholerae C6706str2 ΔvpsL ΔluxO None Waters et. al, 2008

ZDC621 V. cholerae C6706str2 ΔvpsL pZND116 This study ZDC623 V. cholerae C6706str2 ΔvpsL pZND120 This study ZDC631 V. cholerae C6706str2 ΔvpsL luxO-D47E pZND116 This study ZDC633 V. cholerae C6706str2 ΔvpsL luxO-D47E pZND120 This study ZDC641 V. cholerae C6706str2 ΔvpsL ΔluxO pZND122, pZND116 This study ZDC643 V. cholerae C6706str2 ΔvpsL ΔluxO pZND122, pZND120 This study ZDC702 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA None This study ZDC757 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND95, pZND122 This study ZDC775 V. cholerae C6706str2 ΔvpsL pZND143 This study ZDC777 V. cholerae C6706str2 ΔvpsL luxO-D47E pZND143 This study ZDC779 V. cholerae C6706str2 ΔvpsL ΔluxO pZND143, pZND122 This study ZDC811 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND122, pZND129 This study ZDC823 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND122, pZND130 This study ZDC839 V. cholerae C6706str2 ΔvpsL pZND144 This study ZDC841 V. cholerae C6706str2 ΔvpsL luxO-D47E pZND144 This study ZDC845 V. cholerae C6706str2 ΔvpsL ΔluxO pZND144, pZND122 This study ZDC853 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND145 This study ZDC855 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND146 This study ZDC857 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND147 This study ZDC859 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND148 This study ZDC861 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND149 This study ZDC863 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND150 This study ZDC865 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND151 This study ZDC869 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND152 This study ZDC871 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND153 This study ZDC873 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND154 This study ZDC875 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND157 This study ZDE244 E. coli S17-1λpir Wild type pZND95 This study ZDE362 E. coli S17-1λpir Wild type pZND122 This study ZDE429 E. coli S17-1λpir Wild type pZND116 This study ZDE433 E. coli S17-1λpir Wild type pZND120 This study ZDE497 E. coli S17-1λpir Wild type pZND129 This study ZDE499 E. coli S17-1λpir Wild type pZND130 This study

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ZDE540 E. coli S17-1λpir Wild type pZND145 This study ZDE542 E. coli S17-1λpir Wild type pZND146 This study ZDE544 E. coli S17-1λpir Wild type pZND147 This study ZDE546 E. coli S17-1λpir Wild type pZND148 This study ZDE548 E. coli S17-1λpir Wild type pZND149 This study ZDE550 E. coli S17-1λpir Wild type pZND150 This study ZDE552 E. coli S17-1λpir Wild type pZND151 This study ZDE589 E. coli S17-1λpir Wild type pZND152 This study ZDE590 E. coli S17-1λpir Wild type pZND153 This study ZDE592 E. coli S17-1λpir Wild type pZND154 This study ZDE90 E. coli S17-1λpir Wild type pZND45 This study

Table 4.3 Bacterial strains used in this study

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CHAPTER 5:

EXAMINATION OF CQSA GROWTH PHASE REGULATION

 82  

Introduction

Quorum sensing is a communication process used by bacteria to assess their

population size and regulate the expression of genes involved in group behaviors. The

QS system of the human pathogen V. cholerae operates through the production of

autoinducers by the synthases CqsA and LuxS, and detection of autoinducers by the

receptors CqsS and LuxPQ, respectively. Previous chapters have shown that the

autoinducer synthase gene cqsA is not transcribed in E. coli under the native cqsA

promoter, suggesting that an activator of cqsA may exist in V. cholerae, but not in E. coli.

Here, we screen for activators of cqsA and examine candidates for their roles in V.

cholerae QS. This screen identifies multiple genes with potential roles in regulation of the

Qrr sRNAs.

Results Screens for a cqsA transcriptional activator

We have previously shown that cqsA is not expressed from its endogenous

promoter in E. coli. However, cqsA expression can be detected in E. coli when cqsA is

expressed from an IPTG-inducible promoter. We hypothesized that this discrepancy

could be explained if a transcriptional activator of cqsA exists in V. cholerae, but not in E.

coli. In order to test this hypothesis, we devised two screens to isolate a cqsA

transcriptional activator.

In the first screen, we constructed an E. coli strain harboring a PcqsA-gfp

transcriptional fusion. The cells carrying this fusion were dark since this promoter is not

expressed in E. coli. We next introduced a V. cholerae genomic library under an aTc-

inducible Ptet promoter into this E. coli strain. Our rationale was that if a V. cholerae

genomic fragment contained an activator of the cqsA promoter, GFP expression would

 83  

be induced and the cell would be bright. After multiple rounds of FACS sorting and

measurements on a 96-well plate reader, eleven clones were isolated and retested, but

ultimately none consistently exhibited GFP expression above background levels.

In the second screen, we constructed a V. cholerae Tn5 transposon library

carrying a Pqrr4-gfp promoter fusion. We conducted the screen in a V. cholerae ΔluxQ

background to eliminate qrr4 activation that would stem from the AI-2 signaling pathway.

We used a Pqrr4-gfp fusion rather than the PcqsA-gfp fusion because the PcqsA-gfp

fusion does not fluoresce in V. cholerae. Our rationale was that clones expressing cqsA

would be dark because these clones would be in HCD mode and thus, repress the qrr4

promoter. Transposon mutants deficient in cqsA activation caused by transposon

insertion would, by contrast, be in LCD mode, and the qrr4 promoter would be activated,

causing GFP to be expressed. We first confirmed that the pattern of GFP expression

from Pqrr4-gfp matched our expectations of high GFP using a cqsA null allele (Figure

5.1) Subsequently, we screened the V. cholerae Tn5 library and isolated seven targets

from the initial screen, and reconfirmed these clones using a 96-well plate reader assay

and FACS. We found that four of the seven transposon mutants showed a fluorescence

signal similar to that of a ΔluxQ ΔcqsA control strain (Figure 5.2), and we sequenced the

insertion sites in these mutants.

The transposon insertions were located inside three ORFs: VCA0033, VC0122,

and VC1926 (two separate insertions). VCA0033 is a hypothetical gene that has been

suggested to show homology to phoA (Majumdar et al., 2005). VC0122 encodes one of

the two V. cholerae adenylate cyclase genes, cya. VC1926 encodes the gene dctD, a

NtrC-class activator and two-component system response regulator. NtrC-class

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activators bind far upstream of their target promoters and activate transcription through

DNA looping and ATPase activity, and they interact with the alternative sigma factor σ54.

In previous chapters, we have demonstrated that the Qrr sRNAs repress cqsA

through a feedback loop, and therefore mutations that alter Qrr levels can affect cqsA

mRNA levels. Because our experimental system relied on measuring signal from the

qrr4 promoter and not from cqsA itself, we further investigated the mutants from the

present screen by directly measuring RNA levels of both cqsA and qrr4 at exponential

and stationary phases (Figure 5.3). We found that at exponential phase (OD600 = 0.1),

the VCA0033 and VC1926 mutants contained higher cqsA mRNA levels than wild-type,

whereas the VC0122 mutant showed cqsA mRNA levels identical to wild-type. The

ΔrpoN (encodes σ54) mutant was examined because of its connection to VC1926, and

because it is required for LuxO regulation of Qrr sRNA expression. This ΔrpoN mutant

showed very strong cqsA expression. The ΔluxQ ΔcqsA control strain showed no cqsA

expression as expected. All mutants from the screen showed Qrr4 levels similar to wild-

type. The ΔrpoN mutant had no Qrr4 expression since this mutant lacks the ability to

transcribe the Qrr sRNAs. The ΔluxQ ΔcqsA control had the strongest Qrr4 production

since it is locked at LCD. These results alone suggest that VCA0033 and VC1926 may

be affecting cqsA directly as there is no change in Qrr4 levels at exponential phase.

However, the direction of cqsA expression change is opposite of the expectation

(mutations in genes responsible for activation of cqsA should lower cqsA expression).

At stationary phase (OD600 = 1.0) all mutants from the screen and controls

(except the ΔluxQ ΔcqsA mutant) had nearly identical cqsA mRNA levels. Conversely,

many strains differed in Qrr4 levels. All screen mutants showed Qrr4 expression at

stationary phase, suggesting that these mutants lock the V. cholerae QS circuit in a LCD

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configuration since Qrr sRNAs in wild type are undetectable shortly after OD600 = 0.1.

These results suggest that none of the mutants identified in the screen altered the levels

of cqsA expression directly, but instead caused qrr4 expression to remain activated

through entry into stationary phase. Since the screen measured qrr4-GFP levels, it

appears that all of the bright mutants from the screen were the result of Qrr4 alterations

at stationary phase rather than activation of cqsA directly.

Investigating an inverted repeat in the cqsA promoter

We originally hypothesized that if DctD regulates cqsA, its activation mechanism

would follow that of conventional NtrC-class activators (cqsA would be controlled by σ54

and an inverted repeat would be located around 120bp upstream of the cqsA

transcriptional start site). We scanned the region upstream of the cqsA transcriptional

start site by visual inspection and identified an inverted repeat centered at -118bp

(Figure 5.4). We further investigated the inverted repeat upstream of cqsA to assess

whether this genomic feature regulates cqsA in conjunction with other transcriptional

regulators. We tested if the inverted repeat was necessary for cqsA expression by

constructing plasmids that contained the cqsA ORF and 500bp of upstream sequence.

We engineered two mutants, one containing a deletion of the inverted repeat region, and

the other containing scrambled nucleotides in place of the inverted repeat region. These

mutants showed no difference in cqsA expression levels from wild-type, suggesting that

the inverted repeat region does not control cqsA under the conditions tested (data not

shown).

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Discussion

In this study we screened for a transcriptional activator of cqsA. There are a

number of reasons why a cqsA activator may exist, but was not discovered though our

screens. First, if the activation mechanism relies on multiple genes, the screens

conducted here may not be able to identify the activator. Second, transposon insertion of

Tn5 may be biased and not perfectly random, and insertion into the activator site may

not have been achieved. Third, in the case of the PcqsA-gfp reporter used in E. coli, we

were not able to construct a positive control in V. cholerae, so it is not confirmed if this

reporter is capable of cqsA-GFP expression and fluorescence.

Regulation of qrr4

Although our screens were designed to identify regulators of cqsA, because we

used a qrr4-GFP reporter as the readout, we identified potential regulators of qrr4. It

appears from these data that DctD is responsible for repression of Qrr4. This proposed

repression mechanism is contrary to DctD mechanisms that have been reported in

rhizobia (Yurgel and Kahn, 2004) and it differs from NtrC-class activators in general

(Tucker and Sallai, 2007). Previous studies of this family of proteins have shown them to

exclusively behave as transcriptional activators. In this case, the VC1926 allele may

function differently than canonical NtrC-class proteins and could have acquired new

functionality. V. cholerae contains two separate DctBD systems (VC1925/1926, and

VCA0141/0142), and perhaps one of these paralogs behaves as the “true” DctD, while

the system identified in the present screen has evolved a repression mechanism. We

observed that cqsA levels are higher in a ΔrpoN mutant than in wild-type, however, this

likely due to the fact that a ΔrpoN mutant locks the QS circuit into the LCD state via

 87  

inhibition of Qrr sRNA transcription (similar in phenotype to a ΔluxO mutant). Thus,

upregulated cqsA is due to lack of repression of the Qrr sRNAs. The role of DctD in Qrr

sRNA production may also be important for the study of Qrr sRNAs in E. coli, as E. coli

lacks the DctBD system, and the target gene of the DctBD system. DctA is controlled

instead by the DcuR-DcuS regulatory system in E. coli, which is not homolgous to DctBD.

Qrr sRNAs are frequently studied outside of V. cholerae in E. coli to isolate the QS

system from feedback and external regulation, and doing so could ignore the effects of a

potentially important feature of the Qrr sRNA regulatory system.

The VC0122 and VCA0033 mutants also cause upregulation of Qrr4 at HCD.

Similar to the DctBD system discussed above, VC0122 encodes one of two cya paralogs

in V. cholerae. Although glucose metabolism has been implicated in cqsA regulation

previously (Liang et al., 2008), the role that glucose plays in cqsA regulation remains

unclear. We have shown that Δcrp mutants are incapable of activating cqsA, however,

Δcya mutants and glucose induction do not yield significant changes in cqsA mRNA

levels (data not shown). Although the VC0122 mutant shows higher than wild type

expression of Qrr4 at HCD, this difference is not as substantial as the effect for VC1926

and VCA0033. Furthermore, although there have been reports that VCA0033 is the V.

cholerae phoA, PhoA should not be expressed in our experimental conditions. It is

currently unclear the role VCA0033 plays in Qrr sRNA regulation, especially since this

gene does not display any clear homology to any previously studied genes. We leave

this as an avenue for further research.

 88  

Figure 5.1 Pqrr4-GFP expression in V. cholerae mutants

Pqrr4-GFP expression was measured using a 96-well plate reader. Histogram bars

indicate the mean GFP fluorescence of three measurements of V. cholerae cultures

divided by the optical density of each culture.

 89  

Figure 5.2 Pqrr4-GFP expression in V. cholerae screen hits

Pqrr4-GFP expression in V. cholerae cells was measured by FACS. Y-axis indicates

bacterial count and X-axis indicates FITC intensity. The “controls” indicate GFP

expression from wild type V. cholerae, a positive control (ΔluxQ ΔcqsA), and a negative

control (a screen mutant that did not yield high GFP expression). “Screen hits” indicate

GFP expression from mutants that were screened twice and yielded high GFP

expression in 96-well plate reader assays.

 90  

Figure 5.3 RNA expression of screen hits and controls

cqsA mRNA and Qrr4 levels were measured by Northern blotting. Aliquots from cultures

were taken and processed at OD600=0.1 and OD600=1.0.

 91  

Figure 5.4 An inverted repeat exists in the cqsA promoter

Asterisks indicate exact matches for an inverted repeat centered at 118bp upstream of

the cqsA ORF. Orange bars indicate the inverted repeat region, as well as a region

downstream that shares similarity to the 3’ side of the inverted repeat. The red arrow

indicates the 41bp cqsA 5’UTR, and the green bar indicates the N-terminal region of the

cqsA ORF.

 92  

Materials and Methods

Bacterial strains and media

V. cholerae El Tor C6706str2 and isogenic mutant strains were grown at 30°C

and E. coli S17-1λpir (de Lorenzo and Timmis, 1994) and E. coli TOP10 (Invitrogen)

strains were grown at 37°C in Luria-Bertani (LB) medium. Liquid cultures were grown in

flasks or test tubes with shaking for aeration. Strains used in this study are noted in

Table 5.3. Antibiotics were used at the following concentrations: chloramphenicol 10

µg/mL, ampicillin 100ug/mL, kanamycin 100ug/mL. Plasmids were electroporated into

electrocompetent E. coli S17-1λpir and E. coli TOP10 by the MicroPulser (Bio-Rad).

DNA manipulations and plasmids

Plasmid pZND165 was constructed by PCR-amplifying the pZND95 backbone

with primers designed to amplify the DNA flanking the inverted repeat region containing

“TTC GGG GTA GAG TCC CTA CCC CTA A”. The amplicon was phosphorylated and

blunt self-ligated. Plasmid pZND166 was constructed in a similar manner, except that

one of the PCR primers contained the scrambled inverted repeat sequence to replace

the inverted repeat region above. pZND136 was constructed by PCR-amplifying the

backbone of pSLS4, except for the region containing kanR. The cmR region of pJS1194

was PCR-amplified, and the amplicons were restriction digested with NotI-HF and AvrII

and ligated together.

pZND130 was generated by first PCR-amplifying 486bp of the cqsA promoter

upstream of the transcriptional start site. The superfolderGFP ORF and ribosome

binding site (including 21bp upstream of the translational start site) were PCR-amplified

from the pNUT173 template, and SOEing PCR was used to combine this amplicon with

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the cqsA promoter. The cloning vector was generated by PCR-amplifying the backbone

of pZND116, and this amplicon and the insert were restriction digested with EcoRI-HF

and KpnI-HF and ligated together. pZND131 was constructed by PCR-amplifying the

PcqsA-gfp region of pZND130 and subcloning this site onto the pZE12 backbone. The

pZE12 backbone was PCR-amplified, and the insert and vector were restriction digested

with EcoRI-HF and KpnI-HF and ligated together. All plasmids were transformed into E.

coli S17-1λpir, and plasmids were mated to V. cholerae strains as described (Skorupski

and Taylor, 1996).

Genomic library screen in E. coli

The V. cholerae genomic library was constructed by partially digesting V.

cholerae genomic DNA (gDNA) with Sau3AI and ligating the fragments into the pZA31

plasmid digested with BamHI. 5µg V. cholerae gDNA was restriction digested for 15

minutes with 0.1X Sau3AI enzyme, and the DNA appeared as a smear when

electrophoresed on 1% agarose, ranging in size from 0.5kb to 2.5kb. The smear was

excised and partitioned by size into 5 sections (500bp-800bp, 800bp-1kb, 1kb-1.5kb,

1.5kb-2kb, 2kb-2.5kb), and each portion of the partial digest was ligated with pZA31

individually. 2x105 total transformants (4x104 from each section) were pooled and

plasmid mini-prepped to generate the plasmid genomic library. The library was

transformed into ZDE507 (electrocompetent E. coli BW-RI harboring plasmid pZND131).

2x105 transformants were pooled and stocked to make the screening library.

An overnight culture of the genomic library was back-diluted 1:1000, and the

library was expressed by activation of the Ptet promoter through addition of 50ng/mL aTc.

At OD600=0.1, the culture was diluted 1:10 into phosphate buffered saline (PBS), and

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FACS-sorted. In the first round of sorting, GFP production from 25 million cells was

measured, and the 1% brightest cells were sorted and retained. The cells were

immediately resorted, and the 4.5x103 cells that remained bright were grown overnight in

LB. This two-stage sorting process was repeated with the enriched culture the following

day. The cells retained after the second enrichment sort were plated, and 103 were

arrayed into 96-well plates the following day and grown overnight. 96-well plate cultures

were back-diluted 1:100, grown to OD600~0.1, and GFP was assessed on an EnVision

96-well plate reader using fluorescence and OD600. Eight strains with above-average

GFP expression were isolated, and the plasmids from these strains were sequenced to

identify the plasmid-based V. cholerae genomic sequence. The basic local alignment

search tool (BLAST) was used to cross-reference the sequences to the V. cholerae

genome to identify relevant genes.

Transposon library screen in V. cholerae

The V. cholerae Tn5 transposon mutant library was constructed by first mating

ΔluxQ V. cholerae with E. coli S17-1λpir harboring the pRL27 plasmid and selecting

on plates containing polymyxin B and kanamycin. 2x105 random insertion mutants

were pooled and mated again with E. coli S17-1λpir harboring the pZND136 plasmid.

The Tn5 library was subsequently plated, 104 mutants were arrayed into 96-well plates,

grown overnight, and back-diluted 1:1000 into new 96-well plates. Cultures were grown

to OD600~0.1 and examined using a 96-well plate reader (EnVision) for GFP

fluorescence and OD600. Cultures showing high GFP production were re-confirmed on

the plate reader and then examined by flow cytometry (BD Biosciences FACSAria). To

map the transposon insertions, gDNA was isolated from the mutant candidates and

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digested with BamHI, self-ligated, and transformed into E. coli TOP10. Transformants

containing the pRL27 backbone fused to V. cholerae genomic DNA were plasmid mini-

prepped, sequenced, and BLASTed against the V. cholerae genome.

Northern Blots

cqsA riboprobes were synthesized by first PCR-amplifying V. cholerae El Tor

C6706str2 gDNA template with primers in Table 4.2, and then performing T7 in vitro

transcription on the PCR amplicon (Ambion) with 32P-α-UTP. qrr4, spot42, and VSsrna24

oligoprobes were synthesized by end-labeling primers in Table 4.2 with 32P-γ-ATP using

T4 PNK (New England Biolabs). Riboprobes and oligoprobes were purified by Illustra

Microspin columns according to the manufacturer’s guidelines (GE Healthcare). For

Northern blot experiments, overnight cultures were back-diluted 1:1000 into fresh LB and

shaken in test tubes at 30°C, grown to OD=1.0. 5 OD units of culture were added to 20%

RNA stop solution (95% ethanol, 5% phenol), mixed by inversion, and frozen in liquid

nitrogen. Samples were stored at -80°C until processing the following day. To isolate

RNA from the samples, samples were thawed at room temperature and subjected to

centrifugation at 5000x g for 10 minutes at 4°C. Supernatants were discarded and pellets

were processed to extract RNA following the Trizol method (Tu and Bassler, 2007). 10ug

of total RNA was resolved on 6% polyacrylamide (PAA, 7M urea) at 300V for 2 hours.

RNA was transferred to Amersham Hybond-XL nylon membranes (Amersham

Biosciences, Piscataway, NJ) for 1 hour at 50V and at 4°C. For hybridization,

membranes were incubated overnight at 70°C for cqsA riboprobes and at 42°C for qrr4

oligoprobes in 15mL Rapid-hyb buffer (GE Healthcare, Piscataway, NJ). Following

overnight incubation, membranes were washed three times in SSC buffer + 0.1%SDS

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(5X, 1X, and 0.5X at 42°C for oligoprobes; 2X, 1X, and 0.5X at 70°C for riboprobes).

Blots were imaged for 20 hours unless otherwise noted on phosphorimager screens and

subsequently scanned using a Typhoon 9410 instrument (GE Healthcare).

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Plasmid name Description Source

pJS1194 Empty vector J. Schaffer, unpublished pRL27c Tn5 transposon, R6Koriγ (Larsen et al., 2002) pSLS4 pEVS143 with qrr4-gfp (Svenningsen et al., 2008) pZA31 Empty vector (Levine et al., 2007) pZE12 Empty vector (Levine et al., 2007) pZND116 pEVS143 with Plac-cqsA-FLAG This study pZND130 pEVS143 with PcqsA-gfp-FLAG This study pZND131 pZE12 with PcqsA-gfp-FLAG This study pZND136 pSLS4 with cmR This study pZND165 PcqsA-cqsA with inverted repeat deletion This study pZND166 PcqsA-cqsA with inverted repeat scramble This study pZND45 pKAS32-ΔPcqsA This study pZND95 pJS1194 with cqsA-FLAG This study pZND132 pZA31 with V. cholerae genomic library This study

Table 5.1 Plasmids used in this study

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Primer name Sequence Use

527 TTT TGA ATT CTT GCG CAG CCC GAC CCG ATT CT pZND131, pZND130 (insert)

529 TTT TGG TAC CGA TCC GGT GAT TGA TTG AGC pZND130 (vector) 530 TTT TGA ATT CCA GAA CCG TTA TGA TGT CGG CGC pZND130 (vector)

533

AAA AGG TAC CAA AAC GAA AGG CCC AGT CTT TCG ACT GAG CCT TTC GTT TTA CTA TTT ATC GTC ATC TTT GTA GTC GAT ATC ATG ATC TTT ATA ATC ACC GTC ATG GTC TTT GTA GTC TTT GTA GAG CTC ATC CAT GCC ATG TGT AAT C

pZND131, pZND130 (insert)

534 ATC TCC TTA ACT AGG GAG TGA TAT ATT GCG TGA TTA AAA ACG TCT GTC pZND130 (insert)

535 CGC AAT ATA TCA CTC CCT AGT TAA GGA GAT ATA CAT ATG AGC AAA GGA pZND130 (insert)

536 TTT TGG TAC CTG CGG CGA GCG GTA T pZND131 (vector)

537 AAA AGA ATT CAC GAA AGG GCC TCG TGA TAC GCC TA pZND131 (vector)

539 CCA GCC CAA TAC GAA TGT TT cqsA riboprobe amplicon

540 GTT TTT TTA ATA CGA CTC ACT ATA GGG AGG CAA TGA TCC CAG GAC CAT GAC G

cqsA riboprobe amplicon

549 AAA AAG CGG CCG CGA AGA TGC GTG ATC TGA TCC TTC AAC TC pZND136 (insert)

550 AAA AAC CTA GGC TTC CTC GCT CAC TGA CTC GCT AC pZND136 (vector)

551 AAA AAG CGG CCG CGT TGG CTT GGT TTC ATC AGC CAT CCG pZND136 (vector)

600 TGA TTT TTC CTC CCC TCA CCA TCG AGA pZND165 (vector)

601 GAA TAT TCT GAT ATA AAA AAT AAT TTA GGA GTT TAC TGA GC

pZND165, pZND166 (vector)

602 ACG ATA TTA ATC GGA AGG AGT ATT CTG ATT TTT CCT CCC CTC ACC ATC GAG A pZND166 (vector)

KPO-0063 CGT CTA TAA GTG TGA ACA ATG GTG qrr4 oligoprobe

Table 5.2 Primers used in this study

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Strain name Organism Genotype Plasmid Source

??? V. cholerae C6706str2 ΔrpoN This study

WN009 E. coli S17-1λpir Wild type

(de Lorenzo and Timmis, 1994)

ZDC757 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND95, pZND122 This study ZDC767 V. cholerae C6706str2 ΔvpsL pZND136 This study ZDC769 V. cholerae C6706str2 ΔvpsL luxO-D47E pZND136 This study ZDC771 V. cholerae C6706str2 ΔvpsL ΔPcqsA pZND136 This study ZDC773 V. cholerae C6706str2 ΔluxQ pZND136 This study ZDC827 V. cholerae C6706str2 ΔvpsL ΔluxO pZND136 This study ZDC851 V. cholerae C6706str2 ΔluxQ ΔcqsA pZND136 This study

ZDC897 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND122, pZND165 This study

ZDC899 V. cholerae C6706str2 ΔvpsL ΔluxO ΔPcqsA pZND122, pZND166 This study

ZDE203 E. coli BW-RI wild type (Levine et al., 2007)

ZDE507 E. coli BW-RI wild type pZND131, pZND132 This study

Table 5.3 Strains used in this study

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