Environmental Regulation of Bioluminescence in Vibrio

ENVIRONMENTAL REGULATION OF BIOLUMINESCENCE

IN VIBRIO FISCHERI ES114

NOREEN LORETTA LYELL

(Under the Direction of Eric V. Stabb)

ABSTRACT

The pheromone-mediated circuitry that governs bioluminescence in Vibrio fischeri is well understood; however, less is known about the environmental conditions that influence pheromone and light production. The environment has been shown to have a profound effect on luminescence in V. fischeri, as cells in symbiotic association with the Hawaiian bobtail squid, Euprymna scolopes, are ~1000-fold brighter than nonsymbiotic cells despite reaching similar cell densities. In this dissertation, I show that luminescence is governed by a complex regulatory web and that certain environmental conditions mediate the regulation of pheromone circuitry. My first goal was to identify and characterize previously unidentified regulators that control luminescence in V. fischeri strain ES114. I helped develop and employed a transposon mutagenesis system to discover novel negative regulators of luminescence. In this study, I characterized twenty-eight independent luminescence-up mutants with insertions in 14 loci. This work revealed that environmental conditions such as inorganic phosphate and Mg2+ concentrations are integrated into the regulation of the pheromone-dependent lux system.

Furthermore, I showed that competition between the LuxI- and AinS-generated pheromones is an important and density-dependent factor in the level of light produced by V. fischeri ES114 cells, such that C8-HSL inhibits luminescence in dense populations.

The second goal of this work was to clarify the role of cAMP receptor protein (CRP) in luminescence. Attempts to study the effects of glucose on V. fischeri luminescence have been contradictory and inconclusive, possibly due to strain-specific effects. I confirmed that both cAMP and glucose modulate light production in ES114, which is consistent with CRP regulation of luminescence. Previous researchers proposed a model wherein

CRP regulates transcription of luxR, which encodes a pheromone-dependent activator known to induce luminescence. The data reported in this dissertation indicates that CRP is involved in the expression of not only luxR but also ainS. Using qRT-PCR and purified CRP, I show that CRP significantly increases transcript levels of both luxR and ainS, and that the CRP protein binds the upstream promoter regions of both genes. Taken together, this dissertation illustrates and clarifies how environmentally responsive regulators are integrated with pheromone-mediated circuitry in the regulation of V. fischeri bioluminescence.

INDEX WORDS: Aliivibrio, Photobacterium, quorum sensing, luminescence, CRP

ENVIRONMENTAL REGULATION OF BIOLUMINESCENCE

IN VIBRIO FISCHERI ES114

NOREEN LORETTA LYELL

B. S., University of Arizona, 2004

A Dissertation Submitted to the Graduate Faculty of The University of Georgia in Partial

Fulfillment of the Requirements for the Degree

DOCTOR OF PHILOSOPHY

ATHENS, GEORGIA

2011

Noreen Loretta Lyell

ENVIRONMENTAL REGULATION OF BIOLUMINESCENCE

IN VIBRIO FISCHERI ES114

NOREEN LORETTA LYELL

Major Professor: Eric V. Stabb Committee: Timothy R. Hoover Mark A. Schell Lawrence J. Shimkets

Electronic Version Approved:

Maureen Grasso Dean of the Graduate School The University of Georgia August 2011

DEDICATION

I dedicate this dissertation to Loretta, you have given me the strength and courage to accomplish my dreams, and to Aubrey, you will always be my sunshine.

ACKNOWLEDGEMENTS

There are several individuals that have aided me throughout graduate school. I especially have to thank my advisor, Eric Stabb. His support and guidance have been essential to my success. Furthermore, he has enabled me to attend meetings and participate in programs that have been instrumental in defining my career goals. I also thank my committee members, Mark Schell, Larry Shimkets, and Tim Hoover. I am a better researcher because of their insight. In addition, I want to acknowledge Anna Karls as her advice and input have advanced me as an instructor and mentor.

TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS ...... v

LIST OF TABLES ...... ix

LIST OF FIGURES ...... x

CHAPTER

1 Introduction and literature review ...... 1

Overview ...... 2

Pheromone-mediated regulation and common bacterial pheromones ...... 3

Environmental control of AI synthesis ...... 5

Catabolite repression and pheromone-mediated regulation ...... 11

Quorum sensing circuitry and the V. fischeri-E.scolopes symbiosis ...... 12

Purpose of this research ...... 19

References ...... 21

2 Effective mutagenesis of Vibrio fischeri using hyperactive mini-Tn5

derivatives ...... 39

Abstract ...... 40

Introduction ...... 41

Results and Discussion ...... 43

Acknowledgements ...... 52

References ...... 53

3 Bright mutants of Vibrio fischeri ES114 reveal conditions and regulators that

control bioluminescence and expression of the lux operon ...... 62

Abstract ...... 63

Introduction ...... 64

Materials and Methods ...... 68

Results ...... 74

Discussion ...... 92

Acknowledgements ...... 98

References ...... 99

4 CRP regulates pheromone-mediated bioluminescence at multiple levels in

Vibrio fischeri ES114 ...... 111

Abstract ...... 112

Introduction ...... 113

Materials and Methods ...... 115

Results ...... 129

Discussion ...... 142

Acknowledgements ...... 149

References ...... 150

5 Conclusions and future directions ...... 158

Elucidating the regulatory web governing the pheromone-dependent lux

system ...... 160

Elucidating the role of CRP in the regulation of luminescence ...... 163

Conclusion ...... 166

vii

References ...... 167

APPENDICES

A Colonization and motility phenotypes of Vibrio fischeri ES114 luminescence-

up mutants ...... 169

B Method for extracting symbiotic Vibrio fischeri cells from the Euprymna

scolopes light organ ...... 180

viii

LIST OF TABLES

Page

Table 2.1: Oligonucleotides designed for this study ...... 46

Table 3.1: Analysis of luminescence-up transposon mutants ...... 75

Table 3.2: Luminescence analysis of mutants involved in the AinS signaling pathways .85

Table 4.1: Bacterial strains, plasmids, and oligonucleotides used in this study ...... 116

Table A.1: Ability of mutant strains to compete with ES114 in co-cultures ...... 175

LIST OF FIGURES

Page

Figure 1.1: Representative AHL molecule ...... 3

Figure 1.2: Schematic of quorum sensing circuitry and lux regulation in V. fischeri...... 14

Figure 1.3: The E. scolopes-V. fischeri symbiosis ...... 17

Figure 1.4: Luminescence induction in V. fischeri is not regulated entirely by population

density...... 18

Figure 2.1: Schematic representation of mini-Tn5-ermR delivery vectors pEVS168 and

pEVS170 ...... 44

Figure 2.2: Southern blotting of mutants, using probe generated from pEVS170 ...... 48

Figure 3.1: Model of autoinducer-mediated regulation of bioluminescence in

V. fischeri……...... 65

Figure 3.2: Genetic complementation of transposon mutants ...... 79

Figure 3.3: Luminescence of topA mutants in SWTO ...... 81

Figure 3.4: Effects of transposon mutations on luxR and luxI promoter-gfp reporters...... 83

Figure 3.5: Effects of Pi concentration on luminescence and lux promoter-reporters ...... 87

Figure 3.6: Effect of Mg2+ concentration on luminescence ...... 88

Figure 3.7: Effect of topA mutations on aeration-dependent regulation of luminescence

and supercoiling ...... 91

Figure 4.1: Effect of glucose on luminescence ...... 129

Figure 4.2: Effect of cAMP on luminescence ...... 131

Figure 4.3: CRP regulation of luxR and ainS promoter activity ...... 133

Figure 4.4: RT-PCR transcript analysis of luxR and ainS ...... 134

Figure 4.5: Fluorescence polarization analysis of CRP binding to the luxR and ainS

promoter regions ...... 136

Figure 4.6: Both ainS and luxR regulation contribute to the effect of cAMP on

luminescence ...... 138

Figure 4.7: Effects of ARI and ARII crp mutations on luxR, ainS, and luxI promoter

activity in E. coli ...... 139

Figure 4.8: Proposed model of CRP-mediated regulation in V. fischeri luminescence ..148

Figure A.1: Effects of transposon mutations on symbiosis ...... 173

Figure B.1: V. fischeri CFU ml-1 at each step of the extraction assay ...... 183

Figure B.2: Microscopic images of V. fischeri-IgG-protein A magnetic bead

complexes ...... 184

CHAPTER 1

INTRODUCTION AND LITERATURE REVIEW

Overview

Bacteria often modulate group behaviors using pheromone-mediated regulatory circuits (55, 67, 119). The purpose of these systems is often framed in terms of a cell- density dependent phenomenon referred to as “quorum sensing” (38, 39). In a typical description of quorum sensing regulation, small signaling molecules called autoinducers accumulate as cell density increases, and upon reaching a critical threshold concentration the autoinducers trigger specific transcriptional activators. This regulatory mechanism enables an entire population of cells to collectively initiate a group behavior by modulating gene expression in response to the local population density.

Although sometimes overlooked, such pheromone signaling systems also provide a regulatory framework for populations of cells to rapidly respond to environmental stimuli provided there is sufficient cell density. Modulation of gene expression in response to environmental conditions is generally thought of in terms of a single cell‟s stress response; however, intercellular pheromone-mediated signaling can allow environmental cues perceived by a subset of cells to be conveyed to a larger population.

This ability to alter group behavior in response to environmental stimuli using population-wide communication may be vital to bacterial adaptation. Such signaling enables bacterial populations to coordinate gene expression in response to both population density and environmental conditions, and to mount a unified multicellular response when it is beneficial to the population. Though regulation of pheromone synthesis in response to environmental cues is well documented, the significance of these observations is less clear largely because few model systems allow observation of such signaling in natural environments relevant to the ecology of the bacterium.

In the remainder of this chapter I briefly describe common types of bacterial pheromone signaling and provide examples where environmentally responsive regulators help control the production or accumulation of pheromones. In particular I will focus on pheromone-mediated control of bioluminescence in Vibrio fischeri. This bacterium‟s natural symbiosis with the Hawaiian bobtail squid, Euprymna scolopes, can be reconstituted and studied under controlled experimental conditions, making it an ideal model system for examining bacterial pheromone-mediated signaling in a natural infection. I will describe the V. fischeri-squid model system and close this chapter by explaining how the goals of my research were to contribute to our understanding of pheromone signaling in V. fischeri.

Pheromone-mediated regulation and common bacterial pheromones

Several bacterial species recognize pheromone molecules generated by their own and/or other species of bacteria. These small diffusible pheromone molecules, referred to as autoinducers, allow cell-cell communication in bacterial populations and typically are autoregulatory, leading to greater expression of their respective synthases.

In Gram-negative bacteria, one O R major group of autoinducer signaling O molecules is the N-acyl homoserine N H n lactones (AHLs). In the AHL product, O an acyl chain is anchored to a FIG. 1.1: Representative AHL molecule. R can be an H or OH group, or there can be a homoserine lactone ring by an amide double-bonded O (ketone) at this position. bond (Fig. 1.1). Depending on the species of origin, AHLs differ by the length of the

acyl group, by modifications at C-3 of the acyl side chain, and by the degree of saturation of the acyl group (35, 57, 126). AHLs are generated by combining lactone moieties from S-adenosyl methionine and N-acyl side chains from charged acyl-acyl carrier proteins in a reaction usually catalyzed by LuxI-type proteins, although structurally distinct AHL synthases such as AinS in V. fischeri have also evolved to perform essentially the same process.

Over fifty species of bacteria have been demonstrated to produce AHLs that are involved in the regulation of processes such as bioluminescence, antibiotic production, conjugation, and biofilm formation (2, 18, 35). Moreover, some organisms are able to respond to AHLs yet do not produce their own, further illustrating the importance of

AHLs in bacterial communities (99). The number of organisms that are known to produce AHLs is sure to increase as new species are discovered, as new genome sequences become available, and as more sensitive methods for recognizing AHLs, particularly novel AHLs, are developed. Many current methods are only able to identify molecules that are known and therefore novel AHLs are not detectable. In addition, the

AHL levels produced by certain bacteria may be below the level of detection by current methods (103).

An additional bacterial signaling molecule utilized by many diverse bacteria is autoinducer-2 (AI-2). The structure(s) of AI-2 is a complex process. Like AHLs, AI-2 synthesis involves S-adenosyl methionine. Specifically, LuxS catalyzes the final reaction in a three step process to generate 4,5-dihydroxy-2,3-pentanedione (DPD), a cyclic molecule that undergoes spontaneous rearrangement to different chemical forms in equilibrium. One of these structures, a furanosyl borate diester, was found to be the AI-2

signal in Vibrio harveyi (12). Interestingly however, a distinct product of DPD was shown to be the molecule perceived as AI-2 by Salmonella enterica serovar

Typhumirium (S. typhumirium) (71). Thus, AI-2 is not a single molecule but rather a population of DPD-derived molecules, with distinct forms being perceived as by different bacteria. Though AHLs are only produced by Gram-negative bacteria and tend to have species-specific structures, homologs of luxS and AI-2 production have been identified in numerous Gram-negative and Gram-positive species (17, 106, 125, 127). This suggests that AHLs provide a means for signaling and gene regulation within a species, whereas

AI-2 may act as a more universal signal that can be monitored by entire communities.

Within AI-2-producing organisms, the AI-2 signal regulates diverse functions such as bioluminescence, virulence, and biofilm formation (54, 56, 127).

For simplicity I have chosen to focus on AHL‟s and AI-2, which are most relevant to my research; however, many other types of bacterial autoinducer pheromones exist and play important regulatory roles. For example, Gram-positive bacteria generate post-translationally modified peptides that mediate cell density dependent regulation.

These signaling molecules are secreted by dedicated ATP-binding-cassette exporters and influence gene expression following recognition by a specific two-component system sensor kinase (55). Like AHL‟s, signaling peptides regulate diverse cellular processes, such as competence and antimicrobial peptide production.

Environmental control of AI synthesis

Bacteria have developed sophisticated mechanisms to recognize and respond to environmental stimuli such as pH, temperature, osmolarity, oxygen concentration, and

nutrient availability, which allow the bacteria to respond and adapt to their environment

(41). Integration of pheromone signaling and environmentally-mediated regulatory networks provides a mechanism for modulating a bacterial population‟s behavior in response to specific stimuli. Furthermore, signals perceived by a subset of the population can be communicated to the larger group. Interestingly, many bacteria employ environmentally responsive regulators to control pheromone-mediated regulatory circuits.

As examples, I will describe regulatory control of pheromone signaling in Agrobacterium tumefaciens, Pseuodomonas aeruginosa, and S. typhimurium. I will also describe a recurring theme of catabolite repression regulation governing pheromone systems.

A. tumefaciens TraI/TraR

A. tumefaciens is the causative agent of plant tumors in hundreds of dicots (30,

98), with tumors resulting from A. tumefaciens-mediated transformation of host cells.

The DNA to be transferred (T-DNA) is maintained in the bacterium on the conjugative tumor-inducing (Ti) plasmid, which carries both the T-DNA and genes that are not transferred to the host cells (13). The vir genes are required for processing and transferring of T-DNA into the host (50, 51), where it is targeted to the host nucleus and becomes integrated into the genomic DNA. Once integrated in the host genome, the T-

DNA directs rapid cell growth (tumor formation) and the synthesis of opines that are utilized as a carbon and nitrogen source by A. tumefaciens (43). The genes required for catabolizing opines are located on the non-transferred portion of the Ti plasmid, along with traI and traR, which encode an autoinducer synthase and corresponding AI- dependent transcriptional activator, respectively.

The pheromone system directed by the traI and traR genes is required for interbacterial transfer of the Ti plasmid (84). In the absence of the TraI synthesized N-3- oxooctanoyl-homoserine lactone (3-oxo-C8-HSL) autoinducer, TraR remains a monomer that is quickly degraded by proteases. When 3-oxo-C8-HSL binds TraR, the structure of

TraR is altered such that it forms dimers and binds its promoter target sequence (128,

129). The TraR-3-oxo-C8-HSL complex targets “tra boxes” upstream of tra and trb operons, which are essential for conjugation (128).

The traR/traI system is regulated in response to the conditions found in a successful infection. The traR gene is in an operon downstream of genes encoding the enzymes necessary for opine catabolism (85), and transcription of this operon is induced by the presence of opines (37). Integration of T-DNA into the plant cell genome results in infected plant cells producing opines, which serve as a nutrient and also signal the bacterial cells that they are within a successfully transformed host. The bacterial cells then begin producing the enzymes necessary for utilization of opines and increase TraR expression. Moreover TraI is regulated by TraR and is therefore also tied to opine availability (49). Opines are only present within an infected host, so A. tumefaciens‟ conjugation apparatus is only produced during infection when there is both high cell density and biochemical feedback, in the form of opines, that the plasmid being transferred has directed the successful engineering of host cells.

P. aeruginosa LasI/LasR-RhlI/RhlR

P. aeruginosa is a highly adaptable opportunistic pathogen capable of infecting plants, animals, and immunocompromised humans. In humans, it is mostly associated

with chronic infections of cystic fibrosis patients. The regulation of virulence factors, biofilm formation, motility, and efflux pump production is controlled by two pheromone- mediated regulatory networks, LasI/R and RhlI/R (100, 121). Up to 11% of the P. aeruginosa genome is regulated either directly or indirectly by these regulatory circuits

(94, 122).

The autoinducer synthase, LasI produces N-3-oxo-dodecanoyl-homoserine lactone (3-oxo-C12-HSL), which binds the transcriptional activator LasR and promotes the production of virulence factors such as proteases (81, 111). In addition, LasR-3-oxo-

C12-HSL upregulates transcription of rhlI, rhlR, and lasI (19, 95). An additional autoinducer synthase, RhlI, produces N-butanoyl-homoserine lactone (C4-HSL), which complexes with the cognate regulator RhlR to activate transcription of genes required for the synthesis of lectins PA-IL and PA-IIL, and pyocyanin (9, 123). Together the two pheromone systems regulate rhamnolipid and hydrogen cyanide production (81, 82).

Although more virulence genes in P. aeruginosa are regulated by the RhlI/R system, certain genes are preferentially regulated by the lasI signal, or both autoinducers in tandem (92, 94).

Environmental conditions predominantly determine whether the Las or Rhl system is active in P. aeruginosa cells (23). In fact, when AHL signals are added exogenously many AHL-dependent genes still have a delay in expression until the stationary phase of growth, apparently because some components of the medium inhibit these systems and must be metabolized before the autoinducer can stimulate their respective regulons (22, 122, 123). This illustrates that factors other than population density impact the Las or Rhl circuitry. Another example of such regulation is the small

RNA-binding protein RsmA, whose regulatory mechanism is antagonistic to the two- component regulatory system GacS/A (83). In P. aeruginosa, RsmA is a negative transcriptional regulator of the autoinducer synthases LasI and RhlI thereby inhibiting production of both autoinducers (47, 83). The environmental signal recognized by

GacS/A is unknown; however, environmental conditions such as nutrient availability, oxygen levels, and growth phase have been shown to influence expression of quorum sensing genes (23, 93, 118).

Though pheromone signaling is typically only described as population dependent gene regulation, as noted above LasI/R and RhlI/R regulation is influences strongly by the environment (23, 118). Interestingly, the role for environmental regulation in quorum sensing-based virulence seems to be most clear when examining the effects of sputum on gene expression. Sputum is a thick mucous that collects within the airways of cystic fibrosis patients and is composed of molecules produced by the host and bacterial colonizers, such as DNA, mucin, phosphatidylcholine, phosphtidylglycerol, albumin, biofilms and yet unidentified small molecules (23, 117). In the presence of sputum extracts, expression of lasI, lasR, rhlI, and rhlR increases significantly and in a dose-dependent manner despite the levels of AHL being below those required for auto- activation of the Las and Rhl systems during growth in the absence of sputum (23).

Thus, virulence genes are highly expressed in response to the host environment, such that

C4-HSL, C12-HSL, and their respective regulons are not simply tied to cell density.

S. typhimurium LuxS

The role of environmental stimuli in the regulation of AI-2 has been extensively studied in the enteric bacterial pathogen S. typhimurium (104-106). In S. typhimurium as

in other bacteria, production of the AI-2 signaling molecule is regulated strongly by environmental cues and growth phase, leading some researchers to classify AI-2 as having a metabolic function rather than a role in quorum sensing (124). AI-2 regulates genes encoding an ABC transport system Lsr, which mediates AI-2 uptake (109). As AI-

2 signal accumulates, the Lsr uptake system is activated reaching peak expression in mid- exponential growth (108). Expression of the regulator InvF is similarly dependent upon

AI-2 activity (14), and is also highest at mid-exponential to stationary phase (63). InvF is a primary regulator in the expression of genes encoding a type III secretion system clustered within S. typhimurium pathogenicity island 1, which is required for invasion into host epithelial cells (40). At comparable cell density outside a host, AI-2 would not necessarily elicit the same behaviors.

Though AI-2 levels increase with cell density, factors associated with a host environment also mediate production of AI-2. In S. typhimurium, AI-2 is only generated when glucose, or to a lesser extent other phosphotransferase sugars, are available as a carbon source (104). The growth- and nutrient-dependent expression of AI-2 in S. typhimurium suggests that this signal is important in communicating that a host has been colonized (104). Furthermore, expression of luxS is responsive to low pH and high osmolarity following growth on glucose, which are two conditions encountered within the host environment (105). The recognition of these signals and production of AI-2 in response enables S. typhimurium populations to collectively modulate gene expression facilitating their existence inside a host.

Catabolite repression and pheromone-mediated regulation

Pheromone signaling is often influenced by carbon source and catabolite repression systems, including regulation by cAMP receptor protein (CRP) or its homologs. In E. coli CRP is activated by cAMP, which is responsive to glucose availability via the phosphoenolpyruvate-dependent carbohydrate phosphotransferase system (68, 86). When glucose is limited, cAMP levels are elevated and this signaling molecule binds CRP. Together, CRP-cAMP bind DNA to activate transcription by recruiting RNAP to promoters, and in E. coli this occurs at more than one hundred genes

(10). Though CRP is predominantly involved in the regulation of genes relevant to carbon metabolism, in several organisms it also regulates the expression of genes involved in pheromone-mediated circuitry.

In the bioluminescent bacterium, V. harveyi, a CRP mutant is ~1000-fold dimmer than the parent strain, suggesting it is an important activator of luminescence. This effect is due to CRP-cAMP directly binding to the promoter of the pheromone-dependent master regulator gene luxR (unrelated to V. fischeri luxR) and the promoter for luxCDABEGH (11). The Vibrio cholerae quorum-sensing master regulator encoded by hapR is also regulated by CRP. Mutants defective in CRP express lower levels of hapR resulting in increased production of cholera toxin and enhanced biofilm (62, 97). Though

CRP influences transcription of hapR, the promoter does not contain a binding site (97).

Similarly, CRP regulates the pheromone synthase luxS in E. coli (20), and CRP activates transcription of the AHL-dependent regulators lasR and expR in P. aeruginosa and

Erwinia chrysanthemi, respectively, by CRP (1, 87). Furthermore, the Agr quorum

sensing circuitry in Staphylococcus aureus that regulates virulence and antibiotic resistance is modulated by carbon-catabolite protein A (CcpA) (96).

These studies suggest that a conserved relationship exists between carbon metabolism and pheromone-mediated regulatory circuitry. The integration of these regulatory networks may have a role in ensuring quorum sensing systems are only activated in the appropriate environmental conditions.

Quorum sensing circuitry and the V. fischeri-E. scolopes symbiosis

Quorum sensing was originally described as controlling bioluminescence in certain marine bacteria including V. fischeri, which has since become a model system for studying bioluminescence, quorum sensing, and bacteria-animal interactions. In particular, V. fischeri strain ES114 is ideal for studying environmental context-dependent regulation of pheromone signaling in a natural symbiotic infection. ES114 was isolated from the light-emitting organ of a Hawaiian bobtail squid, E. scolopes, and it has become a popular research model because this symbiosis (described below) can be reconstituted experimentally. Today V. fischeri strain ES114 is widely utilized as a model largely due to researchers‟ ability to track the onset of quorum sensing mediated luminescence and host colonization within a controlled laboratory setting. For this reason many genetic tools were developed in ES114, and it was the first V. fischeri strain to have its genome sequenced.

Autoinducers and lux expression in V. fischeri

The quorum sensing system in V. fischeri has been studied extensively, and the genetics and biochemistry of light production are well defined. In all V. fischeri that have been examined, the regulatory genes, luxI and luxR, are arranged in divergently transcribed operons with the genes required for light production, luxCDABEG, cotranscribed with luxI (Fig. 1.2) (28, 29, 69). Together the LuxI/LuxR proteins regulate expression of the bioluminescence genes via pheromone signaling. LuxI, an autoinducer synthase, catalyzes the reaction that generates the membrane-permeable autoinducer N-3- oxohexananoyl-L-homoserine lactone (3-oxo-C6-HSL) (26, 53). In addition to 3-oxo-

C6-HSL, V. fischeri generates an additional AHL autoinducer, N-octanoyl-L-homoserine lactone (C8-HSL) which is synthesized by AinS (58).

A third autoinducer produced by V. fischeri is AI-2 generated by LuxS (64, 91).

It is thought that C8-HSL and AI-2 function through distinct receptors, AinR and LuxPQ, respectively to increase luxR expression via a complex regulatory cascade (Fig. 1.2) (31,

65, 72). In V. harveyi, the homologs of AinR and LuxPQ have both kinase and phosphatase activity, and they act on LuxU to control its phosphorylation state. Upon binding of their respective cognate autoinducers, the kinase (but not phosphatase) activity of the autoinducer receptors decreases, leading to relatively less phosphorylated LuxU

(110). The relative amount of LuxU-P/LuxU in turn is directly connected to the phosphorylation state of LuxO, and LuxO-P is a negative regulator of luminescence (33,

64). Phosphorylated LuxO represses luminescence by acting with σ54 to promote expression of a small RNA (sRNA) qrr, which associates with the Hfq chaperone protein to destabilize litR mRNA (61, 73, 74). LitR is a transcriptional activator of luminescence

that binds within the promoter region of luxR (31). In addition to binding AinR, C8-HSL can bind LuxR directly to promote expression of the luxICDABEG operon (Fig. 1.1) (65).

Although C8-HSL is a weaker activator than 3-oxo-C6-HSL (90), at low cell densities V. fischeri ES114 generates more C8-HSL and this is the primary activator of luminescence in culture (65, 102).

AI-2 - LuxP LuxQ 54 Hfq

LuxO-P qrr

C8 - AinR LuxU LuxO

C8 - LuxR (C8 and AI-2 lead to less phosphorylated LuxO) litR 3OC6 - LuxR LitR

luxR luxI luxC luxD luxA luxB luxE luxG “lux box”

FIG. 1.2: Schematic of quorum sensing circuitry and lux regulation in V. fischeri. The three autoinducers, N-3-oxohexanoyl-HSL, N-octanoyl-HSL, and a furanosyl borate diester, are abbreviated as 3OC6, C8, and AI-2, respectively.

At low concentrations of 3-oxo-C6-HSL or C8-HSL, LuxR is not associated with autoinducer and the N terminus of this regulatory protein prohibits the helix-turn-helix motif within the C terminus from binding DNA (15, 44). As 3-oxo-C6-HSL or C8-HSL accumulate in the environment, due to high cell density and/or upregulation of luxI or ainS, and upon reaching a threshold concentration of autoinducer(s), HSL binds the N terminus of LuxR (44). The LuxR-3-oxo-C6-HSL complex is then able to associate with the „lux box‟ within the luxICDABEG promoter region (15, 16). LuxR-3-oxo-C6-HSL bound to the „lux box‟ activates transcription of the luxICDABEG operon (28, 29, 114) by

directly interacting with RNA polymerase and compensating for a weak -35 promoter element (27, 32, 112).

Induction of the luxICDABEG operon leads to bioluminescence through expression of the genes required to generate luciferase and the proteins involved in regenerating necessary substrates. Light is produced when luciferase, composed of LuxA and LuxB, binds a reduced riboflavin phosphate (FMNH2), oxygen, and an aliphatic aldehyde, then converts these substrates to an oxidized flavin (FMN), water, and an aliphatic acid (45, 113). In this reaction FMNH2 binds luciferase and reacts with O2 to generate a 4a-peroxyflavin. A stable intermediate is formed when this enzyme complex interacts with an aliphatic aldehyde and the slow decay of this association results in emission of blue-green light and oxidation of the FMNH2 and aldehyde substrates (70,

113). LuxC, LuxD, LuxE and LuxG (re)generate luciferase‟s aldehyde and FMNH2 substrates (8, 75).

Because of regulation by LuxI and LuxR, the luxICDABEG operon is expressed only when the population has reached the density needed for accumulation of autoinducer(s) (36). Activation of luxICDABEG above basal levels leads to an increase in LuxI and thereby an increase in the concentration of 3-oxo-C6-HSL. Increased concentrations of this autoinducer further activate LuxR and the result is a positive feedback loop that results in a highly luminescent population of V. fischeri (21, 28). The ability to generate light is required for a successful symbiosis between V. fischeri and its squid host, E. scolopes (6, 115). Specifically non-luminescent mutants do not persist as well as wild type within E. scolopes shortly following colonization (6, 115) despite out- competing luminescent populations in some culture conditions (6).

The E. scolopes-V. fischeri symbiosis

E. scolopes has evolved a specialized organ that becomes colonized by V. fischeri shortly after hatching and emits bacterial bioluminescence throughout the life of the animal (Fig. 1.3A) (67). Juvenile E. scolopes hatch lacking V. fischeri symbionts and must acquire the bacteria horizontally from their environment. An adult E. scolopes light organ contains 107-109 V. fischeri cells, ~90% of which are exuded as part of a diel rhythm (4, 60). The daily expulsion of V. fischeri into the environment promotes populations that are up to 30-fold higher in coastal waters that the squid are known to occupy than in other habitats (59, 60). This suggests that the squid host seeds the environment with symbionts thereby promoting rapid colonization of newly hatched juvenile animals (79). This also population enhancement illustrates a benefit of the symbiosis for V. fischeri.

Immediately after hatching, initiation of the symbiosis between V. fischeri and E. scolopes begins (88). Ciliated appendages on either side of the squid light organ collect

V. fischeri from the seawater by increasing water flow near pores located at the base of these appendages (Fig. 1.3B) (80). In addition to increasing water flow, the ciliated appendages shed mucous that provides a matrix in which the bacterial cells aggregate prior to entering the light organ via the pores (76). Though other bacteria are able to collect within the mucous aggregates, V. fischeri outcompetes these cells and will migrate into the pores as little as two hours post-inoculation (77, 80).

After entering the pores, V. fischeri cells travel through ducts lined with outward beating cilia, into antechambers, and lastly into the crypts (80). Throughout the host‟s life, a persistent population of V. fischeri colonizes these epithelium-lined crypt spaces

within the light organ (Fig. 1.3C) (107). Within the crypts, V. fischeri grows rapidly and fills the extracellular space (78, 89) leading to population densities high enough to induce luminescence approximately eight to twelve hours following hatching (67). Light from

V. fischeri apparently provides a mechanism of camouflage for E. scolopes termed

„counter-illumination‟ (52, 88, 101). In counter-illumination, light is projected downward from the light organ with the effect of hiding the animal‟s silhouette from predators and prey below.

Pore Antechamber A B Duct C Crypt

Ciliated epithelial appendages

FIG. 1.3: The E. scolopes-V. fischeri symbiosis. A. E. scolopes juvenile oriented ventral side up. Light organ (circled) is located ventral to the ink sac within the mantle cavity. B. Illustration depicting one half of the light organ with major components of the structure labeled. C. Electron micrograph showing association between V. fischeri cells and epithelium-lined crypt spaces of the E. scolopes light organ. Panels courtesy of Eric V. Stabb.

For counter-illumination to be successful, the host must maintain a viable and luminescent population of symbionts within the light organ. Accordingly the host provides symbionts with substrates for growth and metabolism. Studies have shown that the matrix within the light organ contains peptides and amino acids (42) that may be utilized by V. fischeri. Mannose, sialomucin, dead cells, and chitin have also been suggested as nutrients provided to V. fischeri cells within the light organ (66, 76, 78). In

this partnership, E. scolopes provides nutrients and a niche to V. fischeri which, in turn, provides its host a means of protection during nocturnal activities.

Environmental regulation of V. fischeri luminescence

Though a quorum is important in V. fischeri light production, cell-density alone does not explain the luminescence generated by a symbiotic population. Despite reaching similar densities, cells within the E. scolopes host are ~1000-fold brighter and produce more 3-oxo-C6-HSL (2) than cells grown in culture (Fig. 1.4). These observations suggest that environmentally-responsive regulators have a critical role in regulating luminescence.

The environmental regulators that influence the pheromone signaling circuitry in

V. fischeri are not entirely understood, particularly in strain FIG. 1.4: Luminescence induction in V. fischeri is not regulated entirely by population density. ES114. Catabolite repression and Despite similar cell densities, V. fischeri cells in the host (left) generate ~2800 photons sec-1 cell-1 cAMP receptor protein (CRP)- and cells in colonies (right) generate ~3 photons sec-1 cell-1. Panels courtesy of Eric V. Stabb. mediated regulation of luxR has been suggested but has predominantly been studied using transgenic lux-containing mutants of E. coli (24, 25). Furthermore, attempts to study the effects of glucose on luminescence in V. fischeri are inconclusive and may be strain specific (3, 34). In strain

MJ-1, glucose represses both luminescence and luciferase production (34), whereas it has been reported that there is no effect in ES114 (3). The role of iron in lux expression also has been suggested but may depend on the strain (3, 46). The redox-responsive

ArcA/ArcB two-component system represses transcription of luxI (5), but the exact environmental conditions responsible for this regulation remain unclear. In contrast, the

GacA response regulator positively regulates luminescence in culture, though this regulation does not appear necessary for luminescence in symbiotic cells (120).

Though genes have been identified that influence ES114 luminescence (48, 116,

120), directed research has not been completed to identify environmentally responsive regulators that control the lux quorum sensing circuitry.

Purpose of this research

Environmental stimuli are important factors in the regulation of pheromone signaling circuitry in several organisms. In fact, population-independent factors modulate not only signal production, but also diffusion and recognition of autoinducer molecules (7). Genes have been identified in V. fischeri that regulate pheromone- mediated bioluminescence (5, 48, 116, 120) and we know that environmental regulation, specifically the host environment, is important in light production because non-symbiotic cells are dimmer than symbiotic cells despite reaching similar population sizes (3).

The initial aim of my research was the identification and characterization of regulators that control luminescence in V. fischeri ES114. I helped develop a transposon mutagenesis system, outlined in Chapter 2, which made large-scale mutant screening and identification feasible. In Chapter 3, I describe how I employed this transposon system to screen for mutants of V. fischeri that were brighter than the wild-type strain in an effort to discover novel negative regulators and leading to a better understanding of environmental factors controlling lux.

An additional goal of this research was to determine the role of CRP and glucose in the regulation of V. fischeri ES114 luminescence. This work, described in Chapter 4, helped determine the effects of both glucose and cAMP-CRP on luminescence in V. fischeri and better mechanistically defined the regulatory role of CRP and its integration into the pheromone-signaling system of V. fischeri.

Finally, in Chapter 5 I will discuss how my research has contributed to our understanding of bioluminescence induction and pheromone signaling in V. fischeri

ES114.

References

1. Albus, A. M., E. C. Pesci, L. J. Runyen-Janecky, S. E. West, and B. H.

Iglewski. 1997. Vfr controls quorum sensing in Pseudomonas aeruginosa. J.

Bacteriol. 179:3928-3935.

2. Bassler, B. L. 1999. How bacteria talk to each other: regulation of gene

expression by quorum sensing. Curr. Opin. Microbiol. 2:582-587.

3. Boettcher, K. J., and E. G. Ruby. 1990. Depressed light emission by symbiotic

Vibrio fischeri of the sepiolid squid Euprymna scolopes. J. Bacteriol. 172:3701-

3706.

4. Boettcher, K. J., E. G. Ruby, and M. J. McFall-Ngai. 1996. Bioluminescence

in the symbiotic squid Euprymna scolopes is controlled by a daily biological

rhythm. J. Comp. Physiol. A. 179:65-73.

5. Bose, J. L., U. Kim, W. Bartkowski, R. P. Gunsalus, A. M. Overley, N. L.

Lyell, K. L. Visick, and E. V. Stabb. 2007. Bioluminescence in Vibrio fischeri is

controlled by the redox-responsive regulator ArcA. Mol. Microbiol. 65:538-553.

6. Bose, J. L., C. S. Rosenberg, and E. V. Stabb. 2008. Effects of luxCDABEG

induction in Vibrio fischeri: enhancement of symbiotic colonization and

conditional attenuation of growth in culture. Arch. Microbiol. 190:169-183.

7. Boyer, M., and F. Wisniewski-Dye. 2009. Cell-cell signalling in bacteria: not

simply a matter of quorum. FEMS Microbiol. Ecol. 70:1-19.

8. Boylan, M., C. Miyamoto, L. Wall, A. Graham, and E. Meighen. 1989. Lux C,

D and E genes of the Vibrio fischeri luminescence operon code for the reductase,

transferase, and synthetase enzymes involved in aldehyde biosynthesis.

Photochem. Photobiol. 49:681-688.

9. Brint, J. M., and D. E. Ohman. 1995. Synthesis of multiple exoproducts in

Pseudomonas aeruginosa is under the control of RhlR-RhlI, another set of

regulators in strain PAO1 with homology to the autoinducer-responsive LuxR-

LuxI family. J. Bacteriol. 177:7155-7163.

10. Busby, S., and R. H. Ebright. 1999. Transcription activation by catabolite

activator protein (CAP). J. Mol. Biol. 293:199-213.

11. Chatterjee, J., C. M. Miyamoto, A. Zouzoulas, B. F. Lang, N. Skouris, and E.

A. Meighen. 2002. MetR and CRP bind to the Vibrio harveyi lux promoters and

regulate luminescence. Mol. Microbiol. 46:101-111.

12. Chen, X., S. Schauder, N. Potier, A. Van Dorsselaer, I. Pelczer, B. L. Bassler,

and F. M. Hughson. 2002. Structural identification of a bacterial quorum-sensing

signal containing boron. Nature 415:545-549.

13. Chilton, M. D., M. H. Drummond, D. J. Merio, D. Sciaky, A. L. Montoya, M.

P. Gordon, and E. W. Nester. 1977. Stable incorporation of plasmid DNA into

higher plant cells: the molecular basis of crown gall tumorigenesis. Cell 11:263-

271.

14. Choi, J., D. Shin, and S. Ryu. 2007. Implication of quorum sensing in

Salmonella enterica serovar typhimurium virulence: the luxS gene is necessary for

expression of genes in pathogenicity island 1. Infect. Immun. 75:4885-4890.

15. Choi, S. H., and E. P. Greenberg. 1991. The C-terminal region of the Vibrio

fischeri LuxR protein contains an inducer-independent lux gene activating

domain. Proc. Natl. Acad. Sci. U. S. A. 88:11115-11119.

16. Choi, S. H., and E. P. Greenberg. 1992. Genetic dissection of DNA binding and

luminescence gene activation by the Vibrio fischeri LuxR protein. J. Bacteriol.

174:4064-4069.

17. De Keersmaecker, S. C., K. Sonck, and J. Vanderleyden. 2006. Let LuxS

speak up in AI-2 signaling. Trends Microbiol. 14:114-119.

18. de Kievit, T. R., and B. H. Iglewski. 2000. Bacterial quorum sensing in

pathogenic relationships. Infect. Immun. 68:4839-4849.

19. de Kievit, T. R., Y. Kakai, J. K. Register, E. C. Pesci, and B. H. Iglewski.

2002. Role of the Pseudomonas aeruginosa las and rhl quorum-sensing systems

in rhlI regulation. FEMS Microbiol. Lett. 212:101-106.

20. De Lay, N., and S. Gottesman. 2009. The Crp-activated small noncoding

regulatory RNA CyaR (RyeE) links nutritional status to group behavior. J.

Bacteriol. 191:461-476.

21. Devine, J. H., G. S. Shadel, and T. O. Baldwin. 1989. Identification of the

operator of the lux regulon from the Vibrio fischeri strain ATCC7744. Proc. Natl.

Acad. Sci. U. S. A. 86:5688-5692.

22. Diggle, S. P., K. Winzer, A. Lazdunski, P. Williams, and M. Camara. 2002.

Advancing the quorum in Pseudomonas aeruginosa: MvaT and the regulation of

N-acylhomoserine lactone production and virulence gene expression. J. Bacteriol.

184:2576-2586.

23. Duan, K., and M. G. Surette. 2007. Environmental regulation of Pseudomonas

aeruginosa PAO1 Las and Rhl quorum-sensing systems. J. Bacteriol. 189:4827-

4836.

24. Dunlap, P. V., and E. P. Greenberg. 1985. Control of Vibrio fischeri

luminescence gene expression in Escherichia coli by cyclic AMP and cyclic AMP

receptor protein. J. Bacteriol. 164:45-50.

25. Dunlap, P. V., and E. P. Greenberg. 1988. Control of Vibrio fischeri lux gene

transcription by a cyclic AMP receptor protein-luxR protein regulatory circuit. J.

Bacteriol. 170:4040-4046.

26. Eberhard, A., A. L. Burlingame, C. Eberhard, G. L. Kenyon, K. H. Nealson,

and N. J. Oppenheimer. 1981. Structural identification of autoinducer of

Photobacterium fischeri luciferase. Biochemistry 20:2444-2449.

27. Egland, K. A., and E. P. Greenberg. 1999. Quorum sensing in Vibrio fischeri:

elements of the luxl promoter. Mol. Microbiol. 31:1197-1204.

28. Engebrecht, J., K. Nealson, and M. Silverman. 1983. Bacterial

bioluminescence: isolation and genetic analysis of functions from Vibrio fischeri.

Cell 32:773-781.

29. Engebrecht, J., and M. Silverman. 1984. Identification of genes and gene

products necessary for bacterial bioluminescence. Proc. Natl. Acad. Sci. U. S. A.

81:4154-4158.

30. Escobar, M. A., and A. M. Dandekar. 2003. Agrobacterium tumefaciens as an

agent of disease. Trends Plant. Sci. 8:380-386.

31. Fidopiastis, P. M., C. M. Miyamoto, M. G. Jobling, E. A. Meighen, and E. G.

Ruby. 2002. LitR, a new transcriptional activator in Vibrio fischeri, regulates

luminescence and symbiotic light organ colonization. Mol. Microbiol. 45:131-

143.

32. Finney, A. H., R. J. Blick, K. Murakami, A. Ishihama, and A. M. Stevens.

2002. Role of the C-terminal domain of the alpha subunit of RNA polymerase in

LuxR-dependent transcriptional activation of the lux operon during quorum

sensing. J. Bacteriol. 184:4520-4528.

33. Freeman, J. A., and B. L. Bassler. 1999. Sequence and function of LuxU: a two-

component phosphorelay protein that regulates quorum sensing in Vibrio harveyi.

J. Bacteriol. 181:899-906.

34. Friedrich, W. F., and E. P. Greenberg. 1983. Glucose repression of

luminescence and luciferase in Vibrio fischeri. Arch. Microbiol. 134:87-91.

35. Fuqua, C., M. R. Parsek, and E. P. Greenberg. 2001. Regulation of gene

expression by cell-to-cell communication: acyl-homoserine lactone quorum

sensing. Annu. Rev. Genet. 35:439-468.

36. Fuqua, C., and S. C. Winans. 1996. Conserved cis-acting promoter elements are

required for density-dependent transcription of Agrobacterium tumefaciens

conjugal transfer genes. J. Bacteriol. 178:435-440.

37. Fuqua, C., and S. C. Winans. 1996. Localization of OccR-activated and TraR-

activated promoters that express two ABC-type permeases and the traR gene of Ti

plasmid pTiR10. Mol. Microbiol. 20:1199-1210.

38. Fuqua, C., S. C. Winans, and E. P. Greenberg. 1996. Census and consensus in

bacterial ecosystems: the LuxR-LuxI family of quorum-sensing transcriptional

regulators. Annu. Rev. Microbiol. 50:727-751.

39. Fuqua, W. C., S. C. Winans, and E. P. Greenberg. 1994. Quorum sensing in

bacteria: the LuxR-LuxI family of cell density-responsive transcriptional

regulators. J. Bacteriol. 176:269-275.

40. Galan, J. E., and A. Collmer. 1999. Type III secretion machines: bacterial

devices for protein delivery into host cells. Science 284:1322-1328.

41. Galperin, M. Y. 2006. Structural classification of bacterial response regulators:

diversity of output domains and domain combinations. J. Bacteriol. 188:4169-

4182.

42. Graf, J., and E. G. Ruby. 1998. Host-derived amino acids support the

proliferation of symbiotic bacteria. Proc. Natl. Acad. Sci. U. S. A. 95:1818-1822.

43. Guyon, P., A. Petit, J. Tempe, and Y. Dessaux. 1993. Transformed plants

producing opines specifically promote growth of opine-degrading Agrobacteria.

Mol. Plant-Microbe Interact. 6:92-98.

44. Hanzelka, B. L., and E. P. Greenberg. 1995. Evidence that the N-terminal

region of the Vibrio fischeri LuxR protein constitutes an autoinducer-binding

domain. J. Bacteriol. 177:815-817.

45. Hastings, J. W., and K. H. Nealson. 1977. Bacterial bioluminescence. Annu.

Rev. Microbiol. 31:549-595.

46. Haygood, M. G., and K. H. Nealson. 1985. Mechanisms of iron regulation of

luminescence in Vibrio fischeri. J. Bacteriol. 162:209-216.

47. Heurlier, K., F. Williams, S. Heeb, C. Dormond, G. Pessi, D. Singer, M.

Camara, P. Williams, and D. Haas. 2004. Positive control of swarming,

rhamnolipid synthesis, and lipase production by the posttranscriptional

RsmA/RsmZ system in Pseudomonas aeruginosa PAO1. J. Bacteriol. 186:2936-

2945.

48. Hussa, E. A., T. M. O'Shea, C. L. Darnell, E. G. Ruby, and K. L. Visick.

2007. Two-component response regulators of Vibrio fischeri: identification,

mutagenesis, and characterization. J. Bacteriol. 189:5825-5838.

49. Hwang, I., P. L. Li, L. Zhang, K. R. Piper, D. M. Cook, M. E. Tate, and S. K.

Farrand. 1994. TraI, a LuxI homologue, is responsible for production of

conjugation factor, the Ti plasmid N-acylhomoserine lactone autoinducer. Proc.

Natl. Acad. Sci. U. S. A. 91:4639-4643.

50. Jin, S. G., R. K. Prusti, T. Roitsch, R. G. Ankenbauer, and E. W. Nester.

1990. Phosphorylation of the VirG protein of Agrobacterium tumefaciens by the

autophosphorylated VirA protein: essential role in biological activity of VirG. J.

Bacteriol. 172:4945-4950.

51. Jin, S. G., T. Roitsch, P. J. Christie, and E. W. Nester. 1990. The regulatory

VirG protein specifically binds to a cis-acting regulatory sequence involved in

transcriptional activation of Agrobacterium tumefaciens virulence genes. J.

Bacteriol. 172:531-537.

52. Jones, B. W., and M. K. Nishiguchi. 2004. Counterillumination in the Hawaiian

bobtail squid, Euprymna scolopes Berry (Mollusca: Cephalopoda). Mar. Biol.

144:1151-1155.

53. Kaplan, H. B., and E. P. Greenberg. 1985. Diffusion of autoinducer is involved

in regulation of the Vibrio fischeri luminescence system. J. Bacteriol. 163:1210-

1214.

54. Kendall, M. M., and V. Sperandio. 2007. Quorum sensing by enteric pathogens.

Curr. Opin. Gastroenterol. 23:10-15.

55. Kleerebezem, M., L. E. Quadri, O. P. Kuipers, and W. M. de Vos. 1997.

Quorum sensing by peptide pheromones and two-component signal-transduction

systems in Gram-positive bacteria. Mol. Microbiol. 24:895-904.

56. Kong, K. F., C. Vuong, and M. Otto. 2006. Staphylococcus quorum sensing in

biofilm formation and infection. Int. J. Med. Microbiol. 296:133-139.

57. Kumari, A., P. Pasini, and S. Daunert. 2008. Detection of bacterial quorum

sensing N-acyl homoserine lactones in clinical samples. Anal. Bioanal. Chem.

391:1619-1627.

58. Kuo, A., N. V. Blough, and P. V. Dunlap. 1994. Multiple N-acyl-L-homoserine

lactone autoinducers of luminescence in the marine symbiotic bacterium Vibrio

fischeri. J. Bacteriol. 176:7558-7565.

59. Lee, K. H., and E. G. Ruby. 1992. Detection of the light organ symbiont, Vibrio

fischeri, in Hawaiian seawater by using lux gene probes. Appl. Environ.

Microbiol. 58:942-947.

60. Lee, K. H., and E. G. Ruby. 1994. Effect of the squid host on the abundance and

distribution of symbiotic Vibrio fischeri in nature. Appl. Environ. Microbiol.

60:1565-1571.

61. Lenz, D. H., K. C. Mok, B. N. Lilley, R. V. Kulkarni, N. S. Wingreen, and B.

L. Bassler. 2004. The small RNA chaperone Hfq and multiple small RNAs

control quorum sensing in Vibrio harveyi and Vibrio cholerae. Cell 118:69-82.

62. Liang, W., A. Pascual-Montano, A. J. Silva, and J. A. Benitez. 2007. The

cyclic AMP receptor protein modulates quorum sensing, motility and multiple

genes that affect intestinal colonization in Vibrio cholerae. Microbiol. 153:2964-

2975.

63. Lundberg, U., U. Vinatzer, D. Berdnik, A. von Gabain, and M. Baccarini.

1999. Growth phase-regulated induction of Salmonella-induced macrophage

apoptosis correlates with transient expression of SPI-1 genes. J. Bacteriol.

181:3433-3437.

64. Lupp, C., and E. G. Ruby. 2004. Vibrio fischeri LuxS and AinS: comparative

study of two signal synthases. J. Bacteriol. 186:3873-3881.

65. Lupp, C., M. Urbanowski, E. P. Greenberg, and E. G. Ruby. 2003. The Vibrio

fischeri quorum-sensing systems ain and lux sequentially induce luminescence

gene expression and are important for persistence in the squid host. Mol.

Microbiol. 50:319-331.

66. McFall-Ngai, M., C. Brennan, V. Weis, and L. Lamarcq. 1998. Mannose

adhesin-glycan interactions in the Euprymna scolopes-Vibrio fischeri symbiosis,

p. 273-276, New developments in marine biotechnology. Plenum Press, New

York, NY.

67. McFall-Ngai, M. J., and E. G. Ruby. 1991. Symbiont recognition and

subsequent morphogenesis as early events in an animal-bacterial mutualism.

Science 254:1491-1494.

68. Meadow, N. D., D. K. Fox, and S. Roseman. 1990. The bacterial

phosphoenolpyruvate: glycose phosphotransferase system. Annu. Rev. Biochem.

59:497-542.

69. Meighen, E. A. 1994. Genetics of bacterial bioluminescence. Annu. Rev. Genet.

28:117-139.

70. Meighen, E. A. 1991. Molecular biology of bacterial bioluminescence.

Microbiol. Rev. 55:123-142.

71. Miller, S. T., K. B. Xavier, S. R. Campagna, M. E. Taga, M. F. Semmelhack,

B. L. Bassler, and F. M. Hughson. 2004. Salmonella typhimurium recognizes a

chemically distinct form of the bacterial quorum-sensing signal AI-2. Mol. Cell

15:677-687.

72. Miyamoto, C. M., Y. H. Lin, and E. A. Meighen. 2000. Control of

bioluminescence in Vibrio fischeri by the LuxO signal response regulator. Mol.

Microbiol. 36:594-607.

73. Miyashiro, T., M. S. Wollenberg, X. Cao, D. Oehlert, and E. G. Ruby. 2010.

A single qrr gene is necessary and sufficient for LuxO-mediated regulation in

Vibrio fischeri. Mol. Microbiol. 77:1556-1567.

74. Moller, T., T. Franch, P. Hojrup, D. R. Keene, H. P. Bachinger, R. G.

Brennan, and P. Valentin-Hansen. 2002. Hfq: a bacterial Sm-like protein that

mediates RNA-RNA interaction. Mol. Cell 9:23-30.

75. Nijvipakul, S., J. Wongratana, C. Suadee, B. Entsch, D. P. Ballou, and P.

Chaiyen. 2008. LuxG is a functioning flavin reductase for bacterial

luminescence. J. Bacteriol. 190:1531-1538.

76. Nyholm, S. V., B. Deplancke, H. R. Gaskins, M. A. Apicella, and M. J.

McFall-Ngai. 2002. Roles of Vibrio fischeri and nonsymbiotic bacteria in the

dynamics of mucus secretion during symbiont colonization of the Euprymna

scolopes light organ. Appl. Environ. Microbiol. 68:5113-5122.

77. Nyholm, S. V., and M. J. McFall-Ngai. 2003. Dominance of Vibrio fischeri in

secreted mucus outside the light organ of Euprymna scolopes: the first site of

symbiont specificity. Appl. Environ. Microbiol. 69:3932-3937.

78. Nyholm, S. V., and M. J. McFall-Ngai. 1998. Sampling the light-organ

microenvironment of Euprymna scolopes: description of a population of host cells

in association with the bacterial symbiont Vibrio fischeri. Biol. Bull. 195:89-97.

79. Nyholm, S. V., and M. J. McFall-Ngai. 2004. The winnowing: establishing the

squid-Vibrio symbiosis. Nat. Rev. Microbiol. 2:632-642.

80. Nyholm, S. V., E. V. Stabb, E. G. Ruby, and M. J. McFall-Ngai. 2000.

Establishment of an animal-bacterial association: recruiting symbiotic Vibrios

from the environment. Proc. Natl. Acad. Sci. U. S. A. 97:10231-10235.

81. Pearson, J. P., E. C. Pesci, and B. H. Iglewski. 1997. Roles of Pseudomonas

aeruginosa las and rhl quorum-sensing systems in control of elastase and

rhamnolipid biosynthesis genes. J. Bacteriol. 179:5756-5767.

82. Pessi, G., and D. Haas. 2000. Transcriptional control of the hydrogen cyanide

biosynthetic genes hcnABC by the anaerobic regulator ANR and the quorum-

sensing regulators LasR and RhlR in Pseudomonas aeruginosa. J. Bacteriol.

182:6940-6949.

83. Pessi, G., F. Williams, Z. Hindle, K. Heurlier, M. T. Holden, M. Camara, D.

Haas, and P. Williams. 2001. The global posttranscriptional regulator RsmA

modulates production of virulence determinants and N-acylhomoserine lactones in

Pseudomonas aeruginosa. J. Bacteriol. 183:6676-6683.

84. Piper, K. R., S. Beck von Bodman, and S. K. Farrand. 1993. Conjugation

factor of Agrobacterium tumefaciens regulates Ti plasmid transfer by

autoinduction. Nature 362:448-450.

85. Piper, K. R., S. Beck Von Bodman, I. Hwang, and S. K. Farrand. 1999.

Hierarchical gene regulatory systems arising from fortuitous gene associations:

controlling quorum sensing by the opine regulon in Agrobacterium. Mol.

Microbiol. 32:1077-1089.

86. Postma, P. W., J. W. Lengeler, and G. R. Jacobson. 1993.

Phosphoenolpyruvate:carbohydrate phosphotransferase systems of bacteria.

Microbiol. Rev. 57:543-594.

87. Reverchon, S., M. L. Bouillant, G. Salmond, and W. Nasser. 1998. Integration

of the quorum-sensing system in the regulatory networks controlling virulence

factor synthesis in Erwinia chrysanthemi. Mol. Microbiol. 29:1407-1418.

88. Ruby, E. G. 1996. Lessons from a cooperative, bacterial-animal association: the

Vibrio fischeri-Euprymna scolopes light organ symbiosis. Annu. Rev. Microbiol.

50:591-624.

89. Ruby, E. G., and L. M. Asato. 1993. Growth and flagellation of Vibrio fischeri

during initiation of the sepiolid squid light organ symbiosis. Arch. Microbiol.

159:160-167.

90. Schaefer, A. L., B. L. Hanzelka, A. Eberhard, and E. P. Greenberg. 1996.

Quorum sensing in Vibrio fischeri: probing autoinducer-LuxR interactions with

autoinducer analogs. J. Bacteriol. 178:2897-2901.

91. Schauder, S., K. Shokat, M. G. Surette, and B. L. Bassler. 2001. The LuxS

family of bacterial autoinducers: biosynthesis of a novel quorum-sensing signal

molecule. Mol. Microbiol. 41:463-476.

92. Schuster, M., and E. P. Greenberg. 2006. A network of networks: quorum-

sensing gene regulation in Pseudomonas aeruginosa. Int. J. Med. Microbiol.

296:73-81.

93. Schuster, M., A. C. Hawkins, C. S. Harwood, and E. P. Greenberg. 2004. The

Pseudomonas aeruginosa RpoS regulon and its relationship to quorum sensing.

Mol. Microbiol. 51:973-985.

94. Schuster, M., C. P. Lostroh, T. Ogi, and E. P. Greenberg. 2003. Identification,

timing, and signal specificity of Pseudomonas aeruginosa quorum-controlled

genes: a transcriptome analysis. J. Bacteriol. 185:2066-2079.

95. Seed, P. C., L. Passador, and B. H. Iglewski. 1995. Activation of the

Pseudomonas aeruginosa lasI gene by LasR and the Pseudomonas autoinducer

PAI: an autoinduction regulatory hierarchy. J. Bacteriol. 177:654-659.

96. Seidl, K., M. Stucki, M. Ruegg, C. Goerke, C. Wolz, L. Harris, B. Berger-

Bachi, and M. Bischoff. 2006. Staphylococcus aureus CcpA affects virulence

determinant production and antibiotic resistance. Antimicrob. Agents Chemother.

50:1183-1194.

97. Silva, A. J., and J. A. Benitez. 2004. Transcriptional regulation of Vibrio

cholerae hemagglutinin/protease by the cyclic AMP receptor protein and RpoS. J.

Bacteriol. 186:6374-6382.

98. Smith, E. F., and C. O. Townsend. 1907. A Plant-Tumor of Bacterial Origin.

Science 25:671-673.

99. Smith, J. L., P. M. Fratamico, and J. S. Novak. 2004. Quorum sensing: a

primer for food microbiologists. J. Food Prot. 67:1053-1070.

100. Smith, R. S., and B. H. Iglewski. 2003. P. aeruginosa quorum-sensing systems

and virulence. Curr. Opin. Microbiol. 6:56-60.

101. Stabb, E. V., and D. S. Millikan. 2009. Is the Vibrio fischeri-Euprymna scolopes

symbiosis a defensive mutualism?, p. 85-98, Defensive mutualism in microbial

symbiosis. CRC Press, Boca Raton, FL.

102. Stabb, E. V., A. Schaefer, J. L. Bose, and E. G. Ruby. 2008. Quorum signaling

and symbiosis in the marine luminous bacterium Vibrio fischeri, p. 233-250,

Chemical communication among microbes. ASM Press, Washington, D.C.

103. Steindler, L., and V. Venturi. 2007. Detection of quorum-sensing N-acyl

homoserine lactone signal molecules by bacterial biosensors. FEMS Microbiol.

Lett. 266:1-9.

104. Surette, M. G., and B. L. Bassler. 1998. Quorum sensing in Escherichia coli and

Salmonella typhimurium. Proc. Natl. Acad. Sci. U. S. A. 95:7046-7050.

105. Surette, M. G., and B. L. Bassler. 1999. Regulation of autoinducer production in

Salmonella typhimurium. Mol. Microbiol. 31:585-595.

106. Surette, M. G., M. B. Miller, and B. L. Bassler. 1999. Quorum sensing in

Escherichia coli, Salmonella typhimurium, and Vibrio harveyi: a new family of

genes responsible for autoinducer production. Proc. Natl. Acad. Sci. U. S. A.

96:1639-1644.

107. Sycuro, L. K., E. G. Ruby, and M. McFall-Ngai. 2006. Confocal microscopy of

the light organ crypts in juvenile Euprymna scolopes reveals their morphological

complexity and dynamic function in symbiosis. J. Morphol. 267:555-568.

108. Taga, M. E., S. T. Miller, and B. L. Bassler. 2003. Lsr-mediated transport and

processing of AI-2 in Salmonella typhimurium. Mol. Microbiol. 50:1411-1427.

109. Taga, M. E., J. L. Semmelhack, and B. L. Bassler. 2001. The LuxS-dependent

autoinducer AI-2 controls the expression of an ABC transporter that functions in

AI-2 uptake in Salmonella typhimurium. Mol. Microbiol. 42:777-793.

110. Timmen, M., B. L. Bassler, and K. Jung. 2006. AI-1 influences the kinase

activity but not the phosphatase activity of LuxN of Vibrio harveyi. J. Biol. Chem.

281:24398-24404.

111. Toder, D. S., S. J. Ferrell, J. L. Nezezon, L. Rust, and B. H. Iglewski. 1994.

lasA and lasB genes of Pseudomonas aeruginosa: analysis of transcription and

gene product activity. Infect. Immun. 62:1320-1327.

112. Trott, A. E., and A. M. Stevens. 2001. Amino acid residues in LuxR critical for

its mechanism of transcriptional activation during quorum sensing in Vibrio

fischeri. J. Bacteriol. 183:387-392.

113. Tu, S. C., and H. I. Mager. 1995. Biochemistry of bacterial bioluminescence.

Photochem. Photobiol. 62:615-624.

114. Urbanowski, M. L., C. P. Lostroh, and E. P. Greenberg. 2004. Reversible

acyl-homoserine lactone binding to purified Vibrio fischeri LuxR protein. J.

Bacteriol. 186:631-637.

115. Visick, K. L., J. Foster, J. Doino, M. McFall-Ngai, and E. G. Ruby. 2000.

Vibrio fischeri lux genes play an important role in colonization and development

of the host light organ. J. Bacteriol. 182:4578-4586.

116. Visick, K. L., T. M. O'Shea, A. H. Klein, K. Geszvain, and A. J. Wolfe. 2007.

The sugar phosphotransferase system of Vibrio fischeri inhibits both motility and

bioluminescence. J. Bacteriol. 189:2571-2574.

117. Voynow, J. A., and B. K. Rubin. 2009. Mucins, mucus, and sputum. Chest

135:505-512.

118. Wagner, V. E., D. Bushnell, L. Passador, A. I. Brooks, and B. H. Iglewski.

2003. Microarray analysis of Pseudomonas aeruginosa quorum-sensing regulons:

effects of growth phase and environment. J. Bacteriol. 185:2080-2095.

119. Waters, C. M., and B. L. Bassler. 2005. Quorum sensing: cell-to-cell

communication in bacteria. Annu. Rev. Cell. Dev. Biol. 21:319-346.

120. Whistler, C. A., and E. G. Ruby. 2003. GacA regulates symbiotic colonization

traits of Vibrio fischeri and facilitates a beneficial association with an animal host.

J. Bacteriol. 185:7202-7212.

121. Whiteley, M., and E. P. Greenberg. 2001. Promoter specificity elements in

Pseudomonas aeruginosa quorum-sensing-controlled genes. J. Bacteriol.

183:5529-5534.

122. Whiteley, M., K. M. Lee, and E. P. Greenberg. 1999. Identification of genes

controlled by quorum sensing in Pseudomonas aeruginosa. Proc. Natl. Acad. Sci.

U. S. A. 96:13904-13909.

123. Winzer, K., C. Falconer, N. C. Garber, S. P. Diggle, M. Camara, and P.

Williams. 2000. The Pseudomonas aeruginosa lectins PA-IL and PA-IIL are

controlled by quorum sensing and by RpoS. J. Bacteriol. 182:6401-6411.

124. Winzer, K., K. R. Hardie, and P. Williams. 2002. Bacterial cell-to-cell

communication: sorry, can't talk now - gone to lunch! Curr. Opin. Microbiol.

5:216-222.

125. Winzer, K., K. R. Hardie, and P. Williams. 2003. LuxS and autoinducer-2:

their contribution to quorum sensing and metabolism in bacteria. Adv. Appl.

Microbiol. 53:291-396.

126. Withers, H., S. Swift, and P. Williams. 2001. Quorum sensing as an integral

component of gene regulatory networks in Gram-negative bacteria. Curr. Opin.

Microbiol. 4:186-193.

127. Xavier, K. B., and B. L. Bassler. 2003. LuxS quorum sensing: more than just a

numbers game. Curr. Opin. Microbiol. 6:191-197.

128. Zhu, J., and S. C. Winans. 1999. Autoinducer binding by the quorum-sensing

regulator TraR increases affinity for target promoters in vitro and decreases TraR

turnover rates in whole cells. Proc. Natl. Acad. Sci. U. S. A. 96:4832-4837.

129. Zhu, J., and S. C. Winans. 2001. The quorum-sensing transcriptional regulator

TraR requires its cognate signaling ligand for protein folding, protease resistance,

and dimerization. Proc. Natl. Acad. Sci. U. S. A. 98:1507-1512.

CHAPTER 2

EFFECTIVE MUTAGENESIS OF VIBRIO FISCHERI USING

HYPERACTIVE MINI-TN5 DERIVATIVES1

1Noreen L. Lyell, Anne K. Dunn, Jeffrey L. Bose, Susan L. Vescovi, and Eric V. Stabb.

2008. App. Environ. Microbiol. 74: 7059-7063. Reprinted here with permission of publisher.

Abstract

We have developed a transposon mutagenesis system for Vibrio fischeri ES114 that utilizes a hyperactive mutant Tn5 transposase (E54K and M56A) and optimized transposon ends. Using a conjugation-based procedure, we obtained independent single- insertion mini-Tn5 mutants at a rate of ~10-6. This simple and inexpensive technique represents a significant improvement over previous methods for transposon mutagenesis of V. fischeri and should be applicable to many other bacteria.

Introduction

Vibrio fischeri is a useful model for studies of bioluminescence (18, 35), pheromone-mediated signaling (39, 46), and beneficial animal-bacterium symbioses (32,

36, 47, 49). Other aspects of V. fischeri biology, including its motility (10, 16, 23, 26, 27,

29, 30), metabolism (1, 13, 42), and ability to form biofilms (9, 14, 19, 53, 55, 56), are also active areas of study. Interest in the light-organ symbiosis between V. fischeri and the Hawaiian bobtail squid Euprymna scolopes has led several researchers to adopt strain

ES114 (4), a wild-type isolate from an E. scolopes light organ, as the type strain of choice for many of these studies. The recently published genome sequence of ES114 (33), and the ongoing development of genetic tools for ES114 (10-12, 38, 40, 48) promise to advance each of the research areas described above.

The goal of this study was to develop a more effective method for transposon mutagenesis of V. fischeri ES114. Transposon mutagenesis systems have been used with

V. fischeri, but these have important limitations. Initially, a Mu-based transposon was used in V. fischeri (16), but in this system, the delivery plasmid was maintained at low selective temperature, and multiple insertions were more common than single insertions at higher temperature. Furthermore, insertions by the Mu dI 1681 transposon were not cloned or localized in the V. fischeri genome (16). A mini-Tn10 based system was then developed and used with V. fischeri (48); however, transposition frequency is relatively low, and unwanted integrants of the plasmid delivery vector are common and can only be identified by molecular methods (K. Visick, personal communication). Mini-Tn5 systems have been used in V. fischeri as well (17, 37), but in one case, we found insertion to be impractically infrequent for large-scale screens (37), and that investigation, along

with another study, relied on mutagenizing an already mutant V. fischeri background (17,

37).

The reason a mutant strain was used as the parent for further mutagenesis stems from each of the systems above relying on conjugation from Escherichia coli to deliver the transposon into V. fischeri, so to counterselect against E. coli, donors a spontaneous rifampicin-resistant derivative of ES114, designated ESR1, has been used (16).

Unfortunately, rifampicin resistance often results in pleiotropic phenotypes (2, 3, 20, 54), and recently, ESR1 was found to be attenuated in both competitive host colonization (24) and in production of bioluminescence (5). We have introduced plasmids directly into wild-type ES114 by conjugation, using low temperature and high salt to enrich against E. coli donors (38), but this is effective only if the desired transconjugants occur frequently relative to the number and growth of E. coli donor cells on the selective medium (e.g. more frequently than the mini-Tn10 or mini-Tn5 insertions described above occur).

Thus, in our experience existing transposon mutagenesis systems have limited utility with wild type V. fischeri ES114.

To improve transposon mutagenesis of V. fischeri, we exploited advances in the understanding of Tn5 by combining optimized Tn5 end sequences (58) and a hyperactive transposase allele (59). The wild-type Tn5 transposase gene has an alternate translational start site, giving rise to an Inh protein that inhibits transposase activity, but an M56A change removes this alternate start and prevents Inh translation (51). An additional E54K change yielded a much more active transposase with improved binding to the transposon ends (59). Moreover, the Tn5 transposase normally acts at inside end and outside end sequences of IS50 elements; however, the transposase proved to be more active with a

hybrid of the inside end and outside end sequences, termed a mosaic end (31, 58). These improvements in transposition efficiency formed the foundation of strategies for in vitro mutagenesis of naked DNA with mini-Tn5 derivatives and the generation of mutants in vivo by electroporation of transposase-DNA complexes (31). However, transformation frequencies by electroporation are extremely low in V. fischeri (45), and we therefore sought to generate a conjugation-based system that makes use of these improvements in

Tn5 efficiency. One such system generated and modified by others has been effective in diverse bacteria (22, 41), although as discussed below, we sought to construct a system with somewhat different attributes, particularly with respect to the selectable and screenable markers compatible with V. fischeri.

Results and Discussion

We combined the optimized transposase allele and end elements with an erythromycin resistance gene (ermR) to generate miniTn5 delivery vectors pEVS168 and pEVS170 (Fig. 2.1). Each of these vectors contains the RP4 origin of transfer (oriT), thereby allowing their mobilization from E. coli into V. fischeri (38). The transposon in pEVS168 also has promoterless chloramphenicol resistance (cat) and green fluorescent protein (gfp) genes, such that insertion in the correct orientation in actively transcribed regions can generate transcriptional fusions to these selectable (cat) and readily screened

(gfp) markers.

oriT

pEVS168 tnp*

(6485 bp) kanR

R6K ermR gfp cat M13F

mini-Tn5 (3296 bp)

oriT

pEVS170 tnp*

(5311 bp) kanR

R6K ermR M13F

mini-Tn5 (2122 bp)

Fig. 2.1: Schematic representation of miniTn5-ermR delivery vectors pEVS168 and pEVS170. The tnp* gene encodes a Tn5 transposase with mutations leading to E54K and M56A substitutions (59). Croos-hatched boxes indicate a mosaic transposon end (5‟- CTG TCT CTT ATA CAC ATC T-3‟) that serves as a more frequent substrate for transposition (31, 58). R6K denotes the origin of replication, oriT indicates the RP4 origin of conjugative transfer, and the kanR and ermR genes encode resistance to kanamycin and erythromycin, respectively. Stem loops indicate the location of bidirectional transcriptional terminators, and M13F shows the location of an M13 forward priming site for sequencing. Restriction sites (presented 5‟-3‟) absent from both transposons include BglII (A/GATCT), EcoRI (G/AATTC), HhaI (GCG/C), XbaI (T/CTAGA), and XhoI (C/TCGAG).

We constructed pEVS168 and pEVS170, using standard cloning techniques and employed reagents and procedures described previously (1). Details of pEVS168 and pEVS170 construction were as follows. The R6K origin, ermR, cat, and gfp genes of pEVS127 (see supplemental material from Dunn et al. (12)) were PCR amplified using primers EVS90 and EVS91 (Table 2.1); BglII sites introduced on the primers were

digested with this enzyme, and the product was self-ligated to form pEVS155.

Optimized 19-bp Tn5 mosaic ends were incorporated into primers EVS90 and EVS91, along with restriction sites, such that most of pEVS155 was within a transposable unit with only the unique BglII and SalI sites outside the mini-transposon. Plasmid pEVS129 was generated by deleting the XbaI-SpeI fragment of pEVS118 (11), and pEVS129 was linearized with BglII and fused to BamHI-digested pRZ5412 (59), which encodes a hyperactive mutant transposase. The transposase gene on the resulting plasmid fusion was PCR amplified using primers EVS62 and EVS92 (Table 2.1), this amplicon was digested with BamHI, and the fragment was cloned into the BglII site of pEVS155, generating pEVS157. The inside-end-containing SalI fragment of pEVS157 was deleted and subsequently replaced with the oriT- and kanR-containing SalI fragment of pEVS76

(38), resulting in pEVS167. pEVS168 was then generated by annealing oligonucleotides

M13LKF and M13LKR (Table 2.1) together and cloning this linker into the XbaI site of pEVS167, thereby introducing a unique ApaI site and a complement for the M13 forward sequencing primer into one end of the transposon. pEVS168, which is similar to pEVS170 (Fig. 2.1) but contains a promoterless cat-gfp transcriptional reporter on the transposon distal to the M13 forward site, was digested with BspEI and BsrGI, the overhangs were filled with Klenow fragment, and the vector self-ligated to delete gfp along with most of cat, generating pEVS170. Translational stop codons were incorporated into primer EVS91 (Table 2.1), so that insertions of the transposon from pEVS168 could generate transcriptional, but not translational, fusions of chromosomal genes to cat and gfp.

Table 2.1: Oligonucleotides designed for this study

Primer name Oligonucleotide sequence (5‟-3‟)a

EVS62 CTT CAG ATC CTC TAC GCC GGA CGC

EVS90 GGG AGA TCT GTC GAC CTG TCT CTT ATA CAC ATC TGC

GGC CGC TCT AGA ACT AGT GGA TCC

EVS91 GGG AGA TCT GTC TCT TAT ACA CAT CTA AGG TAA TCA

GGA GCT AAG GAA GCT AAA ATG G

EVS92 CCC GGA TCC GTA GCG TCC TGA ACG GAA CCT TTC CCG

M13LKF CTA GGG GCC CTG TAA AAC GAC GGC CAG TC

M13LKR CTA GGA CTG GCC GTC GTT TTA CAG GGC CC a Optimized 19-bp Tn5 mosaic ends (5‟-CTG TCT CTT ATA CAC ATC T-3‟) are indicated by single underlines in EVS90 and EVS91. The M13 Forward priming site (5‟- TGT AAA ACG ACG GCC AGT-3‟) and its complement formed by annealing M13LKF and M13LKR together are underlined in those sequences. Translational stops (5‟-TAA- 3‟) present in all three potential reading frames in EVS91, are shown in boldface. Key restriction enzyme recognition sequences are in italics, including ApaI (5‟-GGGCC/C-3‟) in M13LKF and M13LKR, BamHI (5‟-G/GATCC-3‟) in EVS92, BglII (5‟-A/GATCT- 3‟) in EVS90 and EVS91, and SalI (5‟-G/TCGAC-3‟) in EVS90.

This transposon system proved highly effective with V. fischeri. We used a simple mating procedure (38) to introduce pEVS170 into V. fischeri ES114 and found that typically, ~104 miniTn5-ermR mutants were selected as erythromycin resistant from

~1010 total recipients in each small-scale mating, yielding a transposon mutagenesis frequency of 10-6. It is worth noting that erythromycin selection, which was carried out on LBS medium (5) supplemented with 5 μg ml-1 erythromycin, resulted in a clearer distinction between Tn mutants and other cells in the mating mix than did the chloramphenicol selection associated with some previous transposons, and this may have

contributed to our success. Erythromycin-resistant transconjugants were recovered about

104-fold more frequently upon introduction of the stable ermR plasmid pS44S (11), and assuming that conjugative transfer frequency is similar for pEVS170 and pS44S, which each have the same oriT, the frequency of transposition once pEVS170 has entered V. fischeri can be estimated as ~10-4. We identified mutants resulting from plasmid integration, rather than miniTn5-ermR transposition, by screening for kanamycin resistance, which we observed in ~10% of the erythromycin-resistant mutants. Similar frequencies were obtained when the mini-transposon contained promoterless cat and gfp genes, and epifluorescence microscopy revealed that about a quarter of these insertion mutants visibly expressed GFP, with different mutants varying in their degree of green fluorescence (data not shown). This result is consistent with non-specific insertions generating transcriptional fusions to the cat-gfp reporter, assuming that (i) only a fraction of the genome will be transcribed significantly during growth on LBS plates, (ii) approximately half of the insertions in transcribed regions will be oriented in the direction of transcription, and (iii) levels of transcription will vary between loci.

Although the recovery of transposon insertion mutants was more efficient than we and others (K. Visick, personal communication) have observed with other Tn5- or Tn10- based systems in V. fischeri, we were concerned that the hyperactive transposase on pEVS170 might yield multiple miniTn5-ermR insertions in mutants. However, Southern blotting of over fifty mutants revealed that this is not the case. Chromosomal DNA was purified from the transposon insertion mutants using Easy-DNA (Invitrogen, Carlsbad,

Calif.), the DNA was restriction digested, and 5 ng of each digested DNA sample was separated by gel electrophoresis. Digested DNA was transferred from the gel onto

nitrocellulose paper and then prepared for imaging using a digoxigenin High Prime DNA labeling and detection starter kit II (Roche Applied Science, Mannheim, Germany).

Southern blot analysis revealed a single Tn insertion in each strain examined (e.g., Fig.

2.2).

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16

5 4 3 2 1

Fig. 2.2: Southern blotting of mutants, using a probe generated from pEVS170. Arrows and corresponding numbers at right indicate DNA fragment sizes in kb. Lanes 1 and 16 were loaded with 1-kb DNA ladder. Lanes 2 to 14 were loaded with HhaI-digested chromosomal DNA from mutant strains NL1, NL2, NL3, NL4, NL5, NL6, NL8, EMH5, EMH6, EMH7, EMH9, EMH11, EMH12, and EMH13, respectively. Fourteen representative mutants of more than 50 examined are shown.

The composition of the miniTn5-ermR mutant from pEVS170 allowed us to clone it readily from insertion mutants and to identify insertion locations in the V. fischeri genome. The R6K plasmid origin of replication within the mini-transposon does not function in V. fischeri, but it does replicate in E. coli strains harboring the pir gene (21), such as DH5-pir (11). Chromosomal DNA from mutants was digested with HhaI, which does not cut within the transposon, the fragments were self-ligated, and transposons along with flanking chromosomal DNA were transformed as circularized plasmids into DH5-pir. The M13 forward sequencing primer was then used to sequence across transposon-chromosome junctions, and we localized the point of

insertion by comparisons to the genome sequence (33). HhaI digests DNA at a 4-bp recognition sequence (5‟-GCG/C-3‟), and virtually all HhaI fragments of the ES114 genome are small enough to be recovered readily by this cloning procedure yet large enough to allow unambiguous identification of the genomic location.

Multiple lines of evidence suggest that our miniTn5-ermR transposons exhibit a lack of target specificity, similar to that of other Tn5 derivatives (15, 34), which generally insert with some deviation from complete randomness but not enough to present a practical barrier to mutagenesis strategies. For example, Figure 2.2 illustrates a lack of target specificity with the miniTn5-ermR inserting into different-sized chromosomal fragments. In addition, we sequenced the location of more than 60 insertions, and we found them distributed on both V. fischeri chromosomes. Moreover, we have never identified an insertion in the same site from more than one independent mating. Finally, we illustrated this transposon mutagenesis system‟s practical utility recently when we used it to identify the cellobiose utilization cel gene cluster in V. fischeri ES114 (1).

Some deviation from random insertion may be intimated in that study by an apparently disproportionate number of insertions in celK, which coincidentally has a lower GC content (31.8%) than other genes of the cluster (35.3 to 39.6%); however, insertions in all six cel genes were recovered without difficulty (1).

Despite the use of ES114 for nearly 2 decades, to our knowledge, the use of pEVS170 (Fig. 2.1) to identify cellobiose-utilization mutants (1) was the first report of transposon mutagenesis in this wild-type strain and not a pleiotropic and marked derivative such as ESR1. Other laboratories are currently using pEVS168, pEVS170, or their derivatives to make new discoveries in V. fischeri. In a particularly exciting

development, Cheryl Whistler (personal communication) has proposed utilizing this system to generate a comprehensive transposon mutant library with insertions in every nonessential gene of the V. fischeri genome.

The tools we describe here are similar to pRL27 (22), a conjugation-based mini-

Tn5 delivery vector that exploits the same Tn5 mosaic end sequences and hyperactive transposase mutations that are similar to those we used in pEVS168 and pEVS170; however, our constructs differ somewhat from that system. For example, the hyperactive transposase in pRL27 contains an additional L372P allele, which apparently minimizes nonproductive multimerization of transposase and improves transposition frequency when the transposase gene is in trans to the transposon ends (50). However, the L372P allele had no effect on transposition frequency in cis (50), and it therefore would not be expected to influence systems like those described here or in pRL27, where the transposase gene is delivered on the same vector as the mini-Tn5. For our purposes, a more important difference is the use of an erythromycin gene cassette within the transposon, rather than the kanamycin resistance gene in pRL27. We have never observed spontaneous resistance to erythromycin in V. fischeri, but we do observe spontaneous resistance to kanamycin at a frequency rivaling that of transposition, making the erythromycin resistance marker superior for primary selection of transposon hops.

The kanamycin resistance gene functions well as a screenable marker in V. fischeri, and by placing it outside the transposon, we distinguished transposon hops from vector integrants. There is no such marker outside the transposon in pRL27. For unknown reasons we observe a large number of integrants (~10% of Tn-harboring clones) in V. fischeri, and the ability to screen for and discard vector integrants is important, as they

may contain the transposase gene and therefore could accumulate multiple mini-Tn5 insertions. Finally, our constructs contain other advantageous features, including promoterless selectable and screenable markers (cat-gfp) in pEVS68, a universal primer site near the transposon end, and a unique ApaI site within each mini-Tn5-ermR construct that can be used to further engineer this system

Combining optimizing elements in an in vivo conjugation-based mini-Tn5 mutagenesis system has opened up new research opportunities in V. fischeri, and we expect our constructs will be useful in other bacteria for which existing transposon mutagenesis strategies are inefficient. The similar pRL27 system devised by Larsen et al.

(22) has proven useful in many bacteria including Xanthobacter autotrophicus (22),

Pseudomonas stutzeri (22), other Pseudomonas strains (28, 57), Alcaligenes faecalis (22,

52), E. coli (6), Bdellovibrio bacteriovorus (44), Prochlorococcus MIT9313 (43), and

Synechococcus WH8102 (25), while modified versions of pRL27 were also successfully employed in V. parahaemolyticus (41), Citrobacter rodentium (8), and Francisella tularensis (7). It therefore seems likely that our constructs will have similarly broad utility and the different features of our system will extend the range of organisms in which conjugation-based mini-Tn5 mutagenesis is a practical tool.

Acknowledgements

We thank Erica M. Hall and Alecia N. Septer for technical assistance, and

William Reznikoff for providing pRZ5412.

This material is based upon work supported by the National Science Foundation

(NSF) under grant CAREER MCB-0347317 to E. V. S., and by National Institutes of

Health (NIH) grant AI50661 to M. McFall-Ngai. S. L. V. was supported by a NSF

Research Experience for Undergraduates site award (DBI-0453353).

Genomic information was provided by the Vibrio fischeri Genome Project supported by the W. M. Keck Foundation.

References:

1. Adin, D. M., K. L. Visick, and E. V. Stabb. 2008. Identification of a cellobiose

utilization gene cluster with cryptic beta-galactosidase activity in Vibrio fischeri.

Appl Environ Microbiol 74:4059-4069.

2. Björkman, J., D. Hughes, and D. I. Andersson. 1998. Virulence of antibiotic-

resistant Salmonella typhimurium. Proceedings of the National Academy of

Sciences USA 31:3949-3953.

3. Blanc-Potard, A., E. Gari, F. Spirito, N. Figueroa-Bossi, and L. Bossi. 1995.

RNA polymerase (rpoB) mutants selected for increased resistance to gyrase

inhibitors in Salmonella typhimurium. Molecular and General Genetics 247:680-

692.

4. Boettcher, K. J., and E. G. Ruby. 1990. Depressed light emission by symbiotic

Vibrio fischeri of the sepiolid squid Euprymna scolopes. Journal of Bacteriology

172:3701-3706.

5. Bose, J. L., C. S. Rosenberg, and E. V. Stabb. 2008. Effects of luxCDABEG

induction in Vibrio fischeri: enhancement of symbiotic colonization and

conditional attenuation of growth in culture. Archives of Microbiology 190:169-

183.

6. Bradshaw, J. S., and A. Kuzminov. 2003. RdgB acts to avoid chromosome

fragmentation in Escherichia coli. Mol Microbiol 48:1711-1725.

7. Buchan, B. W., M. K. McLendon, and B. D. Jones. 2008. Identification of

differentially regulated Francisella tularensis genes by use of a newly developed

Tn5-based transposon delivery system. Appl Environ Microbiol 74:2637-2645.

8. Coulthurst, S. J., S. Clare, T. J. Evans, I. J. Foulds, K. J. Roberts, M. Welch,

G. Dougan, and G. P. Salmond. 2007. Quorum sensing has an unexpected role

in virulence in the model pathogen Citrobacter rodentium. EMBO Rep 8:698-

703.

9. Darnell, C. L., E. A. Hussa, and K. L. Visick. 2008. The putative hybrid sensor

kinase SypF coordinates biofilm formation in Vibrio fischeri by acting upstream

of two response regulators, SypG and VpsR. J Bacteriol 190:4941-4950.

10. DeLoney-Marino, C. R., A. J. Wolfe, and K. L. Visick. 2003. Chemoattraction

of Vibrio fischeri to serine, nucleosides, and N-acetylneuraminic acid, a

component of squid light-organ mucus. Applied and Environmental Microbiology

69:7527-7530.

11. Dunn, A. K., M. O. Martin, and E. V. Stabb. 2005. Characterization of

pES213, a small mobilizable plasmid from Vibrio fischeri. Plasmid 54:114-134.

12. Dunn, A. K., D. S. Millikan, D. M. Adin, J. L. Bose, and E. V. Stabb. 2006.

New rfp- and pES213-derived tools for analyzing symbiotic Vibrio fischeri reveal

patterns of infection and lux expression in situ Applied and Environmental

Microbiology 72:802-810.

13. Dunn, A. K., and E. V. Stabb. 2008. Genetic analysis of trimethylamine N-oxide

reductases in the light organ symbiont Vibrio fischeri ES114. J Bacteriol

190:5814-5823.

14. Geszvain, K., and K. L. Visick. 2008. The hybrid sensor kinase RscS integrates

positive and negative signals to modulate biofilm formation in Vibrio fischeri. J

Bacteriol 190:4437-4446.

15. Goryshin, I. Y., J. A. Miller, Y. V. Kil, V. A. Lanzov, and W. S. Reznikoff.

1998. Tn5/IS50 target recognition. Proc Natl Acad Sci U S A 95:10716-10721.

16. Graf, J., P. V. Dunlap, and E. G. Ruby. 1994. Effect of transposon-induced

motility mutations on colonization of the host light organ by Vibrio fischeri.

Journal of Bacteriology 176:6986-6991.

17. Graf, J., and E. G. Ruby. 2000. Novel effects of a transposon insertion in the

Vibrio fischeri glnD gene: defects in iron uptake and symbiotic persistence in

addition to nitrogen utilization. Molecular Microbiology 37:168-179.

18. Hastings, J. W., and K. H. Nealson. 1977. Bacterial bioluminescence. Annual

Reviews of Microbiology 31:549-595.

19. Hussa, E. A., C. L. Darnell, and K. L. Visick. 2008. RscS functions upstream of

SypG to control the syp locus and biofilm formation in Vibrio fischeri. J Bacteriol

190:4576-4583.

20. Jin, D. J., and C. A. Gross. 1989. Characterization of the pleiotropic phenotypes

of rifampin-resistant rpoB mutants of Escherichia coli. Journal of Bacteriology

171:5229-5231.

21. Kolter, R., M. Inuzuka, and D. R. Helinski. 1978. Trans-complementation-

dependent replication of a low molecular weight origin fragment from plasmid

R6K. Cell 15:1199-1208.

22. Larsen, R. A., M. M. Wilson, A. M. Guss, and W. W. Metcalf. 2002. Genetic

analysis of pigment biosynthesis in Xanthobacter autotrophicus Py2 using a new,

highly efficient transposon mutagenesis system that is functional in a wide variety

of bacteria. Archives of Microbiology 178:193-201.

23. Lupp, C., and E. G. Ruby. 2005. Vibrio fischeri uses two quorum-sensing

systems for the regulation of early and late colonization factors. Journal of

Bacteriology 187:3620-3629.

24. McCann, J., E. V. Stabb, D. S. Millikan, and E. G. Ruby. 2003. Population

dynamics of Vibrio fischeri during infection of Euprymna scolopes. Applied and

Environmental Microbiology 69:5928-5934.

25. McCarren, J., and B. Brahamsha. 2005. Transposon mutagenesis in a marine

Synechococcus strain: isolation of swimming motility mutants. J Bacteriol

187:4457-4462.

26. Millikan, D. S., and E. G. Ruby. 2002. Alterations in Vibrio fischeri motility

correlate with a delay in symbiosis initiation and are associated with additional

symbiotic colonization defects. Applied and Environmental Microbiology

68:2519-2528.

27. Millikan, D. S., and E. G. Ruby. 2004. Vibrio fischeri flagellin A is essential for

normal motility and for symbiotic competence during initial squid light organ

colonization. Journal of Bacteriology 186:4315-4325.

28. Newman, K. L., S. Chatterjee, K. A. Ho, and S. E. Lindow. 2008. Virulence of

plant pathogenic bacteria attenuated by degradation of fatty acid cell-to-cell

signaling factors. Molecular Plant-Microbe Interactions 21:326-334.

29. O'Shea, T. M., C. R. DeLoney-Marino, S. Shibata, S.-I. Aizawa, A. J. Wolfe,

and K. L. Visick. 2005. Magnesium promotes flagellation of Vibrio fischeri.

Journal of Bacteriology 187:2058-2065.

30. O'Shea, T. M., A. H. Klein, K. Geszvain, A. J. Wolfe, and K. L. Visick. 2006.

Diguanylate cyclases control magnesium-dependent motility of Vibrio fischeri.

Journal of Bacteriology 188:8196-8205.

31. Reznikoff, W. S., I. Y. Goryshin, and J. J. Jendrisak. 2004. Tn5 as a molecular

genetics tool. In W. J. Miller and P. Capy (ed.), Mobile genetic elements:

protocols and genomic applications, vol. 260. Human Press Inc., Totowa, N.J.

32. Ruby, E. G. 1996. Lessons from a cooperative bacterial-animal association: The

Vibrio fischeri-Euprymna scolopes light organ symbiosis. Annual Reviews of

Microbiology 50:591-624.

33. Ruby, E. G., M. Urbanowski, J. Campbell, A. Dunn, M. Faini, R. Gunsalus,

P. Lohstroh, C. Lupp, J. McCann, D. Millikan, A. Schaefer, E. Stabb, A.

Stevens, K. Visick, C. Whistler, and E. P. Greenberg. 2005. Complete genome

sequence of Vibrio fischeri: A symbiotic bacterium with pathogenic congeners.

Proceedings of the National Academy of Sciences USA 102:3004-3009.

34. Shevchenko, Y., G. G. Bouffard, Y. S. Butterfield, R. W. Blakesley, J. L.

Hartley, A. C. Young, M. A. Marra, S. J. Jones, J. W. Touchman, and E. D.

Green. 2002. Systematic sequencing of cDNA clones using the transposon Tn5.

Nucleic Acids Res 30:2469-2477.

35. Stabb, E. V. 2005. Shedding light on the bioluminescence "paradox". ASM News

71:223-229.

36. Stabb, E. V. 2006. The Vibrio fischeri-Euprymna scolopes light organ symbiosis,

p. 204-218. In F. L. Thompson, B. Austin, and J. Swings (ed.), The Biology of

Vibrios. ASM Press, Washington D.C.

37. Stabb, E. V., and E. G. Ruby. 1998. Construction of a mobilizable Tn5

derivative and its utility for mutagenizing, cloning, and mapping loci in the

luminescent symbiont Vibrio fischeri, p. 375-376, American Society for

Microbiology, 98th General Meeting. American Society for Microbiology,

Washington, D.C.

38. Stabb, E. V., and E. G. Ruby. 2002. RP4-based plasmids for conjugation

between Escherichia coli and members of the Vibrionaceae. Methods in

Enzymology 358:413-426.

39. Stabb, E. V., A. Schaefer, J. L. Bose, and E. G. Ruby. 2008. Quorum signaling

and symbiosis in the marine luminous bacterium Vibrio fischeri, p. 233-250. In S.

C. Winans and B. L. Bassler (ed.), Chemical Communication Among Microbes.

ASM Press, Washington D.C.

40. Stabb, E. V., K. L. Visick, D. S. Millikan, A. A. Corcoran, L. Gilson, S. V.

Nyholm, M. McFall-Ngai, and E. G. Ruby. 2001. The Vibrio fischeri-Euprymna

scolopes symbiosis: A model marine animal-bacteria interaction, p. 269-277. In

N. Saxena (ed.), Recent advances in marine science and technology 2000.

PACON International, Honolulu, Hawaii.

41. Stewart, B. J., and L. L. McCarter. 2003. Lateral flagellar gene system of

Vibrio parahaemolyticus. Journal of Bacteriology 185:4508-4518.

42. Studer, S. V., M. J. Mandel, and E. G. Ruby. 2008. AinS quorum sensing

regulates the Vibrio fischeri acetate switch. J Bacteriol 190:5915-5923.

43. Tolonen, A. C., G. B. Liszt, and W. R. Hess. 2006. Genetic manipulation of

Prochlorococcus strain MIT9313: green fluorescent protein expression from an

RSF1010 plasmid and Tn5 transposition. Appl Environ Microbiol 72:7607-7613.

44. Tudor, J. J., J. J. Davis, M. Panichella, and A. Zwolak. 2008. Isolation of

predation-deficient mutants of Bdellovibrio bacteriovorus using transposon

mutagenesis. Appl Environ Microbiol:(in press).

45. Visick, K. G., and E. G. Ruby. 1996. Construction and symbiotic competence of

a luxA-deletion mutant of Vibrio fischeri. Gene 175:89-94.

46. Visick, K. L. 2005. Layers of signaling in a bacterium-host association. J

Bacteriol 187:3603-3606.

47. Visick, K. L., and M. J. McFall-Ngai. 2000. An exclusive contract: Specificity

in the Vibrio fischeri-Euprymna scolopes partnership. Journal of Bacteriology

182:1779-1787.

48. Visick, K. L., and E. G. Ruby. 1997. New genetic tools for use in the marine

bioluminescent bacterium Vibrio fischeri, p. 119-122. In J. W. Hastings, L. J.

Kricka, and P. E. Stanley (ed.), Bioluminescence and Chemiluminescence. John

Wiley and Sons, New York.

49. Visick, K. L., and E. G. Ruby. 2006. Vibrio fischeri and its host: it takes two to

tango. Curr Opin Microbiol 9:632-638.

50. Weinreich, M. D., A. Gasch, and W. S. Reznikoff. 1994. Evidence that the cis

preference of the Tn5 transposase is caused by nonproductive multimerization.

Genes Dev 8:2363-2374.

51. Wiegand, T. W., and W. S. Reznikoff. 1992. Characterization of two

hypertransposing Tn5 mutants. Journal of Bacteriology 174:1229-1239.

52. Wilson, M. M., and W. W. Metcalf. 2005. Genetic diversity and horizontal

transfer of genes involved in oxidation of reduced phosphorus compounds by

Alcaligenes faecalis WM2072. Appl Environ Microbiol 71:290-296.

53. Wolfe, A. J., D. S. Millikan, J. M. Campbell, and K. L. Visick. 2004. Vibrio

fischeri 54 controls motility, biofilm formation, luminescence, and colonization.

Applied and Environmental Microbiology 70:2520-2524.

54. Yanofsky, C., and V. Horn. 1981. Rifampicin resistance mutations that alter the

efficiency of transcription termination at the tryptophan operon attenuator. Journal

of Bacteriology 145:1334-1341.

55. Yip, E. S., K. Geszvain, C. R. DeLoney-Marino, and K. L. Visick. 2006. The

symbiosis regulator RscS controls the syp gene locus, biofilm formation and

symbiotic aggregation by Vibrio fischeri. Mol Microbiol 62:1586-1600.

56. Yip, E. S., B. T. Grublesky, E. A. Hussa, and K. L. Visick. 2005. A novel,

conserved cluster of genes promotes symbiotic colonization and sigma-dependent

biofilm formation by Vibrio fischeri. Mol Microbiol 57:1485-1498.

57. Yun, J. I., K. M. Cho, J. K. Kim, S. O. Lee, K. Cho, and K. Lee. 2007.

Mutation of rpoS enhances Pseudomonas sp. KL28 growth at higher

concentrations of m-cresol and changes its surface-related phenotypes. FEMS

Microbiol Lett 269:97-103.

58. Zhou, M., A. Bhasin, and W. S. Reznikoff. 1998. Molecular genetic analysis of

transposase-end DNA sequence recognition: cooperativity of three adjacent base-

pairs in specific interaction with a mutant Tn5 transposase. Journal of Molecular

Biology 276:913-925.

59. Zhou, M., and W. S. Reznikoff. 1997. Tn5 transposase mutants that alter DNA

binding specificity. Journal of Molecular Biology 271:362-373.

CHAPTER 3

BRIGHT MUTANTS OF VIBRIO FISCHERI ES114 REVEAL

CONDITIONS AND REGULATORS THAT CONTROL

BIOLUMINESCENCE AND EXPRESSION OF THE LUX OPERON1

1Noreen L. Lyell, Anne K. Dunn, Jeffrey L. Bose, and Eric V. Stabb. 2010. J. Bacteriol.

192: 5103-5114. Reprinted here with permission of publisher.

Abstract

Vibrio fischeri ES114, an isolate from the Euprymna scolopes light organ, produces little bioluminescence in culture but is ~1000-fold brighter when colonizing the host. Cell-density-dependent regulation alone cannot explain this phenomenon, because cells within colonies on solid medium are much dimmer than symbiotic cells despite their similar cell densities. To better understand this low luminescence in culture, we screened

~20,000 mini-Tn5 mutants of ES114 for increased luminescence and identified 28 independent “luminescence-up” mutants with insertions in 14 loci. Mutations affecting the Pst phosphate uptake system led to the discovery that luminescence is upregulated under low-phosphate conditions by PhoB, and we also found that ainS, which encodes an autoinducer synthase, mediates repression of luminescence during growth on plates.

Other novel luminescence-up mutants had insertions in acnB, topA, tfoY, phoQ, guaB and two specific tRNA genes. Two loci, hns and lonA, were previously described as repressors of bioluminescence in transgenic Escherichia coli carrying the light-generating lux genes, and mutations in arcA and arcB were consistent with our report that Arc represses lux. Our results reveal a complex regulatory web governing luminescence and show how certain environmental conditions are integrated into regulation of the pheromone-dependent lux system.

Introduction

Vibrio fischeri is a valuable model for examining bioluminescence, pheromone signaling, and symbiotic bacteria-animal interactions. Studies of V. fischeri‟s mutualistic interactions have gained momentum since the discovery that this bacterium‟s light organ symbiosis with the Hawaiian bobtail squid, Euprymna scolopes, can be reconstituted in the laboratory (54, 70, 83). Moreover, the bioluminescence induced by V. fischeri in the host light organ is a pheromone-mediated behavior, making this an attractive system for examining environmental influences on bacterial pheromone signaling in a natural infection. Largely because of interest in this symbiosis, strain ES114, which was isolated from the E. scolopes light organ, has become the experimental strain of choice for many studies of V. fischeri.

The genetic basis of bioluminescence in V. fischeri ES114 is fundamentally similar to that of other characterized V. fischeri strains (31, 32). The lux genes responsible for bioluminescence, luxABCDE and G are found together with the regulatory genes luxR and luxI and are arranged with luxR divergently transcribed from the luxICDABEG operon, as shown in Figure 3.1 (24, 25, 56). Light is generated when luciferase, comprised of LuxA and LuxB, binds to FMNH2, O2, and an aliphatic aldehyde, and converts these substrates to FMN, water, and an aliphatic acid (35, 76).

LuxC, LuxD, LuxE, and LuxG (re)generate luciferase‟s aldehyde and FMNH2 substrates

(12, 64).

3OC6 C8 3OC6 AI-2 AI-2 C8 C8 3OC6 AI-2 C8 3OC6 AI-2 AI-2

AI-2 LuxP C8 AinR LuxQ 54 Hfq P- LuxO sRNA Hfq C8- LuxR 3OC6- LuxR LuxU LuxO

(C8 and AI-2 lead to less phosphorylated LuxO) litR

LuxR LitR

luxR luxI luxC luxD luxA luxB luxE luxG “lux box”

bioluminescence

LuxI HO OH AinS B- O O LuxS O O O O HO N O H N HO O O H O (3OC6) (C8) (AI-2) IM OM

3OC6 C8 AI-2

Fig. 3.1: Model of autoinducer-mediated regulation of bioluminescence in V. fischeri. Labeled open arrows correspond to genes, and the structures of three autoinducer molecules are presented with their respective synthases. The AI-2 structure is inferred from studies in V. harveyi (13) but has not been identified in V. fischeri. Interactions between autoinducers, proteins, genes, and small RNA (sRNA) are indicated. For simplicity, multimerization of proteins is not shown. This model is derived in part from experimental results but in some aspects relies on genomic predictions. For details see reference 73.

The remaining genes in this cluster, luxI and luxR, underlie a pheromone- mediated regulatory mechanism often referred to as quorum sensing. LuxI generates the membrane-permeable autoinducer pheromone N-3-oxohexananoyl-L-homoserine lactone

(3-oxo-C6-HSL) (23, 41). As cell density increases, 3-oxo-C6-HSL accumulates until

reaching a threshold concentration, whereupon it binds LuxR and together they activate transcription of luxICDABEG (24, 25, 77). As a result, the lux genes are most highly expressed at high cell densities, when the bacteria have reached a “quorum” (28). Like many bacterial quorum sensing systems, the lux operon constitutes a positive feedback circuit, because the 3-oxo-C6-HSL produced by LuxI leads to increased transcription of luxICDABEG. As a result, environmental regulatory inputs to the lux system can be amplified and spread in a population.

V. fischeri generates two additional autoinducers; N-octanoyl-L-homoserine lactone (C8-HSL) produced by AinS (44) and AI-2 (46), which may be a furanosyl borate diester as it is in Vibrio harveyi (13). As Figure 3.1 illustrates, AI-2 and C8-HSL presumably function through distinct receptors that both act via LuxU and LuxO, Hfq, and a small RNA to increase levels of LitR, which in turn increases luxR expression (26,

47, 58). C8-HSL can also activate LuxR directly (47). Although C8-HSL is a weaker activator of LuxR than 3-oxo-C6-HSL (67), ES114 produces more C8-HSL in broth cultures (73), and under these conditions it is the main activator of bioluminescence in

ES114 (47).

Interestingly, despite conserved lux circuitry, ES114 and other isolates from E. scolopes are much dimmer in culture than previously studied V. fischeri strains (8).

ES114 colonies on solid media are not visibly luminescent, and cells in these colonies produce ~1000-fold less luminescence than do symbiotic ES114 cells, despite achieving similar high population densities. Thus, cell density alone cannot account for the dim luminescence of ES114 in culture, and environmentally responsive regulators must also play a critical role in regulating the luxICDABEG operon.

Relatively little is known about environmental influences on luxICDABEG expression in V. fischeri, particularly in ES114. CRP-mediated regulation of lux in response to glucose has been documented using transgenic lux-containing Escherichia coli (18-20), although the response of V. fischeri to glucose is less clear and may be strain-specific (8, 27). Similarly, the luminescence of strain MJ1 is inhibited by iron

(36), but iron does not affect ES114 luminescence (8). We found that the redox- responsive ArcA/ArcB system repressses lux (10); however, as we report here,

ArcA/ArcB does not account for aeration-dependent regulation of luminescence (73).

Other genes affecting luminescence in V. fischeri ES114 have been reported (38, 79, 84), but to date no directed study has screened for regulators that could account for the relatively low lux expression by ES114 in culture or its induction/derepression upon entering the host.

Control of bacterial pheromone systems such as lux by environmentally responsive regulators is widespread and has important functional implications, yet the significance of this phenomenon remains obscure. Therefore, our goal is to elucidate regulators and environmental contingencies that control induction of the lux operon. In this study, we screened for “luminescence-up” mutants of V. fischeri ES114, which led to the identification of conditions and regulators that affect bioluminescence and expression of the lux operon.

Materials and Methods

Bacteria and media

V. fischeri strain ES114, originally isolated from an E. scolopes light organ (8), was the wild-type strain used throughout this study. V. fischeri mutants KV2801 (79),

KV2850 (79), KV1651 (38), and KV2655, with disruptions in ptsI, crr, phoB, and phoU, respectively, were obtained from Karen Visick. E. coli strains DH5α (34) or DH5αλpir

(21) were transformed by plasmids in the cloning steps outlined below. Transfer of plasmids to V. fischeri was accomplished by triparental matings using conjugative helper strain CC118λpir pEVS104 as previously described (72). E. coli was grown in LB medium (57) or BHI medium (Bacto) and V. fischeri strains were grown in one of three medium types: (i) LBS, which contained, per liter of water, 10 g of tryptone, 5 g of yeast extract, 20 g of NaCl, and 50 mM Tris (pH 7.5); (ii) SWTO, which contained, per liter of total volume, 5 g of tryptone, 3 g of yeast extract, 3 ml of glycerol, 700 ml of Instant

Ocean mixed to 36 ppt (Aquarium Systems, Mentor, Ohio), and an additional 170 mM

NaCl; (iii) FMM, which contained per liter, 950 ml of water, 378 μl of 1 M NaPO4 (pH

7.5), 50 ml of 1 M Tris (pH 7.5), 3 mg of FeSO4·7H2O, 13.6 g of MgSO4·7H2O, 0.59 g of

NH4Cl, 0.83 g of KCl, 19.5 g of NaCl, 1.62 g of CaCl2·2H2O, 1 g of Casamino Acids, and 3 ml of glycerol. Solid media were prepared with 15 mg ml-1 agar for plating. For selection of E. coli, chloramphenicol (CAM) and kanamycin (KAN) were added to LB at final concentrations of 20 and 40 μg ml-1, respectively, and 150 μg ml-1 erythromycin

(ERM) was added to BHI. For selection of V. fischeri on LBS, CAM, ERM, and KAN were used at concentrations of 2, 5, and 100 μg ml-1, respectively. 5-bromo-4-chloro-3-

indolyl-β-D-galactoside (X-Gal) and isopropyl-β-D-thiogalactoside (IPTG) were added to media at final concentrations of 60 and 50 μg ml-1, respectively.

Transposon mutagenesis

Transposon mutants were generated as previously described (1, 48). Briefly, pEVS170 was transferred to ES114 by triparental matings (72). Following overnight incubation, mating spots were resuspended in LBS and dilution plated onto LBS supplemented with ERM. Plates were incubated overnight at ~24˚C, and images were generated using a Fluor-S Max-2 imager (Bio-Rad Laboratories, Hercules, CA) set to high-sensitivity chemiluminescence mode with a 20-min exposure. Colonies that were more luminescent than JRM100, an ERM-resistant derivative of ES114 (52), were patched onto LBS supplemented with ERM and imaged following overnight incubation to confirm their luminescence phenotype. To ensure that ermR colonies carried transposon insertions and were not harboring pEVS170, mutants were screened for kanamycin resistance, which is present on pEVS170 outside the transposon, and KAN- resistant mutants were discarded.

Genetic techniques and analyses

Plasmids were generated using standard techniques. DNA ligase and restriction enzymes were obtained from New England Biolabs (Beverly, MA). Oligonucleotides were synthesized by Integrated DNA Technologies (Coralville, IA). PCR was performed with an iCycler (Bio-Rad Laboratories) using KOD DNA Polymerase (Novagen,

Madison, WI) or Phusion high-fidelity polymerase (Finnzymes, Finland). Plasmids used

for cloning were purified using Qiagen miniprep kits (Valencia, CA) or the GenElute plasmid miniprep kit (Sigma-Aldrich, Inc., St. Louis MO). For supercoiling analyses, plasmids were purified using the ChargeSwitch-ProDNA plasmid miniprep kit

(Invitrogen, Carlsbad, CA). DNA was purified from PCR, digestion, and ligation reactions with the DNA Clean and Concentrator-5 kit (Zymo Research, Orange, CA). To clone PCR products into the pCR-BluntII-TOPO plasmid, we used the ZeroBlunt-TOPO

PCR cloning kit (Invitrogen) and screened for white colonies on plates containing X-Gal.

Cloned PCR products were sequenced at the University of Michigan DNA Sequencing

Core Facility, and sequences were compared to the ES114 genomic database using

Sequencher 4.1.2 (Gene Codes Corp., Ann Arbor, MI) to ensure that no unintended base pair changes were incorporated.

Plasmids pJLB52 (10), pJLB114, pAS5, pNL4, pNL18, pNL75, pNL6, pNL72, and pCL112 (47) were used to mobilize native copies of arcA, arcB, acnB, topA, lonA, pstA, hns, phoQ, and ainS, respectively, into mutants with transposon insertions in these genes. A description of new complementation plasmid construction follows. In each case the cloned gene was expressed by including ~500 bp of upstream sequence and/or by a promoter(s) on the vector. For example, the pstA and phoQ genes appear to be toward the distal end of operons and were cloned by themselves such that they would be expressed by the lacZ promoter on the vector. arcB was PCR amplified using primers

ARC B1 (5‟ CCG CCC TAG GCA GGG TAT GTT TAT GAA GCA GTT AA 3‟) and

ARC B2 (5‟ GGG GTA CCG TGT TGC GGC AAA TAG TAC CTT CTT C 3‟), and the blunt product was cloned into pCR-BluntII-TOPO to generate pJLB106. pJLB106 was linearized with AvrII and ligated to XbaI-digested pVSV105 (22), generating pJLB110,

which was then digested with KpnI and self-ligated to excise the pCR-BluntII-TOPO vector, generating pJLB114. acnB was PCR amplified using primers JBACNB5 (5‟

GCG CCA TAA GTC GTA TGT TGT TTG TTG TGG G 3‟) and JBACNB7 (5‟ CCA

GCC CT TAA ATA AAA AAG CAG CCA ATT GCC 3‟), and the blunt product was ligated directly into HpaI-digested pJLB103 to generate pJLB130. The vector pJLB103, which contains the R6K origin of replication and encodes KAN resistance, was derived from pVSV104 (22) by deleting the pES213 origin on a BamHI fragment. pJLB130 was digested with AvrII and KpnI, and the fragment containing acnB was ligated into XbaI- and KpnI-digested pVSV105 (22) to generate pAS5. topA was PCR amplified using primers pr_NL3 (5‟ CAG CCT CAG AAA TGG ATT TTT TAT CGC TCA TAA G 3‟) and pr_NL4 (5‟ GAG CCG CAT TTC TGC AGC TCT TTC 3‟), and the blunt product was cloned into pCR-BluntII-TOPO to generate pNL2. pNL2 was digested with EcoRV and the topA-containing fragment was ligated into SmaI-digested pVSV105 (22) to generate pNL4. lonA was PCR amplified using primers pr_NL1 (5‟ GGC CGT TTA

CCT GTA ACA ACA ACG GAC 3‟) and pr_NL2 (5‟ GAG TGA CAA GTC ATT TCG

ACT TGT CAG CCC 3‟), and the blunt product was cloned into pCR-BluntII-TOPO to generate pNL3. pNL3 was digested with XbaI, and the lonA-containing fragment was ligated into SpeI-digested pVSV105 (22) to generate pNL18. hns was PCR amplified using primers pr_NL11 (5‟ GCC AAA CCC AGA GCT ATA AGC GGG GGC 3‟) and pr_NL12 (5‟ TTT CGA GCA ATA ATA CGT TTC TAA ATG TAA TAA AAT GAA A

3‟). pstA was PCR amplified using primers pr_NL13 (5‟ GCG CTA GTT GTT GGC

ATT GCA ATG GGA GCT GC 3‟) and pr_NL14 (5‟ AGA CAG GTG GTT AAC ATC

CAT CGG TGA TAG 3‟). phoQ was PCR amplified using primers pr_NL84 (5‟ GCC

TAA CGG TAC TAA AAA GCA TTC TGT ATG 3‟) and pr_NL85 (5‟ GAT GAA GAG

CAT GAT TAT TAT TCT GAT GGA GAG ATA TTG G 3‟). The blunt hns-, pstA-, and phoQ-containing products were cloned into SmaI-digested pVSV105 (22) to generate pNL6, pNL75, and pNL72 respectively.

q The lacI -PA1/34-luxC allele on pJLB101 was moved into the transposon mutants as described previously (11). Allelic exchange was confirmed by PCR and by IPTG

q inducibility of luminescence in the resulting strains. The lacI -PA1/34-luxC allele from plasmid pJLB101 was crossed into the genomes of mutants EMH3, EMH5, EMH6,

EMH7, EMH9, EMH12, EMH13, SLV4, SLV5, SLV10, SLV15, SLV16, SLV20,

SLV29, SLV30, SLV32, SLV33, SLV41, SLV42, SLV43, NL1, NL3, NL4, NL6 and

NL8 to generate strains NL18, NL19, NL20, NL21, NL22, NL23, NL24, NL25, NL26,

NL27, NL28, NL29, NL30, NL31, NL32, NL33, NL34, NL35, NL36, NL37, NL38,

NL39, NL40, NL41 and NL42, respectively. Similarly, the ∆litR::kanR allele on plasmid pMF7 (26) was crossed into the genome of NL2 using two-step allelic exchange to generate the mutant NL11.

To generate a ∆ainR allele on plasmid pNL30, the 1,350-bp region upstream of ainR was PCR amplified using primers pr_NL27 (5‟ GTA CTC ATA ACA CCA CTA

CCT ATT TTT ACT ATA CTG 3‟) and pr_NL28.3 (5‟ GGG CCT AGG CAT TTA TAT

AAA ACT CAC TGA TTT CGA AGT TT 3‟), and the product was cloned into pCR-

BluntII-TOPO plasmid to generate pNL28. The 1,480-bp region downstream of ES114 ainR was PCR amplified using primers pr_NL29 (5‟ GGG GCC TAG GTA ACA CCG

ATA AAA AAA TAG CCA GAA C 3‟) and pr_NL30 (5‟ CCC CAC TAG TCA TGA

CTC TGT TGC GGG TCT TGA TGA AGC T 3‟). AvrII and SpeI sites incorporated

into the PCR product on primers pr_NL29 and pr_NL30 were digested with these enzymes, and this fragment was ligated into AvrII-digested pEVS118 (21) to generate pNL29. Plasmids pNL28 and pNL29 were linearized with AvrII and ligated together to generate pNL30, which contains upstream and downstream sequences fused at the ∆ainR allele, with the ainR start and stop codons separated only by the 6-bp AvrII recognition sequence. The mutant strain NL43 (∆ainR) was generated by crossing the mutant ∆ainR allele from plasmid pNL30 into ES114. Replacement of ainR with the ∆ainR allele in

NL43 was confirmed by PCR using primers pr_NL35 (5‟ GAG TCC GTT AGC AAG

GTC ACA CTT TGT TG 3‟) and pr_NL36 (5‟ ACC CAA AAC GTA AGA CCA TTG

GTA TGC G 3‟).

The ∆ainSR allele was generated on plasmid pNL32 by PCR amplification of the

1,430-bp region upstream of ainS using primers pr_NL62 (5‟ GGC GCT TTA CCG TTT

GGT GAA AAC TTA CTT C 3‟) and pr_NL63 (5‟ GGG CCT AGG CTA CTC TTT

TAT AAA TTC ATA TTG CAG GTT TT 3‟). This product was cloned into pCR-

BluntII-TOPO plasmid to generate pNL31. Plasmids pNL29 and pNL31 were linearized with AvrII and ligated together to generate pNL32, which contains the upstream and downstream sequences fused at the ∆ainSR allele, with the ainS start and ainR stop codons separated by the 6-bp AvrII recognition sequence. Crossing the mutant ∆ainSR allele from plasmid pNL32 into ES114 generated the mutant strain NL55 (∆ainSR).

Replacement of ainSR with the ∆ainSR allele was confirmed by PCR using primers

DMC2 (5‟ GGC GGT ACC AGA ACC AAG ACC TGC TCG TGC TAA 3‟) and pr_NL36.

Luminescence and fluorescence measurements in culture

Overnight V. fischeri cultures were diluted 1:1,000 in 50 ml of LBS or SWTO in

250-ml flasks and then incubated with shaking (200 rpm) at 24˚C. At regular intervals,

500-μl samples were removed and the optical density at 595 nm (OD595) was measured with a BioPhotometer (Brinkman Instruments, Westbury, NY). Relative luminescence was measured with a TD-20/20 luminometer (Turner Designs, Sunnyvale, CA), immediately following shaking to aerate the sample. Specific luminescence was calculated as the luminescence per OD595 unit. Luminescence images of strains patched on plates and incubated overnight at ~24˚C were obtained with a Bio-Rad Fluor-S Max2 imager. Fluorescence of green fluorescent protein (GFP) expressed from reporter plasmids pJLB36 and pJBL38 (10) was measured with a TD-700 fluorometer (Turner

Designs) with excitation and emission filters of 486 nm and >510 nm, respectively.

Fluorescence values for strains carrying the promoterless vector pVSV33 (22) were subtracted as background. Unless otherwise indicated, the reported mean fluorescence for cultures was between OD595 2.0 and 2.8, a range in which ES114 specific luminescence is approximately constant.

Results

Isolation of luminescence-up mutants of V. fischeri ES114

We screened ~20,000 mini-Tn5 mutants of ES114 for increased luminescence and isolated 30 candidate luminescence-up clones that displayed consistent and significant increases in luminescence. The chromosomal location of the transposon insertions in these strains and their luminescence phenotypes are summarized in Table 3.1. Three of

these mutants are likely siblings, because they have insertions in the same location in hns and originated from the same mating. Thus, our screen yielded 28 independent luminescence-up mutants of ES114 distributed over several loci (Table 3.1).

Table 3.1: Analysis of luminescence-up transposon mutantsa

aHorizontal arrows indicate disrupted genes (shaded) and flanking (open) open reading frames (ORFs). Vertical arrows correspond to locations of transposon insertions, and the insertion location is provided in bp relative to the A of the ATG start codon. All mutants were patched onto LBS and SWTO plates, and representative negative images of patches are shown, such that darker patches indicate greater bioluminescence. †, the contrast was increased to visualize the luminescence phenotype of mutant patches. NV, patches were not visible even with enhanced contrast. VAR, variable result as discussed in the text. All mutants were also grown in LBS and SWTO broth, and the average maximum luminescence/OD595 relative to ES114 ± standard error (n ≥ 2) is reported.

The mutants displayed a wide range of luminescence phenotypes that were from

2-fold to more than 1000-fold brighter than their wild-type parent (Table 3.1). When grown in broth, all of the mutants still displayed cell-density dependent regulation of luminescence (data not shown); however, with the exception of ainS mutants, they achieve brighter luminescence than the wild type in broth and some induce luminescence at lower cell densities (data not shown). Most mutants‟ luminescence phenotypes varied depending on whether the cells were grown on plates or in broth and whether they were grown in LBS or SWTO. Both LBS and SWTO contain tryptone and yeast extract, however, LBS is buffered and supplemented with NaCl, whereas SWTO is unbuffered, supplemented with marine salts and glycerol, and is near marine osmolarity (71).

Mutants with insertions in arcA, arcB, acnB, lonA, tfoY, guaB, and specific tRNA genes showed greater increases in luminescence relative to ES114 when grown in SWTO than in LBS (Table 3.1). In contrast, mutants with insertions in pstA, pstC, hns, and phoQ were comparatively brighter than ES114 when cultured in LBS (Table 3.1). Only the topA mutants showed similar luminescence-up phenotypes in both LBS and SWTO

(Table 3.1). We found an unexpected result, discussed below, in that ainS mutants were much brighter than ES114 on plates (Table 3.1), despite being dimmer than ES114 in broth (Table 3.1), as was previously shown (47).

Comprehensiveness of the screen

Based on previous work with mini-Tn5 derivatives (1, 39), we estimated that the

~20,000 mutants screened should be approaching saturation of nonessential genes.

Consistent with this prediction, multiple independent mutants were isolated at most of the loci identified, including all of the loci where dramatic and easily detected luminescence- up effects were observed (Table 3.1). On the other hand, some mutants of V. fischeri previously identified as having luminescence-up phenotypes in broth were not found in our study. These strains were patched onto LBS to test whether they would have been detected in our screen. Strains CL42 (47) and KV2801 (79), containing mutations within luxO and ptsI (E1), respectively, were dim on plates and would not have been identified in our screen (data not shown). A crr mutant (KV2850) yielded patches that were only marginally brighter on plates than ES114 (data not shown), and the phenotype was inconsistent; therefore, it is not surprising that a crr mutant was not isolated in our screen.

A phoU mutant (KV2655) was brighter than ES114 on plates (data not shown), and this was missed in our screen; however, insertions were identified in pstA and pstC, which are upstream of phoU in what appears to be a phosphate transport operon. Taken together, these results suggest that continuing to screen additional transposon mutants under these conditions would yield relatively few additional new insights, although screening under different growth conditions might reveal novel mutants missed here.

Complementation of mutants

To confirm that the luminescence-up phenotypes resulted from the disruption of the genes identified, we genetically complemented the arcA, arcB, acnB, topA, lonA,

pstA, phoQ, and hns mutants by providing the respective native genes in trans on low- copy-number shuttle plasmids (Fig. 3.2). In all strains except the pstA and phoQ mutants, wild-type luminescence was restored following reintroduction of the native gene. The pstA mutant maintained its luminescence-up phenotype (Fig. 3.2B), and this was likely due to polar effects on the downstream phoU. The relationship between this phosphate- uptake system and luminescence is explored further below. Although the luminescence phenotype of the phoQ mutant was not complemented in LBS medium (Fig. 3.2B), it was complemented by phoQ in trans during growth in minimal medium (see below). There is no apparent cotranscribed gene downstream of phoQ (Table 3.1) making polar effects of this insertion unlikely. We further describe the effects of phoQ and Mg2+ on luminescence below. Because the ainS mutants only displayed a luminescence-up phenotype on plates, we evaluated genetic in trans complementation of ainS in patches using pCL112 (47), which restored wild-type-like dim luminescence (data not shown).

We did not attempt to complement mutants with insertions in tfoY, guaB or tRNA genes, which had relatively modest luminescence-up phenotypes. Given the apparent lack of cotranscribed genes downstream of tfoY (Table 3.1), it seems unlikely that the phenotype of this insertion mutant is caused by polar effects.

Fig. 3.2: Genetic complementation of transposon mutants. Luminescence-up mutants were complemented with the respective complete gene carried on shuttle vector pVSV105. ES114 and mutants carrying the empty pVSV105 vector were included as controls. (A) Complementation of mutants with increased luminescence in SWTO. (B) Complementation of mutants with increased luminescence in LBS. Data for one representative mutant from each locus are shown, including SLV41 (arcA), NL3 (arcB), SLV4 (acnB), EMH12 (topA), SLV32 (lonA), SLV10 (pstA), SLV15 (hns), and SLV16 (phoQ). Error bars represent standard errors (n = 2).

Instability of acnB mutants

The observable phenotypes of the luminescence-up mutants remained stable with the notable exception of mutants with insertions in acnB, which encodes the tricarboxylic acid cycle enzyme aconitase. There is no other aconitase gene apparent in the V. fischeri genome, and acnB mutants all grew relatively poorly on aerobic streak plates. Faster- growing suppressors arose frequently and had dim luminescence, like that of wild type.

We were able to confirm complementation of an acnB mutant in trans (described above) by curing the complementing plasmid with concomitant restoration of slow growth and bright luminescence. However, we found further genetic manipulation of these strains difficult due to the rapid appearance of suppressors. We have therefore omitted acnB mutants from genetic analyses below and will describe the influence of acnB on luminescence in greater detail once the nature of the suppressors is more fully understood.

Dependence of luminescence-up phenotypes on the native luxICDABEG promoter

We hypothesized that the luminescence-up phenotype of the mutants in Table 3.1 was due to regulatory effects on lux gene expression; however, the bright phenotypes of these mutants might alternatively be due to metabolic effects caused, for example, by increasing the availability of the FMNH2 or O2 substrates for LuxAB. To test this

q possibility, we placed a nonnative IPTG-inducible promoter construct, lacI -PA1/34 between luxI and luxCDABEG (11), into the chromosomes of the mutant strains, with the exception of acnB mutants. Transposon insertions in arcA, arcB, lonA, pstA, pstC, hns, tfoY, phoQ, guaB, ainS, and the tRNA‟s had no effect on luminescence when luxCDABEG was expressed from this nonnative promoter, either with or without addition of IPTG (data not shown). Thus, the luminescence-up phenotype associated with most insertions was dependent on the native lux promoter and transcript.

Only topA mutations yielded an increased level of luminescence in the lacIq-

q PA1/34-luxC background. topA mutations in the lacI -PA1/34-luxC background led to enhanced luminescence at low ODs (Fig. 3.3A) but did so to a lesser extent than the ~35

to 40-fold effect seen in the native lux promoter background (Table 3.1 and Fig. 3.3B).

This suggests the topA mutants‟ luminescence-up phenotype may be due to combined effects that are both dependent and independent of the native lux promoter.

Fig. 3.3: Luminescence of topA mutants in SWTO. (A) Specific luminescence phenotype of JB22 as well as topA mutants NL23 and NL24, each of which have the q luxCDABEG genes controlled by lacI and the PA1/34 promoter. JB22 has the PA1/34- luxCDABEG allele in an otherwise wild-type (ES114) background. No IPTG was added. (B) Specific luminescence of wild-type (WT) ES114 as well as topA mutants EMH12 and EMH13. Averages of two replicate flasks are shown.

To test whether the luminescence-up mutations enhanced transcription from the luxI or luxR promoters, we moved reporter plasmids pJLB36 (PluxR-gfp) and pJLB38

(PluxI-gfp) into each mutant. Fluorescence data for these reporters were determined by growing the mutant strains in the medium (SWTO or LBS) that showed the greatest luminescence-up phenotype relative to the wild type. Consistent with our previous report

(10), the PluxR-gfp and PluxI-gfp reporters yielded ~2- and ~15-fold greater fluorescence, respectively, in either the arcA or arcB mutants than in parental strain ES114 (Figs. 3.4A and B). These reporters also yielded higher fluorescence in topA mutants than in ES114, although the relative influence on the PluxR-gfp was greater than in the arcA or arcB backgrounds (Figs. 3.4A and B). The PluxI-gfp reporter yielded ~4- and ~8-fold greater fluorescence in the pstC and hns backgrounds, respectively, though these mutations had no effect on the PluxR-gfp reporter (Figs. 3.4C and D). None of the other transposon insertions, with the possible exception of those in unstable acnB mutants (data not shown), had a significant (P < 0.05) effect on either reporter under these conditions. This lack of effect in most mutants may reflect limitations of these reporters. It is worth noting that arc and topA mutants have much larger effects on luminescence (Table 3.1) than on gfp reporter activity (Fig. 3.4), and similar results were obtained with lacZ reporters (data not shown). These plasmid-borne reporters are maintained in ~9.4 copies per genome (21), potentially titrating out regulators, although at least in the case of ArcA effects of similar magnitude were seen with lux-gfp reporters on a plasmid or on the chromosome in single copy (10). Taken together, it is perhaps not surprising that mutations with moderate effects on luminescence have no discernible influence on these reporters.

Fig. 3.4: Effects of transposon mutations on luxR and luxI promoter-gfp reporters. Specific fluorescence generated from reporter plasmids pJLB36 (PluxR-gfp) (A amd C) or pJLB38 (PluxI-gfp) (B and D) harbored in ES114 or in luminescence-up mutants is shown. Reporters were assayed in mutants grown in either SWTO (A and B) or LBS (C and D), depending on which medium yielded the greatest effect on luminescence. Fluorescence from ES114 harboring the promoterless parent vector pVSV33 was subtracted as background. Reporters were tested in each mutant described in Table 1, and data for one representative mutant from each locus are shown, including SLV41 (arcA), NL3 (arcB), EMH12 (topA), SLV32 (lonA), SLV10 (pstA), SLV30 (pstC), SLV15 (hns), EMH7 (tRNA-Met), NL4 (tRNA-Thr), NL1 (tfoY), SLV16 (phoQ), EMH5 (guaB), and NL2 (ainS). In all panels, data represent the average specific fluorescence when the culture OD595 was between 2.0 and 2.8; a range in which specific luminescence is constant for these strains. Averages and standard errors were calculated from 8 to 12 distinct samples

taken from two independent replicate flasks of each examined strain. Mutants with insertions at the same locus were analyzed together in two separate experiments with similar results, and data for one representative mutant from one experiment are shown.

Role of ainS-mediated signaling in repression of luminescence

The luminescence-up phenotype of ainS mutants on plates was unexpected given their previously reported diminished luminescence in broth (47), and we examined these mutants further. The AinS-produced pheromone C8-HSL is thought to act as a signal through two pathways (Fig. 3.1). In one pathway C8-HSL binds to LuxR and activates transcription, much like the LuxI product 3-oxo-C6-HSL. C8-HSL is a weaker activator of LuxR (67) and can apparently compete with the stronger activator 3-oxo-C6-HSL to dim luminescence (47). In the second pathway, C8-HSL is thought to be sensed by AinR

(30), which as illustrated in Fig. 3.1 ultimately leads to an increase in the regulator LitR

(26, 47, 58).

To explore which C8-HSL-responsive pathway is responsible for the luminescence-up phenotype of ainS mutants on plates, we used targeted mutants lacking components of these signaling pathways and compared their luminescence in patches on solid media. Table 3.2 shows our results, which include the following: (i) The

„luminescence up‟ phenotype associated with loss of ainS was independent of ainR, indicating that this phenotype is not simply due to a lack of C8-HSL-AinR; (ii) the luminescence-up phenotype of the ainS mutant was dependent on luxI, indicating a key role for 3-oxo-C6-HSL in generating the bright luminescence; (iii) the luminescence-up phenotype of ainS mutants was also dependent on both luxS and litR, and such dependence could be relieved by inactivating luxO, even though a luxO mutation itself did not result in bright luminescence on plates (data not shown). When taken together

and compared with the model of signaling presented in Fig. 3.1, these data are consistent with the luminescence-up phenotype of ainS mutants on plates resulting from the removal of competition for LuxR activation between C8-HSL and the stronger inducer 3-oxo-C6-

HSL. Moreover, under these conditions it appears that LuxS and AI-2 are sufficient for the signaling through LuxO necessary to generate LitR, rendering AinS and C8-HSL dispensable in this regard.

Table 3.2: Luminescence analysis of mutants involved in the AinS signaling pathways

Strain Reference Genotype Patches on

LBS platesa

ES114 (8) Wildtype -

NL2 This study ES114 ainS::mini-Tn5-ermR +

NL43 This study ES114 ∆ainR -

NL55 This study ES114 ∆ainSR +

CL24 (47) ES114 luxS::kanR -

NL11 This study ES114 ainS::mini-Tn5-ermR litR::kanR -

CL39 (46) ES114 luxI- ainS::catR -

CL41 (46) ES114 ainS::catR luxS::kanR -

CL64 (47) ES114 ainS::catR luxO::kanR +

CL90 (46) ES114 luxS::kanR luxO::kanR -

CL91 (46) ES114 ainS::catR luxS::kanR luxO::kanR + aAll strains were patched onto LBS plates, and the results of the negative images are reported as follows: +, patches were bright; -, indicating patches were not bright (compared to ainS mutant patches [Table 3.1]).

Role of PhoB and phosphate in regulation of luminescence

The luminescence-up phenotypes of the phoU, pstA, and pstC mutants (Table 3.1 and data not shown) suggested a role for inorganic phosphate (Pi) in the regulation of luminescence. In E. coli, the pst genes encode a high-affinity Pi transport system that is active under low-Pi conditions (86), and mutations in the pst genes result in increased expression of genes associated with the pho regulon in a mechanism dependent on the

PhoR/PhoB two-component regulatory system (74). At low Pi, the sensor PhoR phosphorylates the response regulator PhoB, which then activates the transcription of specific genes within the pho regulon (49, 50). PhoU, which is encoded at the distal end of the pst operon, counteracts PhoB when [Pi] is high (59).

To test whether luminescence and lux regulation are tied to the pho regulon, luminescence was measured for ES114 as well as pst, phoU and phoB transposon- insertion mutants cultured at relatively high and low Pi concentrations (Fig. 3.5A). In a medium with reduced Pi levels, the luminescence of ES114 was ~10-fold higher an OD595 of 0.5, and this response to low Pi was absent in a phoB mutant (Fig. 3.5A). Moreover, mutants with insertions in phoU, pstA, or pstC displayed constitutively high levels of luminescence regardless of [Pi]. These data suggest that luminescence is controlled in part by the PhoR/PhoB system in response to low Pi.

To further examine the mechanism of Pi-mediated lux regulation, ES114 cells containing PluxR-gfp and PluxI-gfp reporter plasmids were grown in high and low Pi media

(Fig. 3.5B). Under conditions with lower Pi levels, the activities of PluxR-gfp and PluxI-gfp each increased ~2-fold. As a control we tested a constitutive promoter-gfp reporter and found no relative effect of high and low Pi (data not shown). Thus, low Pi leads to an

increase in activity from the lux promoters, which may account for the luminescence-up phenotypes observed in phoU, pstA and pstC mutant strains.

Fig. 3.5: Effects of Pi concentration on luminescence and lux promoter-reporters. (A) Specific luminescence at OD595 of 0.5 for strains ES114 (wild type), KV1651 (phoB::pTMB28), KV2655 (phoU::mini-Tn5-ermR), SLV10 (pstA::mini-Tn5-ermR), and SLV30 (pstC::mini-Tn5-ermR) grown in FMM containing 378 μM (open bars) or 37.8 μM (grey bars) phosphate. (B) Specific fluorescence from PluxR-gfp and PluxI-gfp reporters on plasmids pJLB36 and pJLB38, respectively, harbored in ES114 grown in FMM containing 378 μM (open bars) or 37.8 μM (grey bars) phosphate. Fluorescence from ES114 harboring promoterless parent vector pVSV33 was subtracted as background. Data represent average specific fluorescence when culture OD595 values were ~0.5. Bars indicate standard errors (n = 2). *, significant difference between high and low Pi conditions as determined with Student‟s t test (P ≤ 0.01).

Effect of PhoQ and Mg2+ on luminescence

We also identified luminescence-up mutants with insertions in phoQ (Table 3.1).

In other bacteria PhoQ together with the response regulator PhoP act as a two-component regulatory system that responds to low Mg2+ (42, 63, 78). Based on a simple model

drawn from the luminescence-up phenotype of phoQ mutants and the function of PhoQ in other systems, we predicted a PhoQ-dependent repression of luminescence when Mg2+ was relatively low.

Fig. 3.6: Effect of Mg2+ concentration on luminescence. (A) Specific luminescence of V. fischeri strains ES114 (wild type), SLV16 (phoQ::mini-Tn5-ermR), JB22 (ES114 lacIq- q PA1/34-luxC), and NL29 (SLV16 lacI -PA1/34-luxC). (B) Specific luminescence of ES114 and SLV16 carrying the empty pVSV105 vector and SLV16 carrying pNL72, which contains native phoQ allele. Both panels report data from cultures grown in FMM, which contains 55 mM magnesium (open bars), or FMM with low (0.03 mM) Mg2+ (grey bars). *, significant difference in luminescence for that strain grown in low-Mg2+ medium relative to normal FMM as determined with Student‟s t test (P ≤ 0.05).

To further examine the effects of PhoP/PhoQ and Mg2+ on luminescence, we measured the luminescence of wild-type ES114, the constitutive lux-expressing strain

JB22 (11), and their respective phoQ mutants, SLV16 and NL29, grown in a defined medium with differing concentrations of Mg2+. Consistent with our prediction, low Mg2+

led to decreased luminescence in ES114 but not in the phoQ mutant (Fig. 3.6A).

Providing phoQ in trans restored low luminescence to the phoQ mutant (Fig. 3.6B).

However, we also observed an unexpected increase in luminescence at low Mg2+ in the phoQ mutant (Fig. 3.6A), suggesting a PhoQ-independent mechanism that functions counter to PhoQ to increase luminescence at low Mg2+. Moreover, low Mg2+ also decreased luminescence in both strains JB22 and NL29, in which luminescence is expressed from the constitutive nonnative PA1/34-luxC promoter (Fig. 3.6A). These data indicate that luminescence is influenced by Mg2+ through multiple mechanisms that are both PhoQ-dependent and PhoQ-independent.

Effect of GuaB and guanine on luminescence

GuaB and GuaA are required for the synthesis of GMP (55, 75), and we predicted that the insertion disrupting guaB would disrupt guanine synthesis. As predicted, in

FMM medium a severe growth defect of the guaB mutant was observed that was recovered by adding 0.25 mM guanine (data not shown). Furthermore, luminescence of the guaB mutant was restored to dimmer, wild-type levels by adding 0.15 mM guanine to

SWTO (data not shown).

Regulation of luminescence in response to aeration is topA-dependent

When ES114 is poorly aerated, its expression of luminescence is repressed (73); however, no mechanism for this regulation has yet been shown. The isolation of luminescence-up mutants with insertions in arcA, arcB, acnB, and topA (Table 3.1) suggested possible mechanisms for regulation of luminescence in response to culture

aeration, because ArcA/ArcB, aconitase, and DNA supercoiling, which is mediated in part by TopA, have been implicated in redox-dependent control of gene expression in E. coli (2, 6, 33). Due to the problems with suppressors arising in cultures of acnB mutants mentioned above, we limited our investigation to possible effects of ArcA/ArcB or TopA on aeration-dependent regulation of luminescence.

The redox-responsive ArcA/ArcB system represses luminescence presumably in response to reducing conditions (10). However, we found that ArcA cannot account for the repression of luminescence when cultures are grown in poorly aerated broth. In both wild type and a ∆arcA mutant, the luminescence of cells grown in poorly aerated flasks began to diverge from that of well-aerated cells at OD595 of ~0.7 (Fig. 3.7A). Thus, although the ∆arcA mutant is brighter than ES114 under both conditions, the effect of poor aeration on luminescence is similar in both strains.

We next examined the topoisomerase I (topA) mutants. In E. coli, DNA is more negatively supercoiled in anaerobically grown cultures, leading to changes in gene expression (17, 37). Similarly, we found that plasmid supercoiling in V. fischeri is affected by culture aeration (data not shown). Topoisomerase I removes negative supercoils from DNA (43, 80), and we hypothesized that topA mutations might prevent cells from using genomic supercoiling as part of their global response to aeration. We confirmed the supercoiling phenotypes of topA mutants, assaying the level of negative supercoiling by isolating plasmid DNA from the wild type and the topA mutant strain and examining them in chloroquine gels. As predicted, supercoiling was affected in a topA mutant background (Fig. 3.7B). We also examined the luminescence phenotype of a topA mutant grown in different aeration conditions. In contrast to the wild type and arcA

mutants, the luminescence of the topA mutant remained similar in both well aerated and poorly aerated cultures (Fig. 3.7A). This suggests that a topA-dependent effect on the level of negative supercoiling may regulate luminescence in response to aeration.

Fig. 3.7: Effect of topA mutations on aeration-dependent regulation of luminescence and supercoiling. (A) Effect of topA mutation on aeration-mediated luminescence. Specific luminescence of wild-type ES114 (squares), topA mutant EMH12 (triangles), and arcA mutant AMJ2 (diamonds) in well-aerated (closed) and poorly-aerated (open) cultures. Poor aeration conditions were created by growing cultures in 250-ml flasks containing 200 ml of SWTO and well-aerated conditions consisted of 50 ml SWTO in 250-ml flasks. (B) Supercoiling of plasmid DNA isolated from ES114 or topA mutants was assayed by electrophoresis through 0.8% agarose containing 20 μg/ml chloroquine. DNA that is less negatively supercoiled prior to loading will migrate more rapidly at this concentration of chloroquine (29). Laddering on the gel is indicative of topoisomers of the same plasmid

that differ by one linking number. The gel was washed in distilled H2O for 2 h, stained in 1 μg/ml ethidium bromide for 1 h, and then imaged.

Discussion

In many bacteria, achieving a high cell density quorum is necessary but not sufficient to elicit full induction of pheromone-dependent behaviors. Therefore, understanding the biological significance of pheromone-mediated regulation in bacteria will require examining the environmentally responsive regulators that govern these systems. Only by elucidating the environmental contingencies required for induction of pheromone-controlled regulons will we be able to fully appreciate their functions in nature and contributions to bacterial fitness. In V. fischeri, environmental conditions clearly play an important role in modulating lux expression, and the ~1000-fold increase in luminescence of V. fischeri upon colonizing E. scolopes cannot be explained by cell density (8, 9). However, in contrast to a detailed understanding of pheromone-mediated activation of luminescence (Fig. 3.1), control of the luxICDABEG operon by environmentally responsive regulators has been less thoroughly explored.

In this study we initiated a systematic examination of the regulatory mechanisms accounting for the dim luminescence of V. fischeri ES114 in culture in order to elucidate the conditions that promote upregulation of luminescence and autoinducer synthesis.

Using transposon mutagenesis, we isolated 28 independent luminescence-up mutants with insertions in 14 loci. Through the characterization of these mutants along with subsequent analyses of ES114 in different media, we have shown that luminescence in

ES114 is responsive to a broad regulatory web influenced by specific environmental conditions. Given that the autoinducer synthase gene luxI is cotranscribed with the genes

responsible for luminescence, these data support the idea that the quorum-sensing circuitry is regulated in response to the environment through multiple regulatory systems.

Environment-dependent lux regulation

The importance of environmental context in luminescence regulation was underscored by simple analyses of the luminescence-up mutants. For example, in all mutants except topA strains, the luminescence-up phenotype was medium-specific (Table

3.1). In some instances, these medium-dependent effects could be rationalized given, the genetic analysis of the mutants. For instance, mutants SLV16, SLV43, and EMH3 displayed a larger luminescence-up effect relative to ES114 when grown in LBS medium as opposed to SWTO (Table 3.1). These mutants have transposon insertions in phoQ, encoding the response regulator of the PhoPQ two-component regulatory system, which responds to environmental signals, including low [Mg2+] (42, 63, 78). Given the much lower levels of Mg+2 in LBS than in SWTO, the medium-dependent phenotype of these mutants was what one would predict if PhoQ repressed luminescence in response to low

Mg2+. This model was tested further and validated by showing a phoQ-dependent repressive effect of Mg2+ on luminescence (Fig. 3.6). Although in other systems PhoPQ also responds to [Ca2+] (78), antimicrobial peptides (4), and pH (5), these did not show clear PhoQ-dependent effects on luminescence in V. fischeri (data not shown). Hussa et al. previously reported multiple homogs of phoP in V. fischeri (38), and the corresponding regulation may be more complex than in E. coli. Interestingly, our data indicate PhoQ-independent effects on luminescence, and taken together Mg2+ appears to influence luminescence through multiple mechanisms.

Similarly, analysis of luminescence-up mutants led to the identification of Pi as a key environmental factor governing luminescence. Mutations in pstA, pstC, or phoU lead to increased luminescence (Table 3.1 and Fig. 3.5), and by analogy to E. coli, mutants in this Pi uptake operon (16, 69) would be expected to display the PhoB-dependent Pi starvation response (81). Consistent with this model, luminescence in ES114 is elevated in response to low Pi levels in a PhoB-dependent manner (Fig. 3.5). Interestingly, the

“low Pi” concentrations utilized in this work appear to be much higher than levels found in the Hawaiian coastal waters inhabited by E. scolopes. Assays completed to determine the [Pi] (14) of seawater showed phosphate levels of <100 nM (data not shown), which is consistent with previous studies (3, 65, 68). However, this does not account for organic phosphates sources, which along with phosphate availability in the host light organ should be considered in future work.

Finally, the identification of luminescence-up mutations in topA, arcA, arcB, and acnB hinted at an important role for redox in luminescence regulation, as each of these loci has been connected to redox-dependent regulation in other systems. The degree of aeration influences regulation of lux expression in V. fischeri (62, 73), and here we show that this is dependent on topA, suggesting a role for supercoiling in this regulatory phenomenon (Fig. 3.7). Ongoing studies are aimed at understanding whether the luminescence-up mutations in arc or acnB reflect other environmental conditions that influence cellular redox state.

Future experiments will also be aimed at understanding whether environmental conditions influence regulation mediated by genes such as ainS, lonA, hns, and tfoY

(Table 3.1). Preliminary work suggests that at least ainS is subject to regulation in

response to the environment, and the other genes may similarly have greater or lesser effects on luminescence in a context-dependent manner.

Mechanisms of luminescence regulation

Understanding mechanism(s) underlying the regulation of luminescence in luminescence-up mutants is important for at least two reasons. First, regulation of the native luxICDABEG promoter is likely to affect LuxI synthase expression and 3-oxo-C6-

HSL synthesis in addition to affecting luminescence, leading to positive feedback and the potential for cell-cell signaling. Most mutations reported here did exert their effects on luminescence through mechanisms dependent on the native lux promoter, and therefore such regulation has the potential for population-wide 3-oxo-C6-HSL-mediated effects.

Secondly, it will be important to ascertain whether regulators act directly on the lux promoter or indirectly, for example, by modulating one of the direct regulators. This distinction will be important for building robust predictive models of luminescence regulation. For example, interdependence of different regulators would imply that multiple environmental conditions must be sensed simultaneously to elicit a regulatory response.

Little is yet known about the specific mechanisms by which the regulators identified here control lux operon expression. It is thought that Lon targets LuxR (51), and we have previously shown that ArcA binds directly to the lux promoter (10), but mechanisms for the other regulators are less clear and await testing. For example, although luxR promoter activity is increased under low Pi conditions, a search of the lux

intergenic region did not reveal a clear pho box typical of a PhoB binding site, suggesting that it may be regulating luminescence indirectly via an unknown mechanism.

For some mutants, the underlying regulatory mechanism is perplexing. Notably, transposon insertions in tRNAMet and tRNAThr produced mutants with an ~6-fold increase in luminescence during growth in SWTO; however, the significance of this finding is unclear. These interrupted tRNAs and those downstream from them are not unique in the

V. fischeri genome. There are several loci for both tRNAMet and tRNAThr annotated within the genome with at least one additional locus having 100% identity to the mutated genes. Thus, although we cannot rule out a model of regulation by rare tRNAs, such a mechanism is not immediately apparent.

Other implications for pheromone-signaling research

Our discovery of regulators that respond to environmental stimuli and control lux expression has important implications. For example, autoinducer-mediated regulation in

V. fischeri has often been used as a model system for mathematically describing a genetic regulatory circuit; however, the effects of environmental regulation on lux expression have been largely overlooked in these studies. Rather, many mathematical models of lux regulatory circuitry either omit environmental regulation entirely (40, 45, 53, 60, 61, 66,

82, 85) or consider only CRP-mediated regulation in response to glucose (7, 15). In the future, components of the environmentally responsive regulators connected to lux should be incorporated into the models of this regulatory circuit.

This work will also direct research aimed at understanding the environment experienced by V. fischeri inside its host, E. scolopes. With a better appreciation of the

conditions and environmentally responsive regulators that lead to dim luminescence in culture, we can now develop clear hypotheses regarding the environmental cues that lead to lux induction during colonization. For example, the studies described above have

2+ prompted greater interest in examining redox, Mg , and Pi levels in the light organ. Our ability to reconstitute and observe this symbiosis offers a unique opportunity to assess the dynamics and regulation of pheromone-mediated signaling in a context that is ecologically relevant for the bacterium.

Acknowledgements

We thank Erica M. Hall, Susan L. Vescovi, and Alecia N. Septer for technical assistance, and Karen Visick for providing strains.

This material is based upon work supported by the National Science Foundation

(NSF) under grants CAREER MCB-0347317 and IOS-0841480, by the Army Research

Office under grant 49549LSII, and by the National Institutes of Health under grant

AI50661 to M. McFall-Ngai. A. N. S. was supported by a National Defense Science and

Engineering Graduate (NDSEG) Fellowship, 32 CFR 168a, under and awarded by DoD,

Air Force Office of Scientific Research. S. L. V. was supported by a NSF Research

Experience for Undergraduates site award (DBI-0453353). Genome information was provided by the Vibrio fischeri Genome Project supported by the W. M. Keck

Foundation.

References

1. Adin, D. M., K. L. Visick, and E. V. Stabb. 2008. Identification of a cellobiose

utilization gene cluster with cryptic beta-galactosidase activity in Vibrio fischeri.

Appl. Environ. Microbiol. 74:4059-4069.

2. Alexeeva, S., K. J. Hellingwerf, and M. J. Teixeira de Mattos. 2003.

Requirement of ArcA for redox regulation in Escherichia coli under microaerobic

but not anaerobic or aerobic conditions. J. Bacteriol. 185:204-209.

3. Armstrong, F. A. J., and H. W. Harvey. 1950. The cycle of phosphorus in the

waters of the English Channel. J. Marine Biol. Assoc. U. K. 29:145-162.

4. Bader, M. W., S. Sanowar, M. E. Daley, A. R. Schneider, U. Cho, W. Xu, R.

E. Klevit, H. Le Moual, and S. I. Miller. 2005. Recognition of antimicrobial

peptides by a bacterial sensor kinase. Cell 122:461-472.

5. Bearson, B. L., L. Wilson, and J. W. Foster. 1998. A low pH-inducible,

PhoPQ-dependent acid tolerance response protects Salmonella typhimurium

against inorganic acid stress. J. Bacteriol. 180:2409-2417.

6. Bebbington, K. J., and H. D. Williams. 2001. A role for DNA supercoiling in

the regulation of the cytochrome bd oxidase of Escherichia coli. Microbiol.

147:591-598.

7. Belta, C., J. Schug, T. Dang, V. Kumar, G. J. Pappas, H. Rubin, and P.

Dunlap. 2001. Stability and reachability analysis of a hybrid model of

luminescence in the marine bacterium Vibrio fischeri. Proceedings of the 40th

IEEE Conference on Decision and Control 2001 1:869-874.

8. Boettcher, K. J., and E. G. Ruby. 1990. Depressed light emission by symbiotic

Vibrio fischeri of the sepiolid squid Euprymna scolopes. J. Bacteriol. 172:3701-

3706.

9. Boettcher, K. J., and E. G. Ruby. 1995. Detection and quantification of Vibrio

fischeri autoinducer from symbiotic squid light organs. J. Bacteriol. 177:1053-

1058.

10. Bose, J. L., U. Kim, W. Bartkowski, R. P. Gunsalus, A. M. Overley, N. L.

Lyell, K. L. Visick, and E. V. Stabb. 2007. Bioluminescence in Vibrio fischeri is

controlled by the redox-responsive regulator ArcA. Mol. Microbiol. 65:538-553.

11. Bose, J. L., C. S. Rosenberg, and E. V. Stabb. 2008. Effects of luxCDABEG

induction in Vibrio fischeri: enhancement of symbiotic colonization and

conditional attenuation of growth in culture. Arch. Microbiol. 190:169-183.

12. Boylan, M., C. Miyamoto, L. Wall, A. Graham, and E. Meighen. 1989. Lux C,

D and E genes of the Vibrio fischeri luminescence operon code for the reductase,

transferase, and synthetase enzymes involved in aldehyde biosynthesis.

Photochem. Photobiol. 49:681-688.

13. Chen, X., S. Schauder, N. Potier, A. Van Dorsselaer, I. Pelczer, B. L. Bassler,

and F. M. Hughson. 2002. Structural identification of a bacterial quorum-sensing

signal containing boron. Nature 415:545-549.

14. Cogan, E. B., G. B. Birrell, and O. H. Griffith. 1999. A robotics-based

automated assay for inorganic and organic phosphates. Anal. Biochem. 271:29-

35.

100

15. Cox, C. D., G. D. Peterson, M. S. Allen, J. M. Lancaster, J. M. McCollum, D.

Austin, L. Yan, G. S. Sayler, and M. L. Simpson. 2003. Analysis of noise in

quorum sensing. OMICS 7:317-334.

16. Cox, G. B., H. Rosenberg, J. A. Downie, and S. Silver. 1981. Genetic analysis

of mutants affected in the Pst inorganic phosphate transport system. J. Bacteriol.

148:1-9.

17. Dorman, C. J., G. C. Barr, N. N. Bhriain, and C. F. Higgins. 1988. DNA

supercoiling and the anaerobic and growth phase regulation of tonB gene

expression. J. Bacteriol. 170:2816-2826.

18. Dunlap, P. V., and E. P. Greenberg. 1985. Control of Vibrio fischeri

luminescence gene expression in Escherichia coli by cyclic AMP and cyclic AMP

receptor protein. J. Bacteriol. 164:45-50.

19. Dunlap, P. V., and E. P. Greenberg. 1988. Control of Vibrio fischeri lux gene

transcription by a cyclic AMP receptor protein-LuxR protein regulatory circuit. J.

Bacteriol. 170:4040-4046.

20. Dunlap, P. V., and J. M. Ray. 1989. Requirement for autoinducer in

transcriptional negative autoregulation of the Vibrio fischeri luxR gene in

Escherichia coli. J. Bacteriol. 171:3549-3552.

21. Dunn, A. K., M. O. Martin, and E. V. Stabb. 2005. Characterization of

pES213, a small mobilizable plasmid from Vibrio fischeri. Plasmid 54:114-134.

22. Dunn, A. K., D. S. Millikan, D. M. Adin, J. L. Bose, and E. V. Stabb. 2006.

New rfp- and pES213-derived tools for analyzing symbiotic Vibrio fischeri reveal

101

patterns of infection and lux expression in situ. Appl. Environ. Microbiol. 72:802-

810.

23. Eberhard, A., A. L. Burlingame, C. Eberhard, G. L. Kenyon, K. H. Nealson,

and N. J. Oppenheimer. 1981. Structural identification of autoinducer of

Photobacterium fischeri luciferase. Biochemistry 20:2444-2449.

24. Engebrecht, J., K. Nealson, and M. Silverman. 1983. Bacterial

bioluminescence: isolation and genetic analysis of functions from Vibrio fischeri.

Cell 32:773-781.

25. Engebrecht, J., and M. Silverman. 1984. Identification of genes and gene

products necessary for bacterial bioluminescence. Proc. Natl. Acad. Sci. U. S. A.

81:4154-4158.

26. Fidopiastis, P. M., C. M. Miyamoto, M. G. Jobling, E. A. Meighen, and E. G.

Ruby. 2002. LitR, a new transcriptional activator in Vibrio fischeri, regulates

luminescence and symbiotic light organ colonization. Mol. Microbiol. 45:131-

143.

27. Friedrich, W. F., and E. P. Greenberg. 1983. Glucose repression of

luminescence and luciferase in Vibrio fischeri. Arch. Microbiol. 134:87-91.

28. Fuqua, C., S. C. Winans, and E. P. Greenberg. 1996. Census and consensus in

bacterial ecosystems: the LuxR-LuxI family of quorum-sensing transcriptional

regulators. Annu. Rev. Microbiol. 50:727-751.

29. Garner, M. M., G. Felsenfeld, M. H. O'Dea, and M. Gellert. 1987. Effects of

DNA supercoiling on the topological properties of nucleosomes. Proc. Natl. Acad.

Sci. U. S. A. 84:2620-2623.

102

30. Gilson, L., A. Kuo, and P. V. Dunlap. 1995. AinS and a new family of

autoinducer synthesis proteins. J. Bacteriol. 177:6946-6951.

31. Gray, K. M., and E. P. Greenberg. 1992. Physical and functional maps of the

luminescence gene cluster in an autoinducer-deficient Vibrio fischeri strain

isolated from a squid light organ. J. Bacteriol. 174:4384-4390.

32. Gray, K. M., and E. P. Greenberg. 1992. Sequencing and analysis of luxR and

luxI, the luminescence regulatory genes from the squid light organ symbiont

Vibrio fischeri ES114. Mol. Mar. Biol. Biotech. 1:414-419.

33. Gruer, M. J., and J. R. Guest. 1994. Two genetically-distinct and differentially-

regulated aconitases (AcnA and AcnB) in Escherichia coli. Microbiol. 140:2531-

2541.

34. Hanahan, D. 1983. Studies on transformation of Escherichia coli with plasmids.

J. Mol. Biol. 166:557-580.

35. Hastings, J. W., and K. H. Nealson. 1977. Bacterial bioluminescence. Annu.

Rev. Microbiol. 31:549-595.

36. Haygood, M. G., and K. H. Nealson. 1985. Mechanisms of iron regulation of

luminescence in Vibrio fischeri. J. Bacteriol. 162:209-216.

37. Hsieh, L. S., R. M. Burger, and K. Drlica. 1991. Bacterial DNA supercoiling

and [ATP]/[ADP]. Changes associated with a transition to anaerobic growth. J.

Mol. Biol. 219:443-450.

38. Hussa, E. A., T. M. O'Shea, C. L. Darnell, E. G. Ruby, and K. L. Visick.

2007. Two-component response regulators of Vibrio fischeri: identification,

mutagenesis, and characterization. J. Bacteriol. 189:5825-5838.

103

39. Jacobs, M. A., A. Alwood, I. Thaipisuttikul, D. Spencer, E. Haugen, S. Ernst,

O. Will, R. Kaul, C. Raymond, R. Levy, L. Chun-Rong, D. Guenthner, D.

Bovee, M. V. Olson, and C. Manoil. 2003. Comprehensive transposon mutant

library of Pseudomonas aeruginosa. Proc. Natl. Acad. Sci. U. S. A. 100:14339-

14344.

40. James, S., P. Nilsson, G. James, S. Kjelleberg, and T. Fagerstrom. 2000.

Luminescence control in the marine bacterium Vibrio fischeri: an analysis of the

dynamics of lux regulation. J. Mol. Biol. 296:1127-1137.

41. Kaplan, H. B., and E. P. Greenberg. 1985. Diffusion of autoinducer is involved

in regulation of the Vibrio fischeri luminescence system. J. Bacteriol. 163:1210-

1214.

42. Kato, A., H. Tanabe, and R. Utsumi. 1999. Molecular characterization of the

PhoP-PhoQ two-component system in Escherichia coli K-12: identification of

extracellular Mg2+-responsive promoters. J. Bacteriol. 181:5516-5520.

43. Kirkegaard, K., and J. C. Wang. 1985. Bacterial DNA topoisomerase I can

relax positively supercoiled DNA containing a single-stranded loop. J. Mol. Biol.

185:625-637.

44. Kuo, A., N. V. Blough, and P. V. Dunlap. 1994. Multiple N-acyl-L-homoserine

lactone autoinducers of luminescence in the marine symbiotic bacterium Vibrio

fischeri. J. Bacteriol. 176:7558-7565.

45. Kuttler, C., and B. A. Hense. 2008. Interplay of two quorum sensing regulation

systems of Vibrio fischeri. J. Theor. Biol. 251:167-180.

104

46. Lupp, C., and E. G. Ruby. 2004. Vibrio fischeri LuxS and AinS: comparative

study of two signal synthases. J. Bacteriol. 186:3873-3881.

47. Lupp, C., M. Urbanowski, E. P. Greenberg, and E. G. Ruby. 2003. The Vibrio

fischeri quorum-sensing systems ain and lux sequentially induce luminescence

gene expression and are important for persistence in the squid host. Mol.

Microbiol. 50:319-331.

48. Lyell, N. L., A. K. Dunn, J. L. Bose, S. L. Vescovi, and E. V. Stabb. 2008.

Effective mutagenesis of Vibrio fischeri by using hyperactive mini-Tn5

derivatives. Appl. Environ. Microbiol. 74:7059-7063.

49. Makino, K., H. Shinagawa, M. Amemura, T. Kawamoto, M. Yamada, and A.

Nakata. 1989. Signal transduction in the phosphate regulon of Escherichia coli

involves phosphotransfer between PhoR and PhoB proteins. J. Mol. Biol.

210:551-559.

50. Makino, K., H. Shinagawa, M. Amemura, S. Kimura, A. Nakata, and A.

Ishihama. 1988. Regulation of the phosphate regulon of Escherichia coli.

Activation of pstS transcription by PhoB protein in vitro. J. Mol. Biol. 203:85-95.

51. Manukhov, I. V., V. Kotova, and G. B. Zavil'genskii. 2006. Role of

GroEL/GroES chaperonin system and Lon protease in regulation of expression

Vibrio fischeri lux genes in Escherichia coli cells. Mol. Biol. (Mosk.) 40:277-283.

52. McCann, J., E. V. Stabb, D. S. Millikan, and E. G. Ruby. 2003. Population

dynamics of Vibrio fischeri during infection of Euprymna scolopes. Appl.

Environ. Microbiol. 69:5928-5934.

105

53. McCollum, J. M., G. D. Peterson, C. D. Cox, M. L. Simpson, and N. F.

Samatova. 2006. The sorting direct method for stochastic simulation of

biochemical systems with varying reaction execution behavior. Comput. Biol.

Chem. 30:39-49.

54. McFall-Ngai, M. J., and E. G. Ruby. 1991. Symbiont recognition and

subsequent morphogenesis as early events in an animal-bacterial mutualism.

Science 254:1491-1494.

55. Mehra, R. K., and W. T. Drabble. 1981. Dual control of the gua operon of

Escherichia coli K12 by adenine and guanine nucleotides. J. Gen. Microbiol.

123:27-37.

56. Meighen, E. A. 1994. Genetics of bacterial bioluminescence. Annu. Rev. Genet.

28:117-139.

57. Miller, J. H. 1992. A short course in bacterial genetics. Cold Spring Harbor

Laboratory Press Plainview, NY.

58. Miyamoto, C. M., Y. H. Lin, and E. A. Meighen. 2000. Control of

bioluminescence in Vibrio fischeri by the LuxO signal response regulator. Mol.

Microbiol. 36:594-607.

59. Muda, M., N. N. Rao, and A. Torriani. 1992. Role of PhoU in phosphate

transport and alkaline phosphatase regulation. J. Bacteriol. 174:8057-8064.

60. Muller, J., C. Kuttler, and B. A. Hense. 2008. Sensitivity of the quorum sensing

system is achieved by low pass filtering. Biosystems 92:76-81.

106

61. Muller, J., C. Kuttler, B. A. Hense, M. Rothballer, and A. Hartmann. 2006.

Cell-cell communication by quorum sensing and dimension-reduction. J. Math.

Biol. 53:672-702.

62. Nealson, K. H., and J. W. Hastings. 1977. Low oxygen is optimal for luciferase

synthesis in some bacteria. Ecological implications. Arch. Microbiol. 112:9-16.

63. Newcombe, J., J. C. Jeynes, E. Mendoza, J. Hinds, G. L. Marsden, R. A.

Stabler, M. Marti, and J. J. McFadden. 2005. Phenotypic and transcriptional

characterization of the meningococcal PhoPQ system, a magnesium-sensing two-

component regulatory system that controls genes involved in remodeling the

meningococcal cell surface. J. Bacteriol. 187:4967-4975.

64. Nijvipakul, S., J. Wongratana, C. Suadee, B. Entsch, D. P. Ballou, and P.

Chaiyen. 2008. LuxG is a functioning flavin reductase for bacterial

luminescence. J. Bacteriol. 190:1531-1538.

65. Redfield, A. C., H. P. Smith, and B. H. Ketchum. 1937. The cycle of organic

phosphorus in the Gulf of Maine. Biol. Bull. 73:421-443.

66. Romero-Campero, F. J., and M. J. Perez-Jimenez. 2008. A model of the

quorum sensing system in Vibrio fischeri using P systems. Artif. Life 14:95-109.

67. Schaefer, A. L., B. L. Hanzelka, A. Eberhard, and E. P. Greenberg. 1996.

Quorum sensing in Vibrio fischeri: probing autoinducer-LuxR interactions with

autoinducer analogs. J. Bacteriol. 178:2897-2901.

68. Solorzano, L., and J. H. Sharp. 1980. Determination of total dissolved

phosphorus and particulate phosphorus in natural waters. Limnol. Oceanogr.

25:754-758.

107

69. Sprague, G. F., Jr., R. M. Bell, and J. E. Cronan, Jr. 1975. A mutant of

Escherichia coli auxotrophic for organic phosphates: evidence for two defects in

inorganic phosphate transport. Mol. Gen. Genet. 143:71-77.

70. Stabb, E. V. (ed.). 2006. The Vibrio fischeri-Euprymna scolopes light organ

symbiosis. ASM Press, Washington, D. C.

71. Stabb, E. V., M. S. Butler, and D. M. Adin. 2004. Correlation between

osmolarity and luminescence of symbiotic Vibrio fischeri strain ES114. J.

Bacteriol. 186:2906-2908.

72. Stabb, E. V., and E. G. Ruby. 2002. RP4-based plasmids for conjugation

between Escherichia coli and members of the Vibrionaceae. Methods Enzymol.

358:413-426.

73. Stabb, E. V., A. Schaefer, J. L. Bose, and E. G. Ruby. 2008. Quorum signaling

and symbiosis in the marine luminous bacterium Vibrio fischeri, p. 233-250,

Chemical communication among microbes. ASM Press, Washington, D.C.

74. Steed, P. M., and B. L. Wanner. 1993. Use of the rep technique for allele

replacement to construct mutants with deletions of the pstSCAB-phoU operon:

evidence of a new role for the PhoU protein in the phosphate regulon. J. Bacteriol.

175:6797-6809.

75. Tiedeman, A. A., and J. M. Smith. 1984. Isolation and characterization of

regulatory mutations affecting the expression of the guaBA operon of Escherichia

coli K-12. Mol. Gen. Genet. 195:77-82.

76. Tu, S. C., and H. I. Mager. 1995. Biochemistry of bacterial bioluminescence.

Photochem. Photobiol. 62:615-624.

108

77. Urbanowski, M. L., C. P. Lostroh, and E. P. Greenberg. 2004. Reversible

acyl-homoserine lactone binding to purified Vibrio fischeri LuxR protein. J.

Bacteriol. 186:631-637.

78. Véscovi, E. G., F. C. Soncini, and E. A. Groisman. 1996. Mg2+ as an

Extracellular Signal: Environmental Regulation of Salmonella Virulence. Cell

84:165-174.

79. Visick, K. L., T. M. O'Shea, A. H. Klein, K. Geszvain, and A. J. Wolfe. 2007.

The sugar phosphotransferase system of Vibrio fischeri inhibits both motility and

bioluminescence. J. Bacteriol. 189:2571-2574.

80. Wang, J. C. 1971. Interaction between DNA and an Escherichia coli protein . J.

Mol. Biol. 55:523-526.

81. Wanner, B. L. 1993. Gene regulation by phosphate in enteric bacteria. J. Cell

Biochem. 51:47-54.

82. Ward, J. P., J. R. King, A. J. Koerber, P. Williams, J. M. Croft, and R. E.

Sockett. 2001. Mathematical modelling of quorum sensing in bacteria. IMA J.

Math Appl. Med. Biol. 18:263-292.

83. Wei, S. L., and R. E. Young. 1989. Development of symbiotic bacterial

bioluminescence in a nearshore cephalopod, Euprymna scolopes. Mar. Biol.

103:541-546.

84. Whistler, C. A., and E. G. Ruby. 2003. GacA regulates symbiotic colonization

traits of Vibrio fischeri and facilitates a beneficial association with an animal host.

J. Bacteriol. 185:7202-7212.

109

85. Williams, J. W., X. Cui, A. Levchenko, and A. M. Stevens. 2008. Robust and

sensitive control of a quorum-sensing circuit by two interlocked feedback loops.

Mol. Syst. Biol. 4:234.

86. Willsky, G. R., and M. H. Malamy. 1980. Characterization of two genetically

separable inorganic phosphate transport systems in Escherichia coli. J. Bacteriol.

144:356-365.

110

CHAPTER 4

CRP REGULATES PHEROMONE-MEDIATED BIOLUMINESCENCE

AT MULTIPLE LEVELS IN VIBRIO FISCHERI ES1141

1Noreen L. Lyell, Deanna M. Colton, Jeffrey L. Bose, Melissa P. Tumen, and Eric V.

Stabb. To be submitted to Journal of Bacteriology.

111

Abstract

Bioluminescence in Vibrio fischeri ES114 is tightly controlled by pheromone- mediated regulation, and this system serves as a model for bacterial cell-cell signaling.

As in other bacteria, pheromone concentration increases with cell density, but pheromone synthesis and perception are also modulated in response to environmental stimuli.

Previous studies, primarily using transgenic Escherichia coli, suggested that expression of the pheromone-dependent bioluminescence activator LuxR is regulated in response to glucose by cAMP-receptor protein (CRP). Consistent with this model, we found that bioluminescence in V. fischeri ES114 is modulated by glucose and stimulated by cAMP.

In addition, a ∆crp mutant was ~100-fold dimmer than ES114 and no longer increased luminescence in response to cAMP, even though cells lacking crp were still metabolically capable of producing luminescence. Using transcriptional reporters and qRT-PCR we found that CRP activates transcription of not only luxR, but also the pheromone synthase gene ainS. DNA binding studies with His-tagged V. fischeri CRP supported bioinformatic predictions of CRP binding sites upstream of both luxR and ainS. Luminescence increased in response to cAMP if either the ainS or luxR systems were under native regulation, suggesting cAMP-CRP significantly increases luminescence through both systems. Finally, using transgenic E. coli we examined expression of a reporter for luxI, which encodes another pheromone synthase, and found

LuxR-independent basal transcription was enhanced by CRP. These results reveal that

CRP is even more integral to the pheromone-mediated regulation of bioluminescence in

V. fischeri ES114 than was previously appreciated.

112

Introduction

The observation that pheromones govern bioluminescence in Vibrio fischeri was a fundamental discovery that shed light on bacterial cell-cell communication. It is now appreciated that diverse bacteria similarly utilize pheromones to regulate numerous processes (43); however, the study of bioluminescence and its regulation in V. fischeri still serves as a powerful model, enabling researchers to explore the mechanisms by which pheromone signals and their regulatory circuits are used by bacteria. In particular, strain ES114 has become a widely used experimental model for V. fischeri, because this isolate‟s bioluminescent symbiosis with the Hawaiian bobtail squid, Euprymna scolopes, can be reconstituted in the laboratory.

The genes responsible for bioluminescence in V. fischeri are separated into two divergent transcripts composed of luxR and luxICDABEG (25, 26, 41). LuxI and LuxR underpin pheromone-mediated regulation of bioluminescence. LuxI encodes a pheromone synthase that generates N-3-oxo-hexanoyl-homoserine lactone (C6-oxo-HSL) also referred to as autoinducer (22). As bacterial population density increases, the membrane permeable C6-oxo-HSL accumulates and at a threshold concentration it binds to the transcriptional regulator LuxR (25, 31, 55). This LuxR-C6-oxo-HSL complex binds the lux box located within the intergenic region between luxR and luxI to activate transcription of luxICDABEG.

Two additional autoinducer pheromones, N-octanoyl-HSL (C8-HSL) generated by AinS and AI-2 generated by LuxS, also regulate bioluminescence (33, 38). C8-HSL and AI-2 are thought to function through distinct receptors that both act via LuxU, LuxO,

Hfq, and the small RNA qrr to increase levels of LitR, which then activates luxR

113

expression (27, 39, 44, 45). C8-HSL can also activate LuxR directly, although it is a weaker activator than 3-oxo-C6-HSL (39).

Although pheromone-mediated regulation such as that controlling bioluminescence is frequently framed in terms of cell-density dependent “quorum- sensing”, environmentally responsive regulators often govern these systems. As a result, a high cell density may be necessary but not sufficient to achieve stimulatory pheromone levels. For example, symbiotic V. fischeri ES114 cells are ~1000-fold brighter and produce more 3-oxo-C6-HSL than cultured cells grown to similar density (3, 4), suggesting that as yet unknown environmental conditions are important in luxICDABEG expression in the squid. Several studies have suggested a role for the environmentally responsive cAMP receptor protein (CRP) in regulating V. fischeri bioluminescence (3,

15-17). CRP was originally identified in Escherichia coli for its role in catabolite repression (24, 60) and has since been shown to activate transcription from numerous promoters (9, 29, 32). When glucose concentrations are low, adenylate cyclase (Cya) activity and intracellular cAMP levels are relatively high, and this effector molecule associates with CRP (8). Together the cAMP-CRP complex binds target DNA and promotes transcription (9, 35).

Though previous research supports a link between catabolite repression and quorum sensing-mediated regulation of luminescence, the exact relationship and mechanism have remained unclear, particularly in V. fischeri ES114. In transgenic E. coli cells carrying the lux operon from strain MJ-1, CRP and cAMP stimulate the induction of luminescence, while the addition of glucose causes a 1-2 hr delay in the onset of luciferase activity (16). Analyses of transcriptional regulators further suggested

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that the cAMP-CRP complex activates transcription of luxR (16, 17) and CRP from E. coli showed binding to DNA upstream of luxR (51). Studies in V. fischeri have suggested possible strain-specific differences. Luminescence in strain MJ-1 is decreased in the presence of glucose (28); however, it was reported that in strain ES114 glucose had no effect (3). In both strains the addition of cAMP increased light production (3, 28).

Furthermore, undefined cya- and crp-like mutants of MJ-1 produce low levels of luminescence, and in the cya-like mutant luminescence was restored by supplying cAMP exogenously (15).

In this study, we examined the regulatory role of CRP in V. fischeri ES114. We found that light production is regulated by CRP and is responsive to addition of both cAMP and glucose. Furthermore, we provide evidence that CRP regulates both pheromone synthesis and the response to pheromone, suggesting a multi-layered integration of carbon metabolism and pheromone-mediated regulation.

Materials and Methods

Bacteria and media

Bacterial strains used in this study are listed in Table 4.1. V. fischeri ES114 was the wild-type strain used throughout (3). Plasmids were transformed into E. coli strains

DH5α (30) or DH5αλpir (19). E. coli was grown in LB (42) or brain heart infusion medium (BHI), and V. fischeri strains were grown in LBS medium (53) or SWTO medium (5). Solid media were prepared with 15 mg ml-1 agar for plating. For selection of E. coli, ampicillin (amp), chloramphenicol (cam), kanamycin (kan), and tetracycline

(tet) were added to LB at final concentrations of 100, 20, 40, and 12 μg ml-1, respectively,

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and 150 μg ml-1 erythromycin (erm) was added to BHI. For selection of V. fischeri on

LBS, cam, erm, and kan were used at concentrations of 2, 5, and 100 μg ml-1, respectively. Where specified, glucose, cyclic adenosine monophosphate (cAMP), and/or

C8-HSL were added to cultures at final concentrations of 20 mM, 10 mM, and 250 nM, respectively.

Table 4.1: Bacterial strains, plasmids, and oligonucleotides used in this study

Strain, plasmid, or Relevant characteristicsa Source or oligonucleotide reference

E. coli

DH5αλpir DH5α lysogenized with λpir (19)

M182 K12 ∆(lacIPOZY) (11)

M182 ∆crp K12 ∆(lacIPOZY), ∆crp (10)

UQ3811 K12 ∆(lacIPOZY), crp::camR, ilv::Tn10 cya (57)

V. fischeri

ES114 Wild-type isolate from E. scolopes (3)

CL53 ES114 ∆luxR::ermR (38)

q DC1 ES114 ∆crp lacI PA1/34-luxCDABEG This study

q JB22 ES114 lacI PA1/34-luxCDABEG (6)

JB24 ES114 ∆crp (5)

NL60 ES114 ∆ainS This study

NL62 ES114 ∆luxR::ermR, ∆ainS This study

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Plasmidsb pCL149 ∆luxR::ermR allele; ColE1, oriTRP4, camR (38) pCR-BluntII-TOPO PCR-product cloning vector; ColE1, kanR Invitrogen pDCRP E. coli crp in pBR322 derivative; ColEI, ampR (56) pDCRP TA158 E. coli crp variant TA158 in pBR322 derivative; (56)

ColEI, ampR pDCRP HL159 E. coli crp variant HL159 in pBR322 derivative; (56)

ColEI, ampR pDCRP HY19 E. coli crp variant HY19 in pBR322 derivative; (49)

ColEI, ampR pDCRP KE101 E. coli crp variant KE101 in pBR322 derivative; (49)

ColEI, ampR pDU9 ∆crp in pBR322 derivative; ColEI, ampR (2) pEVS122 Suicide vector; R6Kγ, oriTRP4, ermR (19) pEXT20 Ptac expression vector; ColEI, ampR (21) pJLB36 PluxR-gfp reporter in pVSV33; pES213, R6Kγ, (5)

oriTRP4, kanR pJLB113 luxR in pCR-BluntII-TOPO; ColE1, kanR This study pJLB117 V. fischeri ∆crp allele; ColE1, R6Kγ, oriTRP4, (5)

camR, kanR pJLB123 Porf1-luxR in pVSV104; pES213, R6Kγ, oriTRP4, (5)

kanR

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pMTV1 Pcon-gfp reporter in pVSV33; pES213, R6Kγ, This study

oriTRP4, kanR pMTV2 PainS-gfp reporter in pVSV33; pES213, R6Kγ, This study

oriTRP4, kanR pNL12 V. fischeri crp in pCR-BluntII-TOPO; ColE1, kanR This study pNL15 Porf1-crp in pVSV105; pES213, R6Kγ, oriTRP4, This study

camR pNL19 Ptac-crp (His6 tag) in pEXT20; ColEI, ampR This study pNL31 ainS upstream fragment in pCR-BluntII-TOPO; (40)

ColE1, kanR pNL53 ainS downstream fragment in pEVS122; R6Kγ, This study

oriTRP4, ermR pNL62 ∆ainS allele; ColEI, R6Kγ, oriV, oriTRP4, kanR, This study

ermR pNL66 129-269 bp of luxR in pCR-BluntII-TOPO; ColE1, This study

kanR pNL73 PluxR-lacZ reporter in pRW50; oriV, tetR This study pNL76 PainS-lacZ reporter in pRW50; oriV, tetR This study pNL81 98-260 bp of lpxA in pCR-BluntII-TOPO; ColE1, This study

kanR pNL82 PluxI-lacZ reporter in pRW50; oriV, tetR This study pNL84 PluxI-lacZ reporter in pRW50; oriV, tetR, luxR This study pNL85 litR in pCR-BluntII-TOPO; ColE1, kanR This study

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pNL86 PainS-lacZ reporter in pRW50; oriV, tetR, litR This study pNL90 91-244 bp of ainS in pCR-BluntII-TOPO; ColE1, This study

kanR pRW50 Promoterless lacZYA; oriV, tetR (37) pVSV33 Promoterless camR-gfp; pES213, R6Kγ, oriTRP4, (20)

kanR pVSV104 Shuttle vector; pES213, R6Kγ, oriTRP4, kanR, (20)

lacZα pVSV105 Shuttle vector; pES213, R6Kγ, oriTRP4, camR, (20)

lacZα

Oligonucleotidesc

DMC2 GGC GGT ACC AGA ACC AAG ACC TGC TCG This study

TGC TAA

JBCRP4 TGC AGG GCA ACG TTG TAC TTG TGC (5)

JBCRP5 GCA TCC TCC AGC AGC CAT TAA GAC C (5)

JBLUXR1 GAA GGA GAT ATA CAT ATG AAC ATT AAA (5)

AAT ATA AAT GC

JBLUXR2 CGC CAA GAT TTT ATG GAA ATG TAT GAG (5)

JBPromoter1 CTA GTT GAC ATG ATA GAA GCA CTC TAC This study

TAT ATT

JBPromoter2 CTA GAA TAT AGT AGA GTG CTT CTA TCA This study

TGT CAA

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Lux1 GGG GTC TAG AGC TTT AGA AAT ACT TT (6)

Lux2 GGA TCC GCT AGG GCG GCC GCC TAA CT (6) pr_MTV1 CGC GGA ATT CAG GAA CTA TAA ACT ATG This study

GTT CTA GGT AAA CCT CAA ACA pr_MTV2d GCG CAA GCT TAT TAA TGG TGA TGG TGA This study

TGG TGA TGA TAA CGA TGA CC GTA AAC

TAC AAT pr_NL54 CGC TAT TAT CTA TCC TCA CTC AAT AAT This study

TAA ACC TGA TG pr_NL55 GAA TGA TGG GAC TTA GAG TAA TCG ACT This study

ACA GGG TC pr_NL58 GGC AGG CCT CAT CAG TTG TTG AAG TAA This study

ATT AAA ATT CTG CG pr_NL78.2 GCG CCC TAG GTG ACT TTT ATA TAA ATG This study

TTA ACT ACT TTA C pr_NL79 CCT CGA CT AAT CGA ATA GAT ATA GAA This study

CTT TTA T pr_NL80 GCG CGA ATT CCA TGA TCA TAA CAA ACT This study

GAT GCA TTA CGG pr_NL81 GCG CAA GCT TGT TCA TTT TTT GTT CAC This study

CTA GCT TAT TGT TA pr_NL86 GCG CGA ATT CAG AAC CAA GAC CTG CTC This study

GTG CTA A

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pr_NL87 GCG CAA GCT TGA TCA GTT GTT GAA GTA This study

AAT TAA AAT TCT GCG pr_NL88 GCA TCC ACG TTT TCT AGG CGG TCG CC 3 This study pr_NL89 AAA TCT AAG GGT TTA CCT TTG TCC GCT This study

CTC pr_NL94 CTG GTA ATG TAA CGA TTG GCG AAG GTA This study

CGG AAG TG pr_NL95 CCG ATA ACA ACT GTT GTT GCT TCA CCA This study

CCA TAT TTC pr_NL96 GCG CGA ATT CCA CCA ATT TGG AGG TTT This study

GGT GAT ATC GC pr_NL97 GCG CAA GCT TTA TCA TTA CAG CCA TGC This study

AAC CTC TCT TAT TT pr_NL100 GCG CCC TAG GGG TTT CCC GAC TGG AAA This study

GCG GGC pr_NL101 GCG CTC TAG AGG TTT CCC GAC TGG AAA This study

GCG GGC pr_NL102 CTG TTG AAT ATA TAA GCA GCC TTA TGT This study

AAA GTT ATT GAG pr_NL103 TCA GCG GTT TTT TAT TAT TAT CAT TTC This study

ATG AAT CCT G pr_NL108 GGC GGA ACG ATT GGA AAT TTG GAA TAC This study

TTA TTT TCA ACA TC

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pr_NL109 CAG TAC TGC ATT TCA AAA GAC AAC CAA This study

AAA CTT TGA TAG CC a. Drug resistance abbreviations used: ampR, ampicillin resistance (bla); camR, chloramphenicol resistance (cat); ermR, erythromycin resistance; kanR, kanamycin resistance (aph); and tetR, tetracycline resistance (tetM). b. All alleles cloned in this study are from V. fischeri strain ES114. Replication origin(s) of each vector are listed as R6Kγ, ColE1, oriV and/or pES213. Plasmids based on pES213 are stable and do not require antibiotic selection for maintenance (Dunn et al., 2005; 2006). c. All oligonucleotides are shown 5‟ to 3‟. d. underlined bases indicate C-terminal His6 tag and newly created stop codon used in protein purification.

Genetic techniques and analyses

Plasmids (Table 4.1) were constructed using standard molecular techniques.

DNA ligase and restriction enzymes were obtained from New England Biolabs (Beverly,

MA). Oligonucleotides used for PCR and cloning (Table 4.1) were synthesized by

Integrated DNA Technologies (Coralville, IA). PCR products were generated using

KOD HiFi DNA Polymerase (Novagen, Madison, WI) or Phusion High-Fidelity DNA

Polymerase (New England Biolabs, Beverly, MA) with an iCycler (BioRad Laboratories,

Hercules, CA). Plasmids used in cloning were purified with the GenElute Plasmid

Miniprep Kit (Sigma-Aldrich, Inc., St. Louis, MO) and DNA from PCR, digestion, and ligation reactions was purified with the DNA Clean and Concentrator-5 Kit (Zymo

Research, Orange, CA). When specified, PCR products were cloned into the pCR-

BluntII-TOPO vector using ZeroBlunt-TOPO PCR Cloning Kit (Invitrogen, Carlsbad,

CA) and white colonies were identified on plates containing 5-bromo-4-chloro-3-indolyl-

β-D-galactoside at a final concentration of 60 μg ml-1. All cloned PCR products were sequenced at the University of Michigan DNA Sequencing Core Facility and sequences were confirmed using Sequencher 4.1.2 (Gene Codes Corp., Ann Arbor, MI).

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To construct a constitutively expressed gfp as a control for promoter-gfp reporters, oligonucleotides JBPromoter1 and JBPromoter2 were designed to anneal together forming an E. coli consensus promoter and single-stranded overhangs compatible with

SpeI-digested DNA. The oligonucleotides were combined in a 1:1 ratio, heated to 100°C and then annealed by allowing the mix to cool slowly at room temperature. This fragment was then ligated into SpeI-digested pVSV33 (20) to generate pMTV1. To construct the PainS-gfp reporter, the 427-bp region upstream of ainS was PCR amplified using primers DMC2 and pr_NL58. The PCR product was digested with KpnI and StuI then ligated into KpnI and StuI-digested pVSV33 (20) to generate pMTV2.

Mutant strain construction

Plasmids carrying mutant alleles were transferred from E. coli into V. fischeri by triparental matings using conjugative helper strain CC118λpir pEVS104 (54).

Recombinational insertion or marker exchange was identified by screening for antibiotic resistance, and putative mutants were tested by PCR. To generate a ∆ainS allele, the

1,833-bp region downstream of ES114 ainS was PCR amplified using primers pr_NL78.2 and pr_NL79. This product was cloned into SmaI-digested pEVS122 (19) to generate pNL53. Plasmid pNL31, which contains the region upstream of ainS (40), and pNL53 were linearized with AvrII and ligated together to generate pNL62, which contains the upstream and downstream sequences fused at the ∆ainS allele, with the ainS start and stop codons separated by the 6-bp AvrII recognition sequence. The ∆ainS allele from pNL62 was crossed into ES114 and CL53 (38) to generate strains NL60 and NL62, respectively, and the deletion of ainS was confirmed by PCR using primers pr_NL88 and

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pr_NL89. To generate DC1, the ∆crp allele on pJLB117 (5) was crossed into the genome of JB22 (6) and allelic exchange was confirmed by PCR and by IPTG inducibility of luminescence in the resulting strain.

Luminescence and fluorescence measurements

Overnight cultures of V. fischeri were diluted 1:1000 in 20 ml or 50 ml of SWTO in 125 ml or 250-ml flasks, respectively, and then incubated with shaking (200 rpm) at

24˚C. At regular intervals, 500-μl samples were removed to measure optical density at

595 nm (OD595) with a BioPhotometer (Brinkman Instruments, Westbury, NY). Relative luminescence was measured with a Glomax TD-20/20 luminometer (Promega, Madison,

WI), immediately following shaking to aerate the sample (6). Specific luminescence was calculated as the luminescence per OD595. Fluorescence expressed from reporter plasmids carrying gfp was measured with a TD-700 fluorometer (Turner Designs,

Sunnyvale, CA) using excitation and emission filters of 486 nm and >510 nm, respectively. Fluorescence of strains carrying the promoterless vector pVSV33 (20) was subtracted as background. Specific fluorescence was calculated as the fluorescence per

OD595, and the reported mean specific fluorescence is for cultures at ~2.0 OD595.

Quantitative PCR and transcript analysis

Overnight cultures of V. fischeri ES114 or JB24 were diluted 1:1000 in 20 ml of

SWTO in 125-ml flasks and then incubated with shaking (200 rpm) at 24˚C. At ~0.5

OD595, 5-ml samples were removed and 1/5 volume 5% acidic phenol and 95% ethanol

(v/v) was added. Samples were incubated on ice for 30 min, and then RNA was isolated

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using the Stratagene Absolutely RNA Miniprep Kit (Agilent Technologies, La Jolla, CA).

Isolated RNA was further treated with Ambion TURBO DNase (Applied Biosystems,

Foster City, CA). cDNA was generated from 1 μg of DNase-treated RNA using

Superscript VILO cDNA Synthesis Kit (Invitrogen, Carlsbad, CA). RT-PCR reactions were completed with iQ SYBR Green Supermix (Biorad, Hercules, CA) and 10 ng cDNA was analyzed using the MyiQ real-time PCR detection system (BioRad, Hercules, CA).

The luxR cDNA was amplified using primers pr_NL54 and pr_NL55, and the ainS cDNA was amplified using primers pr_NL108 and pr_NL109. The lpxA cDNA was amplified using primers pr_NL94 and pr_NL95. All RNA and cDNA concentrations were determined with a Gen5 microplate reader (BioTek, Winooski, VT).

To generate standard curves, the primer sets above (pr_NL54 x pr_NL55, pr_NL108 x pr_NL109, and pr_NL94 x pr_NL95) were used to PCR amplify luxR, ainS, and lpxA, respectively, using ES114 DNA as a template. The blunt luxR, ainS, and lpxA

PCR products were ligated into the pCR-BluntII-TOPO vector to generate pNL66, pNL90, and pNL81, respectively. These plasmids were utilized as standards in real-time

PCR analyses. The efficiency of each primer set was calculated to be ~98%, ~96%, and

~95% using the standard curves generated from serial dilutions of pNL66, pNL90, and pNL89, respectively. To ensure that chromosomal DNA contamination was not affecting the real-time analyses, „no reverse transcriptase‟ control samples were also examined using each primer set and „no template‟ controls were included in each assay.

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Purification of V. fischeri CRP

For CRP purification, V. fischeri crp was PCR amplified using primers JBCRP4 and JBCRP5 and the blunt product was cloned into pCR-BluntII-TOPO to generate pNL12. The wild-type crp allele from pNL12 was then used a template for PCR with primers pr_MTV1 and pr_MTV2. The latter primer mutated the crp stop codon, added a

C-terminal His6 tag, and created a new stop codon (Table 4.1). The PCR product was digested with HindIII and EcoRI then ligated into HindIII- and EcoRI-digested pEXT20 expression vector to generate pNL19.

V. fischeri crp was expressed from pNL19 in E. coli strain UQ3811 (57).

Overnight cultures were diluted 1:100 and grown with shaking (200 rpm) in LB at 37°C to ~0.5 OD595. At this OD595, CRP overexpression was induced by adding isopropyl β-D-

1-thiogalactopyranoside to a final concentration of 1 mM. Following induction, cultures were incubated 5 h, and then cells were pelleted by centrifugation. Cell pellets were resuspended in 50 mM Tris-HCl (pH 8.0), 500 mM KCl, and 1mM EDTA and lysed using a French pressure apparatus. Cell debris was pelleted by centrifugation at 10,000 g for 10 min. The cell lysate was loaded onto a Ni-MAC Cartridge (Novagen, EMD

Chemicals Inc., Gibbstown, NJ), the column was washed with ten volumes 50 mM Tris-

HCl (pH 8.0), 500 mM KCl, and 50 mM imidazole, and then eluted with 50 mM Tris-

HCl (pH 8.0), 500 mM KCl, and 250 mM imidazole in 1 ml fractions. To remove imidazole, elution fractions containing protein were dialyzed overnight in 50 mM Tris-

HCl (pH 8.0), 100 mM KCl, and 1mM EDTA.

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Fluorescence polarization DNA-binding assays

DNA binding assays were completed using 5‟-fluorescein labeled oligonucleotides annealed to unlabeled complement oligonucleotides, both synthesized by Sigma-Aldrich (Saint Louis, MO). Oligonucleotides were resuspended to a final concentration of 1 mM then combined in a 1:1 ratio, boiled for five minutes, and allowed to anneal by cooling slowly at room temperature. Specified concentrations of purified

CRP were added to binding assays to contain a final concentration of 50 mM Tris-HCL pH (8.0), 100 mM KCl, 1 mM EDTA, 15 nM oligo probe, and 20 ng ml-1 salmon sperm

DNA. Binding assay mixtures were incubated at room temperature for 15 min, and then samples were excited at 485 nm and emission measured at 528 nm using a Gen5 microplate reader set to read the extent of fluorescence polarization. Each sample was assayed two additional times, after 25 min and 35 min, and all three measurements were averaged to generate the anisotropy values reported.

Beta-galactosidase transcriptional-reporter assays

Beta-galactosidase transcriptional reporters were constructed using the promoterless lacZ plasmid pRW50 (37). The luxR promoter was amplified with primers pr_NL80 and pr_NL81, the ainS promoter was amplified with primers pr_NL86 and pr_NL87, and the luxI promoter was amplified with primers pr_NL96 and pr_NL97. The

PCR products were digested with EcoRI and HindIII then ligated into EcoRI- and

HindIII-digested pRW50 to generate pNL73, pNL76, and pNL82, respectively.

Because these reporters were used in E. coli, which lacks specific key regulators found in V. fischeri, the complete luxR and litR genes were added to the PluxI-lacZ and

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PainS-lacZ reporter plasmids, respectively. To construct prNL84, luxR was amplified using primers JBLUXR1 and JBLUXR2, and then the blunt PCR product was ligated into the pCR-BluntII-TOPO vector, generating pJLB113. The vector-derived lacZ promoter and luxR insert from pJLB113 were amplified with primers pr_NL100 and pr_NL101, and the PCR product was digested with AvrII and ligated into AvrII-digested pNL82. To construct pNL86, litR was amplified using primers pr_NL102 and pr_NL103, and the blunt PCR product was ligated into the pCR-BluntII-TOPO vector, generating pNL85.

The vector-derived lacZ promoter and litR insert were amplified from pNL85 using primers pr_NL100 and pr_NL101, and the PCR product was digested with AvrII and ligated into AvrII-digested prNL76.

All reporter plasmids were co-transformed into strain M182 ∆crp with either pDCRP, pDCRP-TA158, pDCRP-HL159, pDCRP-HY19, pDCRP-KE101, or pDU9, and the co-transformants were tested in beta-galactosidase assays. Overnight cultures of co- transformants were subcultured 1:100 into 20 ml of LB containing amp and tet in 125-ml flasks and then incubated with shaking (200 rpm) at 28ºC. At OD595 ~0.5 for pNL73, pNL82, and pNL84 assays and OD595 ~1.0 for pNL76 and pNL86 assays, 1-ml samples were removed, pelleted, and stored at -80ºC. For beta-galactosidase assays, pelleted cells were resuspended in 1 ml of Z buffer and 0.5 ml of each resuspension was assayed as previously described (42).

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Results

Luminescence in ES114 is influenced by glucose

To determine the effect of glucose on V. fischeri ES114 luminescence, we examined cultures grown without and with 20 mM glucose (Fig. 4.1A). We found that glucose decreased luminescence when culture OD595 was below 2.0, with a peak effect of

~15-fold. Surprisingly, a brief peak in luminescence was observed in the glucose- supplemented culture at OD595 ~2.3, and was followed by a rapid decline in light production. At this peak, during a brief but reproducible window in cell density, cultures grown with glucose were ~3-fold brighter than cultures grown without glucose (Fig.

4.1A).

1000000 595 100000

10000

1000 ES114ES114

LuminescenceOD / ES114ES114 20mM 20 mM glucoseglucose 100 0 1 2 3 OD B 595 10000000

595 1000000

100000

10000 JB22JB22

1000 JB22JB22 20mM 20 mM glucoseglucose LuminescenceOD / 100 0 1 2 3 OD595

Fig. 4.1: Effect of glucose on luminescence. (A) Specific luminescence for ES114 (wild type) grown in SWTO without (open circles) and with (filled circles) 20 mM glucose. The average of two replicate flasks is shown. (B) Specific luminescence for strain JB22 q (lacI -PA1/34-luxC) grown in SWTO without (open) and with (closed) 20 mM glucose. The average of two replicate flasks is shown.

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The effect of glucose on the luminescence of ES114 in Figure 4.1A might be due to either regulation of genes involved in light production or to metabolic effects caused by growth in glucose. For example, the physiological effects of growing on glucose might alter the availability of substrates for luciferase. To distinguish between these possibilities, we examined the luminescence of mutant JB22, which contains the

q constitutive promoter, lacI -PA1/34, inserted between luxI and luxCDABEG (6). In the

q lacI -PA1/34-luxC background, glucose had no effect on luminescence below an OD595 of

~2.5 (Fig. 4.1B), suggesting that at these culture densities glucose affects ES114 luminescence by controlling gene expression rather than by altering substrate availability.

In both ES114 and JB22, luminescence declines rapidly at higher cell density (Fig. 4.1), which may be due to the build-up of acid in cultures utilizing glucose.

Luminescence in ES114 is affected by cAMP and crp

Previous work using transgenic E. coli carrying the lux genes from V. fischeri strain MJ-1 suggested that cAMP-CRP activated luxR transcription and luminescence

(16, 17). In E. coli, cAMP levels are relatively low in the presence of glucose, which is consistent with less activation of CRP and therefore lower luminescence when glucose is present. The exogenous addition of 4 mM cAMP to lux-containing E. coli stimulated luminescence (16), presumably because enough cAMP entered into cells to activate CRP.

Similarly, it has been reported that addition of 10 mM cAMP stimulates luminescence in

V. fischeri cultures (3, 18). We likewise found that addition of 10 mM cAMP to ES114 stimulated luminescence ~10-fold (Fig. 4.2A). It was difficult to interpret the effects of glucose on luminescence in a V. fischeri ∆crp mutant, because this strain grew very

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poorly without glucose; however, we could test the effect of cAMP addition on both

ES114 and the ∆crp mutant in media with added glucose. In glucose-containing media we found that the ∆crp mutant was both dimmer than wild type and unresponsive to cAMP (Fig. 4.2B).

A 10000000

595 1000000

100000

10000 ES114

1000 ES114 10mM10 mM cAMPcAMP LuminescenceOD / 100 0 1 2 3

OD595 B ES114ES114 glucose ES114ES114 both 10 mMcAMP

10000000 JB24JB24 glucose JB24JB24 both 10 mM cAMP

1000000 595

100000

10000

1000

LuminescenceOD / 100 0 1 2 3 OD595 C ES114 JB24 10000000 JB22 595 1000000 DC1

100000

10000

1000 LuminescenceOD / 100 0 1 2 3 OD595

Fig. 4.2: Effect of cAMP on luminescence. (A) Specific luminescence of ES114 (wild type) grown in SWTO without (open circles) and with (filled circles) 10 mM cAMP added. The average of two replicate flasks is shown. (B) Specific luminescence of ES114 and ∆crp mutant JB24 grown in SWTO containing 20 mM glucose without (open circles) or with (filled circles) 10 mM cAMP. The average of two replicate flasks is

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q shown. (C) Specific luminescence ES114, JB22 (lacI -PA1/34-luxC), ∆crp mutant JB24, q and ∆crp lacI -PA1/34-luxC mutant DC1 grown with 20 mM glucose. The average of two replicate flasks is shown.

We considered the possibility that pleiotropic effects associated with the ∆crp mutation might underlie the low luminescence of this mutant, and we therefore tested the effect of the ∆crp allele in the JB22 strain background, with lux expressed from constitutive non-native PA1/34 promoter. We found that deleting crp led to a ~15-fold decrease in luminescence in the constitutive-lux background (Fig. 4.2C); however, this does not account for the ~1000-fold decrease in luminescence seen when crp is deleted in

ES114 (Fig. 4.2C). Furthermore, the ∆crp constitutive-lux strain DC1 is ~100-fold brighter than the ∆crp mutant JB24, suggesting the attenuation of luminescence in the latter does not simply reflect a metabolic limitation caused by deleting crp.

CRP influences luxR and ainS transcription

To identify genes involved in luminescence that may be controlled by cAMP-

CRP, we used Virtual Footprint (46) to analyze the V. fischeri ES114 genome (data not shown). As expected, a putative CRP binding site was identified upstream of luxR. More surprisingly, a potential CRP binding site was also identified upstream of ainS, which encodes the autoinducer synthase responsible for generating C8-HSL (33). In fact, the putative CRP binding site upstream of ainS was a better match to the weighted consensus

CRP binding site than the site in the lux intergenic region.

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1000 ES114 100

595 JB24

Fluorescence / OD Fluorescence / 0.1

0.01

PPconcon-gfp PPluxRluxR-gfp PPainSainS-gfp

Fig. 4.3: CRP regulation of luxR and ainS promoter activity. (A) Specific fluorescence generated from the reporters Pcon-gfp (pMTV1), PluxR-gfp (pJLB36) and PainS-gfp (pMTV2) carried within ES114 (grey bars) or crp mutant JB24 (black bars). Fluorescence from each strain harboring the promoterless vector, pVSV33, was subtracted as background. Data represent the average specific fluorescence when the culture OD595 was ~2.0. Averages and standard errors were calculated from replicate flasks of each examined strain.

The effect of CRP on the expression of the luxR and ainS promoters was examined with the PluxR-gfp and PainS-gfp promoter-reporters. Fluorescence expressed from each reporter was measured in wild type and the ∆crp mutant (Fig. 4.3). The PluxR- gfp and PainS-gfp reporters yielded a statistically significant (p ≤ 0.05) ~15-fold and ~10- fold decrease fluorescence in the ∆crp mutant, respectively. A constitutively active promoter-reporter with a consensus promoter driving gfp expression was also examined in the wild type and crp mutant to ensure that the ∆crp allele did not decrease fluorescence of all gfp reporters. Fluorescence of the Pcon-gfp reporter was actually higher in the ∆crp mutant, suggesting that, if anything, the effects of crp of the PluxR- and

PainS-gfp reporters may underestimate the effects of crp on transcription of these genes.

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A 1000000 luxR

100000 cDNA 10000 1000 * 100 10

Transcript / 10ng / Transcript 1 ES114 JB24 B 10000 ainS

cDNA 1000 *

100

Transcript / 10ng / Transcript 1 ES114 JB24 C 1000000 lpxA

100000 cDNA 10000 1000 100 10

Transcript / 10ng / Transcript 1 ES114 JB24

Fig. 4.4: RT-PCR transcript analysis of luxR and ainS. Copy number of (A) luxR, (B) ainS, and (C) lpxA transcripts in 10 ng total cDNA generated from ES114 (wild-type) or crp mutant JB24 RNA. Bars indicate standard error (n=3). * Indicates a significant difference in transcript number between wild type and crp mutant transcript copy numbers as determined by a Student‟s T-test (p≤0.0005).

To directly measure whether CRP controls the transcript levels of luxR and ainS, we utilized quantitative real-time PCR to determine the copy number of both luxR and ainS transcripts in V. fischeri ES114 and in the ∆crp mutant (Figs. 4.4A and B). The copy number of the luxR and ainS transcripts was decreased by ~250-fold and ~7-fold,

134

respectively. To test whether all transcripts appear decreased in the ∆crp mutant, we also examined the lpxA transcript as a control. LpxA is involved in lipid A biosynthesis in

Gram-negative bacteria, and is an essential housekeeping gene (13) that is not part of the

CRP regulon in E. coli (58). The transcript level of lpxA was equivalent in both the wild type and the ∆crp mutant (Fig. 4.4C). These results support the model that CRP regulates luxR and ainS expression.

CRP binds upstream of luxR and ainS

Previous work showed that E. coli CRP can bind upstream of luxR from V. fischeri strain ATCC7744 (51), and we sought to test whether this would also be true for

CRP from V. fischeri binding the ES114 lux promoter, and whether binding upstream of ainS could also be detected. Using purified V. fischeri CRP, we tested in vitro binding to

5‟ fluorescein labeled DNA fragments (Fig. 4.5A). The melR target was previously shown to associate with CRP, and we used it in this study as a positive control (Fig.

4.5A). The melR shuffled target contains the same nucleotides as the melR target, but they were randomized, thereby serving as a negative binding control. With increased anisotropy serving as a measure of target binding, we observed cAMP-dependent binding of CRP to the melR target and no binding to the melR shuffled target (Figs. 4.5B and C).

We next tested if CRP binds the predicted CRP target sequences upstream of luxR and ainS. As with melR, we observed a cAMP-dependent increase in anisotropy as CRP was added with both the luxR and ainS targets, indicating that CRP binds sequences found upstream of these genes (Figs. 4.5D and E). This supports the model that cAMP-

CRP binds near the promoters of luxR and ainS.

135

A Target Oligonucleotide target sequence

melR Flc-GTAAATGTGATGTACATCACATGGAT melR shuffled Flc-GTAAAGAGACGCTTTAAGTGCTATAT luxR Flc-GTAAATTCGATCTGGGTCACATTTAT ainS Flc-GTAATTATGAAAAACTTCACACTTAT

Consensus AATGTGATCTAGATCACATTT B C 130 melR target 130 melR shuffled target

120 120

110 110

Anisotropy Anisotropy

100 100 0 1000 2000 3000 4000 5000 0 1000 2000 3000 4000 5000 [CRP] Monomer (nM) [CRP] Monomer (nM) D E 150 luxR target 130 ainS target 140 120 130 120

110 Anisotropy Anisotropy 110 100 100 0 1000 2000 3000 4000 5000 0 1000 2000 3000 4000 5000 [CRP] Monomer (nM) [CRP] Monomer (nM)

Fig. 4.5: Fluorescence polarization analysis of CRP binding to the luxR and ainS promoter regions. (A) Fluorescein labeled oligonucleotides utilized for binding analyses. * Indicates position of fluorescein modification at the 5‟ end of the oligo target sequences. The melR sequence was previously shown to bind CRP (57) and is used in this study as a positive control. The melR shuffled sequence was generated using melR sequence and the shuffle DNA application from The Sequence Manipulation Suite (http://www.bioinformatics.org/sms/). The CRP binding sequences upstream of luxR and ainS were identified using the regulon analysis application of Virtual Footprint (http://prodoric.tu-bs.de/vfp/). The V. fischeri ES114 genome was analyzed using the CRP-specific position weight matrix from E. coli K12. Underlined region of the melR target is the previously identified CRP binding site and underlined regions of the luxR and ainS targets are the binding sequences identified by Virtual Footprint. Fluorescence polarization curves for (B) melR oligo target, (C) shuffled melR oligo target, (D) luxR oligo target, and (E) ainS oligo target measured without (open) and with (closed) 1 mM cAMP. All curve data points represent the average of two independent replicate samples that were each measured three times.

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Regulation of both luxR and ainS may contribute to cAMP activation of luminescence

Previous research suggested that the cAMP-mediated induction of luminescence is due to cAMP-CRP promoting transcription of luxR (17), however, this work was done in E. coli lacking the ainS autoinducer synthase. To test whether regulation of the luxR promoter alone could account for the effect of cAMP-CRP on luminescence in V. fischeri we determined the effect of cAMP on luminescence of a luxR mutant (CL53) (38) complemented by luxR constitutively expressed in trans (pJLB123) (5) (Fig. 4.6). Using this construct eliminated regulation of luxR from the CRP-regulated native promoter and maintained constitutive expression of luxR (20). When 10 mM cAMP was added to

CL53 pJLB123, luminescence increased ~15-fold (Fig. 4.6). Thus, the cAMP-mediated induction of luminescence is not solely due to CRP regulation of luxR.

We also examined the luminescence of an ainS mutant supplemented with C8-

HSL (NL60) without and with 10 mM cAMP (Fig. 4.6). With constant C8-HSL, the ainS mutant‟s luminescence increased ~5-fold when 10 mM cAMP was added (Fig. 4.6).

Thus, as with luxR above, the cAMP-mediated induction of luminescence appears to not be solely due to CRP regulation of ainS either. We then examined the effect of cAMP on a luxR ainS double mutant (NL62) complemented with pJLB123 and supplemented with

C8-HSL. In this strain, neither C8-HSL or LuxR levels should be subject to direct regulatory control by cAMP-CRP, and a cAMP-mediated increase in luminescence was not observed (Fig. 4.6). Taken together, these results suggest that cAMP-CRP control of both ainS and luxR is significant in the luminescence of V. fischeri ES114.

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No cAMP 10 mMcAMP 100000000

* 595 10000000

1000000 *

100000 LuminescenceOD /

10000 JB22JB22 CL53,CL53 123 NL60,NL60 C8 NL62,NL62 123 PA1/34-luxC pJLB123 C8-HSL pJLB123 luxR::ermR ∆ainS C8C8-HSL pluxR luxR::ermR, ∆ainS pluxR

Fig. 4.6: Both ainS and luxR regulation contribute to the effect of cAMP on luminescence. Maximal specific luminescence is shown for ES114, JB22, a luxR mutant complemented with a constitutive luxR in trans (CL53 pJLB123), an ainS mutant biochemically complemented with 250 mM C8-HSL (NL60 C8-HSL), and luxR ainS double mutant complemented with luxR in trans and 250 mM C8-HSL (NL62 pJLB123 C8-HSL). Cultures were grown in SWTO without (grey bars) and with (black bars) 10 mM cAMP added. Bars indicate standard error (n=2). * Indicates a significant difference between cultures grown without and with cAMP as determined by a Student‟s T-test (p≤0.05).

CRP activates transcription of luxR, luxI, and ainS promoters through different mechanisms in E. coli

CRP usually regulates transcription by recruiting RNA polymerase to promoters following binding to a target sequence (9). CRP-dependent promoters are grouped into three classes: Class I-type promoters require one point of contact with RNAP at activating region I (ARI); Class II-type require two points of contact between CRP and

RNAP at both activating region I and activating region II (ARI and ARII); and Class III- type promoters either require activating regions I or II if they contact RNAP, or at some

Class III-type promoters CRP exerts an effect on a second regulatory protein through

138

DNA binding or protein-protein interaction, in which case ARI and ARII are not required

(9, 23, 47, 59).

A P -lacZ 1000 luxR

100

10 Miller UnitsMiller * * * 1 pNL73 pNL73 pNL73 pNL73 pNL73 pNL73 pDCRP pDU9 TA158 HL159 HY19 KE101 B P -lacZ, litR 10 ainS

* Miller UnitsMiller

1 pNL86 pNL86 pNL86 pNL86 pNL86 pNL86 pDCRP pDU9 TA158 HL159 HY19 KE101 C P -lacZ 10 ainS * *

* Miller UnitsMiller

1 pNL76 pNL76 pNL76 pNL76 pNL76 pNL76 pDCRP pDU9 TA158 HL159 HY19 KE101

D P -lacZ 100 luxI *

10 * * Miller UnitsMiller

1 pNL82 pNL82 pNL82 pNL82 pNL82 pNL82 pDCRP pDU9 TA158 HL159 HY19 KE101

Fig. 4.7: Effects of ARI and ARII crp mutations on luxR, ainS and luxI promoter activity in E. coli. Miller units generated from the reporters (A) PluxR-lacZ (pNL73), (B) PainS- lacZ (pNL76), (C) PainS-lacZ, litR (pNL76), and (C) PluxI-lacZ (pNL82). Bars indicate

139

standard error (n=2). * Indicates a significant difference between pDCRP and empty vector pDU9 or AR variants as determined by a Student‟s T-test (p≤0.05).

V. fischeri promoters were examined using a reporter system developed in E. coli wherein known ARI and ARII crp mutations are used to classify CRP-regulated promoters (2, 49, 56). In this dual plasmid system, promoter-lacZ reporters are generated in pRW50 and crp alleles are expressed from a compatible plasmid (37). Plasmid pDCRP contains the wild-type E. coli crp and is used as a positive control, whereas plasmid pDU9 lacks crp and is used as a negative control. The crp alleles on pDCRP-

TA158 and pDCRP-HL159 alleles are mutated at ARI and the crp alleles pDCRP-HY19 and pDCRP-KE101 alleles are mutated at ARII. To ensure that E. coli CRP activates transcription from V. fischeri promoters, we tested whether the wild-type E. coli crp would complement the growth and luminescence defect of the V. fischeri crp mutant.

When the E. coli wild-type crp allele was expressed in trans, luminescence and growth in the absence of glucose were restored in the V. fischeri crp mutant (data not shown).

Expression from the PluxR-lacZ (pNL73) promoter-reporter increased ~100-fold in cells co-transformed with the wild-type CRP allele relative to co-transformation with the empty vector control (Fig. 4.7A). This increase in activity is also evident in cells co- transformed with ARII variant alleles but not in cells co-transformed with ARI variant alleles. This indicates that the effect of CRP on luxR expression requires only ARI and therefore suggests a CRP Class I-type promoter.

We next attempted to classify the ainS promoter using a PainS-lacZ reporter.

Because previous research suggests a role for LitR in ainS expression (39), litR was cloned into the PainS-lacZ reporter plasmid. In the presence of litR, the wild-type CRP

140

allele increased beta-galactosidase activity from the PainS-lacZ reporter ~9-fold relative to co-transformation with the empty vector control (Fig. 4.7B). This increase in activity was also observed in cells co-transformed with either the ARI or ARII variant alleles.

Because the ARI and ARII variants did not alter expression from the PainS-lacZ reporter, our results suggest that this enhancement of expression is due a Class III-type promoter interaction.

We also examined expression of the ainS reporter in the absence of litR (Fig.

4.7C). These experiments confirmed that litR contributes to expression of PainS-lacZ, because there was less LacZ activity in the construct lacking litR (Figs. 4.7B and C).

Surprisingly, when litR was absent we also observed an increase in PainS-lacZ activity in the absence of CRP or in the presence of CRP with ARI mutant alleles (Fig. 4.7C).

These results suggest an additional layer of regulation, at least in E. coli, whereby CRP can mediate negative regulation of ainS through a Class I-type promoter, perhaps by activating a repressor of ainS.

Finally, we used this reporter system to test the effect of CRP on the luxI promoter. The CRP binding site upstream of luxR is likewise upstream of the divergently transcribed luxI. Previous work, also in transgenic E. coli, suggested that in the absence of luxR CRP has a negative effect on expression of the luxI promoter from V. fischeri strain MJ-1 (16). We found that our PluxI-lacZ reporter derived from strain ES114 increased ~3-fold in expression in the presence of the wild-type crp or with the ARII mutants (Fig. 4.7D), suggesting that CRP activates LuxR-independent basal expression of luxI through a mechanism that requires ARI. We also added luxR to the PluxI-lacZ reporter plasmid under control of a non-native constitutive promoter, and observed the

141

expected autoinducer-dependent increase in PluxI-lacZ expression (data not shown); however, when the 3-oxo-C6-HSL autoinducer combined with LuxR to enhance transcription, no further increase was seen when crp was present (data not shown).

Discussion

V. fischeri ES114 regulates luminescence using a pheromone-mediated regulatory circuitry similar to many other bacterial systems in that signaling is both cell-density dependent and regulated in response to the environment. Although high cell density may be necessary for lux induction in ES114, it is not sufficient to achieve stimulatory pheromone levels. Symbiotic V. fischeri ES114 cells are ~1000-fold brighter and produce more 3-oxo-C6-HSL than cultured cells at equivalent densities (3), illustrating that environmental conditions are also important in luxICDABEG expression. Previous reports have suggested a role for glucose and CRP-mediated regulation of V. fischeri bioluminescence (3, 15-17, 51). These previous studies used V. fischeri strain MJ-1 (15), the lux genes cloned from MJ-1 into E. coli (16, 17), undefined spontaneous mutants of

MJ-1 (15), or purified E. coli CRP protein in conjunction with lux DNA from V. fischeri

ATCC7744 (51).

Our goal in this study was to test and define the role of glucose and CRP in lux regulation in V. fischeri ES114. Recently, ES114 has become a popular experimental model, and the genetic tools developed for this strain allowed us to explore this area in ways that were not possible when the effects of glucose and CRP on lux were first examined. It seemed especially prudent not to assume that previous conclusions about

CRP‟s role in lux regulation held in ES114, because the lux system in ES114 has

142

diverged significantly from lux in strains like MJ-1 (7), it has been reported that ES114

does not respond to glucose as MJ-1 does (3), and it is now understood than key

regulators of lux would be absent in transgenic E. coli. Our results underscore this last

point, as we found that CRP regulates the ainS pheromone synthase system, which was

absent in studies with transgenic E. coli. However, we also generated support for the

previous conclusion that CRP regulates luxR in ES114, much as it does in transgenic E.

coli. Both of these phenomena appear significant in luminescence regulation in ES114.

Taken together our data agree with previous studies that cAMP-CRP enhances

luminescence in V. fischeri, but the regulatory mechanisms appear more complex.

Regulation by CRP or similar proteins is integral to pheromone circuitry in diverse

systems

Interestingly, the role of CRP in modulating the pheromone signaling circuitry of

V. fischeri apparently illustrates a widespread trend. Several bacteria with mechanistically

diverse pheromone signaling systems integrate CRP, CRP-like proteins, or catabolite

repression into regulation of their pheromone signaling systems. For example, Vibrio

cholerae and Vibrio harveyi lack the V. fischeri LuxI/LuxR system, but the quorum-

sensing master regulator HapR in V. cholerae and its homolog in V. harveyi are also

regulated by CRP (12, 36, 52). V. cholerae mutants defective in CRP express lower

levels of hapR resulting in increased production of cholera toxin and enhanced biofilm

(36, 52). Similarly, CRP indirectly regulates the pheromone synthase luxS in E. coli (14),

and CRP activates transcription of the acyl homoserine lactone-dependent regulators

encoded by lasR and expR in Pseudomonas aeruginosa and Erwinia chrysanthemi,

143

respectively (1, 48). Furthermore, the Agr quorum sensing circuitry, which regulates virulence and antibiotic resistance in Staphylococcus aureus, is modulated by carbon- catabolite protein A (CcpA) (50). Taken together these results suggest that pheromone levels in many systems integrate both carbon source availability and cell density.

Role of glucose in luminescence regulation

The regulatory role of glucose in V. fischeri has been difficult to interpret due in part to potential strain-specific effects. Previous research showed luminescence in strain

MJ-1 is decreased when glucose is added (28), yet glucose was reported to have no effect in ES114 (3). We found that glucose modulates luminescence in V. fischeri ES114, and using a constitutive lux construct this appears to be a regulatory and not simply metabolic phenomenon. When glucose is added to cultures of ES114, luminescence is decreased compared to cultures without glucose at low cell density, but at OD595 ~2.3 luminescence peaks briefly, with cultures actually getting more luminescent in the presence of glucose

(Fig. 4.1A). Thus, depending on when a culture is sampled, and perhaps on other growth conditions, one might potentially see no apparent effect of glucose on ES114.

The initial decrease in luminescence upon addition of glucose is easy to explain by lower CRP-mediated ainS and luxR expression, discussed below. In ES114, at low to moderate cell densities, luminescence expression is primarily driven by LuxR combined with the AinS product C8-HSL (39). As cell density increases, C8-HSL competes with the stronger activator 3-oxo-C6-HSL for binding with LuxR, such that C8-HSL becomes effectively an inhibitor of luminescence (39, 40), as was seen in MJ-1 (33, 34). We speculate that in ES114 grown with glucose, cAMP production is inhibited, and there is

144

less active cAMP-CRP to promote expression of ainS and luxR, leaving cells dimmer.

Because of less C8-HSL production however, at higher cell densities the stronger activator 3-oxo-C6-HSL may elicit the sharper increase in luminescence due to a lack of competition from C8-HSL.

CRP and regulation of luxR

It was previously suggested that the cAMP-CRP complex promotes luminescence by increasing transcription of luxR (16, 17). This is supported by research that showed

CRP from E. coli binds upstream of luxR (51). Our data support the model that cAMP-

CRP directly regulates luxR (Figs. 4.3, 4 and 5). It should be noted that although the intergenic lux sequence has diverged significantly between ES114 and MJ-1, the CRP binding site in this region is largely conserved (7). In addition, using a transgenic E. coli system, we found that the effect of CRP on luxR requires only the CRP activating region

I, indicating this effect is mediated by a class I-type promoter interaction. We were surprised to find that when luxR expression was artificially held constant, addition of cAMP still influenced luminescence (Fig. 4.6) leading us to investigate other components of the V. fischeri pheromone signaling system that might be controlled by CRP.

CRP and regulation of ainS

Bioinformatic analysis of the V. fischeri ES114 genome revealed a potential CRP binding site upstream of ainS that returned a better score than the site upstream of luxR.

We then found experimental evidence that CRP regulates ainS (Figs. 4.3, 4 and 5). In addition, we show that LitR is important for CRP-dependent ainS expression (Fig. 4.7B),

145

and in fact this effect appears independent of CRP activating regions I or II, suggesting a possible class III-type promoter with CRP influencing LitR rather than RNAP.

Moreover, at least in E. coli, an additional regulator controlled by a CRP class I-type promoter apparently influences ainS expression (Fig. 4.7C). In the future, improved methods for assaying C8-HSL from V. fischeri cultures would allow us to better test how such regulatory mechanisms affect the ultimate accumulation of this pheromone.

CRP and regulation of luxI

Interestingly, we also provide data suggesting that CRP may directly regulate transcription of luxI. In previous work with transgenic E. coli and the MJ-1 lux genes,

CRP appeared to inhibit luxI transcription in the absence of LuxR (17), however in our transgenic E. coli system using an ES114-derived lux promoter, we observed the opposite effect (Fig. 4.7D). The spacing of the CRP binding site to the luxI promoter differs by one nucleotide between strains ES114 and MJ-1 (7). It is not clear whether any direct regulation of luxI by CRP is relevant in V. fischeri, where the effects of CRP on luxR and ainS may overpower any direct modulation of the luxICDABEG promoter. If cAMP-

CRP has a significant direct influence on expression from the luxI promoter it would probably be at very low cell densities where a lack of pheromone accumulation renders

LuxR inactive and effects on ainS expression irrelevant. In the future, studies focused at low cell density may reveal if CRP plays a role in modulating the basal LuxR- independent expression of the luxICDABEG operon.

146

The role of cAMP in luminescence regulation

The addition of cAMP to cultures of ES114 resulted in a ~10-fold increase in luminescence (Fig. 4.2A), and this effect was dependent on crp (Fig. 4.2B). We hypothesize that exogenous cAMP led to greater intracellular cAMP and more cAMP-

CRP-mediated activation of luxR and ainS. We further speculate that the effect of glucose on luminescence may similarly reflect intracellular cAMP levels. As an intracellular signal, cAMP levels will probably be much lower than what we add to cultures, and future work should be directed at measuring intracellular cAMP levels and how they are affected by carbon sources or other growth conditions.

Studying cAMP-CRP in the ES114 model

We have found that cAMP-CRP regulates luminescence in V. fischeri ES114, and that the mechanisms underlying this regulation are more complex than what was previously understood. A new model for this CRP-mediated regulation of luminescence is presented in Figure 4.8, although we cannot rule out the possibility that CRP is integrated with the pheromone-signaling system of V. fischeri ES114 at other levels as well. Moreover, it remains to be determined exactly how environmental conditions are connected to cAMP levels in this bacterium. By laying the groundwork in strain ES114, it should be possible to explore these areas and examine the relevance of cAMP-CRP- mediated regulation in a natural symbiotic infection.

147

glucose cAMP LitR

litR CRP † X

luxR luxICDABEG ainS LuxR C6 C8 C8-LuxR

C6-LuxR

Fig. 4.8: Proposed model of CRP-mediated regulation in V. fischeri luminescence. Regulation of luxI and potential negative regulator of ainS (designated „X‟) is suggested only by data from transgenic E.coli and is therefore shown with dashed lines. The autoinducers 3-oxo-C6-HSL and C8-HSL are abbreviated as C6 and C8, respectively. † C8-HSL functions through a phosphorelay involving AinR, LuxU, LuxO, Hfq, and the small RNA qrr that increases levels of LitR to activate luxR expression (27, 45). C8- HSL can also activate LuxR directly, although it is a weaker activator than 3-oxo-C6- HSL (39) and this is represented by the relative weight of the arrows.

148

Acknowledgements

We thank Anne Weeks and Alecia N. Septer for technical assistance and Stephen

J. W. Busby for E. coli strains M182 and M182∆crp, the pRW50 promoterless lacZα vector, and all pDCRP crp constructs utilized in the beta-galactosidase promoter-reporter experiments. We also thank Gary P. Roberts for the pEXT20 expression vector, and E. coli strain UQ3811 utilized in the purification of CRP.

This research was supported by the National Science Foundation (NSF) under grants CAREER MCB-0347317 and OCE-0929081.

149

References

1. Albus, A. M., E. C. Pesci, L. J. Runyen-Janecky, S. E. West, and B. H.

Iglewski. 1997. Vfr controls quorum sensing in Pseudomonas aeruginosa. J.

Bacteriol. 179:3928-3935.

2. Bell, A., K. Gaston, R. Williams, K. Chapman, A. Kolb, H. Buc, S. Minchin,

J. Williams, and S. Busby. 1990. Mutations that alter the ability of the

Escherichia coli cyclic AMP receptor protein to activate transcription. Nucleic

Acids Res. 18:7243-7250.

3. Boettcher, K. J., and E. G. Ruby. 1990. Depressed light emission by symbiotic

Vibrio fischeri of the sepiolid squid Euprymna scolopes. J. Bacteriol. 172:3701-

3706.

4. Boettcher, K. J., and E. G. Ruby. 1995. Detection and quantification of Vibrio

fischeri autoinducer from symbiotic squid light organs. J. Bacteriol. 177:1053-

1058.

5. Bose, J. L., U. Kim, W. Bartkowski, R. P. Gunsalus, A. M. Overley, N. L.

Lyell, K. L. Visick, and E. V. Stabb. 2007. Bioluminescence in Vibrio fischeri is

controlled by the redox-responsive regulator ArcA. Mol. Microbiol. 65:538-553.

6. Bose, J. L., C. S. Rosenberg, and E. V. Stabb. 2008. Effects of luxCDABEG

induction in Vibrio fischeri: enhancement of symbiotic colonization and

conditional attenuation of growth in culture. Arch. Microbiol. 190:169-183.

7. Bose, J. L., M. S. Wollenberg, D. M. Colton, M. J. Mandel, A. N. Septer, A.

K. Dunn, and E. V. Stabb. 2011. Contribution of rapid evolution of the luxR-luxI

150

intergenic region to the diverse bioluminescence outputs of Vibrio fischeri strains

isolated from different environments. Appl. Environ. Microbiol. 77:2445-2457.

8. Botsford, J. L., and J. G. Harman. 1992. Cyclic AMP in prokaryotes.

Microbiol. Rev. 56:100-122.

9. Busby, S., and R. H. Ebright. 1999. Transcription activation by catabolite

activator protein (CAP). J. Mol. Biol. 293:199-213.

10. Busby, S., D. Kotlarz, and H. Buc. 1983. Deletion mutagenesis of the

Escherichia coli galactose operon promoter region. J. Mol. Biol. 167:259-274.

11. Casadaban, M. J., and S. N. Cohen. 1980. Analysis of gene control signals by

DNA fusion and cloning in Escherichia coli. J. Mol. Biol. 138:179-207.

12. Chatterjee, J., C. M. Miyamoto, A. Zouzoulas, B. F. Lang, N. Skouris, and E.

A. Meighen. 2002. MetR and CRP bind to the Vibrio harveyi lux promoters and

regulate luminescence. Mol. Microbiol. 46:101-111.

13. Crowell, D. N., M. S. Anderson, and C. R. Raetz. 1986. Molecular cloning of

the genes for lipid A disaccharide synthase and UDP-N-acetylglucosamine

acyltransferase in Escherichia coli. J. Bacteriol. 168:152-159.

14. De Lay, N., and S. Gottesman. 2009. The Crp-activated small noncoding

regulatory RNA CyaR (RyeE) links nutritional status to group behavior. J.

Bacteriol. 191:461-476.

15. Dunlap, P. V. 1989. Regulation of luminescence by cyclic AMP in cya-like and

crp-like mutants of Vibrio fischeri. J. Bacteriol. 171:1199-1202.

151

16. Dunlap, P. V., and E. P. Greenberg. 1985. Control of Vibrio fischeri

luminescence gene expression in Escherichia coli by cyclic AMP and cyclic AMP

receptor protein. J. Bacteriol. 164:45-50.

17. Dunlap, P. V., and E. P. Greenberg. 1988. Control of Vibrio fischeri lux gene

transcription by a cyclic AMP receptor protein-LuxR protein regulatory circuit. J.

Bacteriol. 170:4040-4046.

18. Dunlap, P. V., and J. M. Ray. 1989. Requirement for autoinducer in

transcriptional negative autoregulation of the Vibrio fischeri luxR gene in

Escherichia coli. J. Bacteriol. 171:3549-3552.

19. Dunn, A. K., M. O. Martin, and E. V. Stabb. 2005. Characterization of

pES213, a small mobilizable plasmid from Vibrio fischeri. Plasmid 54:114-134.

20. Dunn, A. K., D. S. Millikan, D. M. Adin, J. L. Bose, and E. V. Stabb. 2006.

New rfp- and pES213-derived tools for analyzing symbiotic Vibrio fischeri reveal

patterns of infection and lux expression in situ. Appl. Environ. Microbiol. 72:802-

810.

21. Dykxhoorn, D. M., R. St Pierre, and T. Linn. 1996. A set of compatible tac

promoter expression vectors. Gene 177:133-136.

22. Eberhard, A., A. L. Burlingame, C. Eberhard, G. L. Kenyon, K. H. Nealson,

and N. J. Oppenheimer. 1981. Structural identification of autoinducer of

Photobacterium fischeri luciferase. Biochemistry 20:2444-2449.

23. Ebright, R. H. 1993. Transcription activation at Class I CAP-dependent

promoters. Mol. Microbiol. 8:797-802.

152

24. Emmer, M., B. deCrombrugghe, I. Pastan, and R. Perlman. 1970. Cyclic

AMP receptor protein of E. coli: its role in the synthesis of inducible enzymes.

Proc. Natl. Acad. Sci. U. S. A. 66:480-487.

25. Engebrecht, J., K. Nealson, and M. Silverman. 1983. Bacterial

bioluminescence: isolation and genetic analysis of functions from Vibrio fischeri.

Cell 32:773-781.

26. Engebrecht, J., and M. Silverman. 1984. Identification of genes and gene

products necessary for bacterial bioluminescence. Proc. Natl. Acad. Sci. U. S. A.

81:4154-4158.

27. Fidopiastis, P. M., C. M. Miyamoto, M. G. Jobling, E. A. Meighen, and E. G.

Ruby. 2002. LitR, a new transcriptional activator in Vibrio fischeri, regulates

luminescence and symbiotic light organ colonization. Mol. Microbiol. 45:131-

143.

28. Friedrich, W. F., and E. P. Greenberg. 1983. Glucose repression of

luminescence and luciferase in Vibrio fischeri. Arch. Microbiol. 134:87-91.

29. Grainger, D. C., D. Hurd, M. Harrison, J. Holdstock, and S. J. Busby. 2005.

Studies of the distribution of Escherichia coli cAMP-receptor protein and RNA

polymerase along the E. coli chromosome. Proc. Natl. Acad. Sci. U. S. A.

102:17693-17698.

30. Hanahan, D. 1983. Studies on transformation of Escherichia coli with plasmids.

J. Mol. Biol. 166:557-580.

153

31. Kaplan, H. B., and E. P. Greenberg. 1985. Diffusion of autoinducer is involved

in regulation of the Vibrio fischeri luminescence system. J. Bacteriol. 163:1210-

1214.

32. Kolb, A., S. Busby, H. Buc, S. Garges, and S. Adhya. 1993. Transcriptional

regulation by cAMP and its receptor protein. Annu. Rev. Biochem. 62:749-795.

33. Kuo, A., N. V. Blough, and P. V. Dunlap. 1994. Multiple N-acyl-L-homoserine

lactone autoinducers of luminescence in the marine symbiotic bacterium Vibrio

fischeri. J. Bacteriol. 176:7558-7565.

34. Kuo, A., S. M. Callahan, and P. V. Dunlap. 1996. Modulation of luminescence

operon expression by N-octanoyl-L-homoserine lactone in ainS mutants of Vibrio

fischeri. J. Bacteriol. 178:971-976.

35. Lawson, C. L., D. Swigon, K. S. Murakami, S. A. Darst, H. M. Berman, and

R. H. Ebright. 2004. Catabolite activator protein: DNA binding and transcription

activation. Curr. Opin. Struct. Biol. 14:10-20.

36. Liang, W., A. Pascual-Montano, A. J. Silva, and J. A. Benitez. 2007. The

cyclic AMP receptor protein modulates quorum sensing, motility and multiple

genes that affect intestinal colonization in Vibrio cholerae. Microbiol. 153:2964-

2975.

37. Lodge, J., J. Fear, S. Busby, P. Gunasekaran, and N. R. Kamini. 1992. Broad

host range plasmids carrying the Escherichia coli lactose and galactose operons.

FEMS Microbiol. Lett. 74:271-276.

38. Lupp, C., and E. G. Ruby. 2004. Vibrio fischeri LuxS and AinS: comparative

study of two signal synthases. J. Bacteriol. 186:3873-3881.

154

39. Lupp, C., M. Urbanowski, E. P. Greenberg, and E. G. Ruby. 2003. The Vibrio

fischeri quorum-sensing systems ain and lux sequentially induce luminescence

gene expression and are important for persistence in the squid host. Mol.

Microbiol. 50:319-331.

40. Lyell, N. L., A. K. Dunn, J. L. Bose, and E. V. Stabb. 2010. Bright mutants of

Vibrio fischeri ES114 reveal conditions and regulators that control

bioluminescence and expression of the lux operon. J. Bacteriol. 192:5103-5114.

41. Meighen, E. A. 1994. Genetics of bacterial bioluminescence. Annu. Rev. Genet.

28:117-139.

42. Miller, J. H. 1992. A short course in bacterial genetics. Cold Spring Harbor

Laboratory Press Plainview, NY.

43. Miller, M. B., and B. L. Bassler. 2001. Quorum sensing in bacteria. Annu. Rev.

Microbiol. 55:165-199.

44. Miyamoto, C. M., Y. H. Lin, and E. A. Meighen. 2000. Control of

bioluminescence in Vibrio fischeri by the LuxO signal response regulator. Mol.

Microbiol. 36:594-607.

45. Miyashiro, T., M. S. Wollenberg, X. Cao, D. Oehlert, and E. G. Ruby. 2010.

A single qrr gene is necessary and sufficient for LuxO-mediated regulation in

Vibrio fischeri. Mol. Microbiol. 77:1556-1567.

46. Munch, R., K. Hiller, A. Grote, M. Scheer, J. Klein, M. Schobert, and D.

Jahn. 2005. Virtual Footprint and PRODORIC: an integrative framework for

regulon prediction in prokaryotes. Bioinformatics 21:4187-4189.

155

47. Niu, W., Y. Kim, G. Tau, T. Heyduk, and R. H. Ebright. 1996. Transcription

activation at class II CAP-dependent promoters: two interactions between CAP

and RNA polymerase. Cell 87:1123-1134.

48. Reverchon, S., M. L. Bouillant, G. Salmond, and W. Nasser. 1998. Integration

of the quorum-sensing system in the regulatory networks controlling virulence

factor synthesis in Erwinia chrysanthemi. Mol. Microbiol. 29:1407-1418.

49. Rhodius, V. A., D. M. West, C. L. Webster, S. J. Busby, and N. J. Savery.

1997. Transcription activation at class II CRP-dependent promoters: the role of

different activating regions. Nucleic Acids Res. 25:326-332.

50. Seidl, K., M. Stucki, M. Ruegg, C. Goerke, C. Wolz, L. Harris, B. Berger-

Bachi, and M. Bischoff. 2006. Staphylococcus aureus CcpA affects virulence

determinant production and antibiotic resistance. Antimicrob. Agents Chemother.

50:1183-1194.

51. Shadel, G. S., J. H. Devine, and T. O. Baldwin. 1990. Control of the lux regulon

of Vibrio fischeri. J. Biolumin. Chemilumin. 5:99-106.

52. Silva, A. J., and J. A. Benitez. 2004. Transcriptional regulation of Vibrio

cholerae hemagglutinin/protease by the cyclic AMP receptor protein and RpoS. J.

Bacteriol. 186:6374-6382.

53. Stabb, E. V., K. A. Reich, and E. G. Ruby. 2001. Vibrio fischeri genes hvnA

and hvnB encode secreted NAD(+)-glycohydrolases. J. Bacteriol. 183:309-317.

54. Stabb, E. V., and E. G. Ruby. 2002. RP4-based plasmids for conjugation

between Escherichia coli and members of the Vibrionaceae. Methods Enzymol.

358:413-426.

156

55. Urbanowski, M. L., C. P. Lostroh, and E. P. Greenberg. 2004. Reversible

acyl-homoserine lactone binding to purified Vibrio fischeri LuxR protein. J.

Bacteriol. 186:631-637.

56. West, D., R. Williams, V. Rhodius, A. Bell, N. Sharma, C. Zou, N. Fujita, A.

Ishihama, and S. Busby. 1993. Interactions between the Escherichia coli cyclic

AMP receptor protein and RNA polymerase at class II promoters. Mol. Microbiol.

10:789-797.

57. Youn, H., R. L. Kerby, M. Conrad, and G. P. Roberts. 2006. Study of highly

constitutively active mutants suggests how cAMP activates cAMP receptor

protein. J. Biol. Chem. 281:1119-1127.

58. Zheng, D., C. Constantinidou, J. L. Hobman, and S. D. Minchin. 2004.

Identification of the CRP regulon using in vitro and in vivo transcriptional

profiling. Nucleic Acids Res. 32:5874-5893.

59. Zhou, Y., X. Zhang, and R. H. Ebright. 1993. Identification of the activating

region of catabolite gene activator protein (CAP): isolation and characterization of

mutants of CAP specifically defective in transcription activation. Proc. Natl.

Acad. Sci. U. S. A. 90:6081-6085.

60. Zubay, G., D. Schwartz, and J. Beckwith. 1970. Mechanism of activation of

catabolite-sensitive genes: a positive control system. Proc. Natl. Acad. Sci. U. S.

A. 66:104-110.

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CHAPTER 5

CONCLUSIONS AND FUTURE DIRECTIONS

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The purpose of this dissertation was to examine how regulators that respond to environmental conditions govern the pheromone-mediated regulatory circuitry in V. fischeri ES114. In this chapter, I will discuss how my research has furthered the appreciation for environmental regulation in the V. fischeri lux system, and I will suggest future directions for research in this area.

In Chapter 2, I discussed the development of a mini-Tn5 transposon mutagenesis system that allows large-scale mutant screening in V. fischeri. This transposon mutagenesis system was utilized to generate the luminescence-up mutants characterized in Chapter 3 and in Appendix A. Twenty-eight mutants were identified with disruptions in fourteen loci that resulted in brighter than wild-type luminescence. These effects on luminescence were partly or entirely dependent on expression of the lux genes from the native luxI promoter. Disruptions in acnB, topA, pstA, pstC, specific tRNA‟s, tfoY, and guaB identified novel regulators of V. fischeri luminescence whereas insertions in arcA, arcB, lonA, and hns confirmed previously identified negative regulators. In the case of lonA and hns, studies completed in E. coli were validated.

For certain novel regulators, I completed experiments to determine the environmental conditions mediating lux regulation, leading to the discovery that [Pi] and

[Mg2+] influence light production (Figs. 3.5 and 3.6). Also, experiments comparing wild type and the topA mutant under different aeration regimes suggested a role for supercoiling in redox-dependent regulation of lux.

In a surprising result, the ainS mutant was isolated in the luminescence-up screen despite ainS mutants being dimmer than wild type in broth cultures. By determining the luminescence phenotypes of mutants lacking components of the C8-HSL-mediated

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signaling pathway, I showed that the importance of competition between 3-oxo-HSL and

C8-HSL for LuxR binding depends on growth context in V. fischeri ES114 luminescence.

Specifically, in broth cultures C8-HSL acts primarily as an inducer of luminescence whereas in growth on plates C8-HSL plays a greater role in blocking 3-oxo-C6-HSL- mediated induction of LuxR.

Taken together, these mutants revealed a complex regulatory web that governs the lux system and also provided insight into environmental conditions that mediate light production in V. fischeri ES114.

In Chapter 4, I expanded upon previous research that suggested exogenous cAMP or glucose can influence luminescence through CRP-mediated regulation. I provide evidence that CRP is more integral to the pheromone-mediated regulation of luminescence than was previously proposed. Specifically, I showed that CRP activates transcription of both luxR and ainS (Figs. 4.3 and 4.4). By examining the association between purified CRP and the putative CRP binding sites upstream of luxR and ainS I confirmed that CRP binds within the promoter regions of these genes, which presumably activates their transcription. Other experiments suggest a possible role for CRP in the direct regulation of the luxI promoter, although this conclusion is less clear and requires further examination.

Elucidating the regulatory web governing the pheromone-dependent lux system

Though the research discussed in Chapter 3 identified regulators and conditions that promote upregulation of luminescence and autoinducer synthesis, little is yet understood about the specific mechanisms these regulators use to control light

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production. For example, the pstA/C and phoQ mutants led to the discovery that luminescence is affected by Pi in a manner dependent on PhoB, but it remains to be seen whether or not PhoB directly regulates lux. Though Pi limitation promoted an increase in

PluxR-gfp expression (Fig. 3.5B), a search of the lux intergenic region did not identify an apparent pho box characteristic of a PhoB binding site. In future studies, footprinting assays of the intergenic region could identify an atypical pho box or confirm the absence of the PhoB binding site. If the intergenic region lacks a pho box then bioinformatic analyses of the Pho regulon may provide further insight into the regulatory mechanism leading to the increase in luminescence evident at low Pi concentrations.

I also showed that low Mg2+ mediated a decrease in luminescence (Fig. 3.6A), and that this decrease in luminescence appears to be PhoQ dependent; however, an unexpected increase in light production also occurs in the phoQ mutant when grown in low [Mg2+]. This could suggest that an unknown Mg2+-responsive regulator promotes luminescence at low [Mg2+], whereas PhoQ inhibits luminescence in these conditions.

Microarray analyses of V. fischeri ES114 grown in low Mg2+ may provide clues as to the unknown regulators that mediate lux expression. Future work examining the currently unidentified Mg2+-responsive regulators would expand what is currently known concerning the environmental cues involved in lux regulation and provide a more complete model of how luminescence is controlled in response to environmental stimuli.

2+ Interestingly, I found that [Pi] and [Mg ] affect luminescence in various V. fischeri strains (data not shown), suggesting that these environmental signals are part of a conserved regulatory web governing bioluminescence, and potentially pheromone- mediated circuitry, in organisms other than ES114.

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Previous research has shown that the degree of aeration influences luminescence in V. fischeri (6), and in Chapter 3 I show that this is dependent on topA. In poorly aerated cultures, ES114 luminescence decreases at OD595 ~0.5, whereas luminescence in the topA mutant is unaffected (Fig. 3.8A). Because topA mutants have more highly supercoiled DNA than wild type (Fig. 3.8B), these data support a model in which DNA topology is important in lux regulation.

The role of aeration in luminescence is important, because studies suggest the environment within the light organ is microaerobic (1, 3). In experiments not reported in

Chapter 3, I showed that negative supercoiling is increased and that PtopA-gfp expression is decreased in V. fischeri ES114 when cells are grown anaerobically (data not shown).

Given these data, I hypothesize that topA-mediated relaxation of negative supercoiling may be important in the induction of luminescence observed when cells colonize E. scolopes. To further examine the DNA topology of symbiotic cells, I developed a method for extracting V. fischeri from the host tissue, which is discussed in Appendix B.

Future research using this method could provide insight into whether DNA topology influences luminescence during colonization, and it could also give further insight into the oxygen levels within the light organ.

In addition to further exploring the mechanisms employed by the regulators identified in Chapter 3, the transposon mutagenesis system discussed in Chapter 2 can be utilized to identify positive regulators of luminescence. Because the screen used in this dissertation identified bright mutants, only negative regulators were discovered. To isolate mutants containing disruptions in positive regulators, a bright strain of V. fischeri could be mutagenized and mutants screened for a „luminescence-down‟ phenotype.

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Alternatively, mutants in the dim wild-type background could be screened using quantitative and sensitive measurements in a microplate-format luminometer. Conditions such as low Pi that increase luminescence of ES114 would enhance the likelihood of identifying luminescence-down mutants. Such experiments would draw an even more complete picture of the regulatory web that controls bioluminescence in V. fischeri.

Elucidating the role of CRP in the regulation of luminescence

In Chapter 4, I provide evidence that CRP regulates both luxR and ainS, demonstrating that the role of CRP in V. fischeri ES114 luminescence is more complex than previously appreciated. The role of CRP was examined by determining the expression of luxR and ainS. Because AinS is the autoinducer synthase that generates

C8-HSL, analysis of ainS expression is not an exact measure of autoinducer concentration. A small increase in ainS expression at one point in growth might reflect a much larger change in C8-HSL accumulation after long-term growth in batch culture. To better examine the effect of CRP on autoinducer concentration, a C8-HSL bioassay is necessary. A tool for measuring C8-HSL concentration would not only aid in determining the effect of CRP on C8-HSL, but would also provide insight into the competition that exists between 3-oxo-C6-HSL and C8-HSL for LuxR binding. In

Chapter 3 I show that an ainS mutant is brighter than the wild-type parent due to the absence of C8-HSL competition (Table 3.2). Finally, a C8-HSL bioassay would be instrumental in examining other C8-HSL-responsive pheromone-mediated circuitry.

Unfortunately, the only available C8-HSL bioassay (4) has proved difficult and experimentally incompatible with V. fischeri culture supernatents.

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Consistent with the model that cAMP-CRP leads to activation of lux expression, I found that high exogenous cAMP concentrations can activate CRP-mediated regulation of luminescence in V. fischeri ES114. When 10 mM cAMP is added to cultures, luminescence increases 10-fold (Fig. 4.2A). In these experiments an excess of cAMP is added, which is consistent with previous methodologies in other organisms. To determine the concentration of cAMP necessary to activate CRP and induce luminescence, future work should be aimed at determining the intracellular concentration of cAMP in V. fischeri. Also, because CRP mutants have a colonization defect (Deanna

M. Colton, personal communication), measuring the intracellular levels of cAMP may help define the role of CRP in symbiotic cells. Analysis of genes upregulated in response to the host environment suggests that chitin is an available carbon source (7). CRP is important in the expression of genes necessary for chitin metabolism in E. coli (5), and if this is true in V. fischeri the inability of crp mutants to fully colonize and persist within the E. scolopes light organ could be at least in part because these cells are unable to utilize the available chitin.

In addition to finding that CRP controls ainS expression, I also provide evidence that CRP-dependent regulation of ainS requires LitR. In E. coli cells carrying litR in trans, the PainS-lacZ reporter expression increased ~5-fold (Fig. 4.7B). In the absence of

LitR, CRP did not activate the ainS promoter (Fig. 4.7C). This effect of LitR may be explained by the presence of a putative LitR binding site upstream of ainS (Rahul

Kulkarni, personal communication). This putative LitR binding site is positioned 6 bp upstream from the CRP binding site confirmed in Chapter 4 (Fig. 4.5E). At Class III- type promoters, CRP can interact directly with second activators through protein-protein

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interactions. In this mechanism, the activating region I and activating region II residues have little to no role in the protein-protein interaction (2). Therefore, the relationship between CRP and LitR in the activation of ainS is further supported by the PainS-lacZ reporter studies that show wild-type crp levels of expression when activating region variant crp alleles were tested (Fig. 4.7B). To examine the potential co-regulation of ainS by CRP and LitR, co-immunoprecipitation experiments using the His6-tagged CRP generated in Chapter 4 could determine if these proteins associate with one another.

Furthermore, amino acid substitutions could be made to determine which residues are important in the CRP-LitR association.

An additional layer of regulation suggested by the PainS-lacZ reporter analysis is the potential existence of a negative regulator of ainS that itself is positively regulated by

CRP (Fig. 4.7C). Because this assay was completed in E. coli it is possible that this regulation is not relevant to ainS in V. fischeri; however, experiments to test this model are important in developing a more complete understanding of CRP-mediated regulation of ainS. Experiments are also necessary to confirm that the ainS promoter is Class III- type.

The lacZ promoter-reporter assay utilized in Chapter 4 to assess the residues important in CRP-mediated activation of luxR and ainS does not conclusively define these promoters as Class I, II, or III-type. Rather, these experiments examine which activating regions associate with RNAP and, therefore, are important in promoting transcription. Though the activating regions suggest the mechanism of CRP regulation at these promoters, the location of the CRP binding site relative to the transcriptional start site is a more definitive method used to classify CRP regulated promoters. Class I-type

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promoters have only one point of contact with CRP at the α subunit C-terminal domain, and therefore can be located upstream of the transcriptional start site on the same DNA face, whereas in Class II-type promoters, the CRP binding site overlaps the -35 site.

Class III-type promoters are more complex in that spacing from the transcriptional start site is only relevant if CRP contacts RNA polymerase (2). Future studies identifying the transcriptional start sites of luxR and ainS would provide insight into the mechanism of

CRP regulation at these promoters. Also, mapping these transcript start sites may inform research examining the interactions between other regulators that bind within the luxR and ainS promoter regions. For example, binding sites for both LitR and FNR have been identified upstream of luxR and, as discussed above, a putative LitR binding site was identified upstream of ainS.

Conclusion

Overall, in this dissertation I have shown that environmental conditions and specific regulators are integrally associated with the pheromone-mediated regulatory circuitry in V. fischeri ES114. I have presented work that identified and characterized novel regulators in addition to work that further defined the role of CRP in V. fischeri

ES114 luminescence. As this research is carried forward, additional unknown regulators and better defined mechanisms utilized by the regulators identified here are sure to greatly expand our knowledge regarding the importance of cell-cell communication in bacteria.

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References

1. Boettcher, K. J., and E. G. Ruby. 1995. Detection and quantification of Vibrio

fischeri autoinducer from symbiotic squid light organs. J. Bacteriol. 177:1053-

1058.

2. Busby, S., and R. H. Ebright. 1999. Transcription activation by catabolite

activator protein (CAP). J. Mol. Biol. 293:199-213.

3. Dunn, A. K., and E. V. Stabb. 2008. Genetic analysis of trimethylamine N-oxide

reductases in the light organ symbiont Vibrio fischeri ES114. J. Bacteriol.

190:5814-5823.

4. Flavier, A. B., L. M. Ganova-Raeva, M. A. Schell, and T. P. Denny. 1997.

Hierarchical autoinduction in Ralstonia solanacearum: control of acyl-

homoserine lactone production by a novel autoregulatory system responsive to 3-

hydroxypalmitic acid methyl ester. J. Bacteriol. 179:7089-7097.

5. Plumbridge, J., and O. Pellegrini. 2004. Expression of the chitobiose operon of

Escherichia coli is regulated by three transcription factors: NagC, ChbR and

CAP. Mol. Microbiol. 52:437-449.

6. Stabb, E. V., A. Schaefer, J. L. Bose, and E. G. Ruby. 2008. Quorum signaling

and symbiosis in the marine luminous bacterium Vibrio fischeri, p. 233-250,

Chemical communication among microbes. ASM Press, Washington, D.C.

7. Wier, A. M., S. V. Nyholm, M. J. Mandel, R. P. Massengo-Tiasse, A. L.

Schaefer, I. Koroleva, S. Splinter-Bondurant, B. Brown, L. Manzella, E. Snir,

H. Almabrazi, T. E. Scheetz, F. Bonaldo Mde, T. L. Casavant, M. B. Soares,

J. E. Cronan, J. L. Reed, E. G. Ruby, and M. J. McFall-Ngai. 2010.

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Transcriptional patterns in both host and bacterium underlie a daily rhythm of anatomical and metabolic change in a beneficial symbiosis. Proc. Natl. Acad. Sci.

U. S. A. 107:2259-2264.

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APPENDIX A

COLONIZATION AND MOTILITY PHENOTYPES

OF VIBRIO FISCHERI ES114 LUMINESCENCE-UP MUTANTS

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Introduction

Vibrio fischeri is a model organism studied for its pheromone-mediated regulation, its bioluminescence, and its association with the Hawaiian bobtail squid

Euprymna scolopes. V. fischeri strain ES114 is widely used, because its symbiosis with

E. scolopes can reconstituted in the laboratory (10, 14). Furthermore, the onset of quorum sensing mediated luminescence can be measured in both symbiotic cells and in cells cultured outside of the host.

The ability of V. fischeri to generate light is critical to the symbiosis suggesting that bioluminescence and the interaction with E. scolopes are intimately related (2, 12).

In colonization studies, wild-type ES114 cells reach population levels 3-4-fold higher than non-luminescent mutants (2). One hypothesis to explain this phenomenon is that the inability of dark mutants to persist within the host is due to the role of luminescence in alleviating oxidative stress (2, 12). In this model, the luciferase-driven light reaction lowers oxygen concentrations as an antioxidant defense mechanism (1).

In addition, studies have shown that some mutants with aberrant luminescence phenotypes also have altered motility, suggesting that the two are co-regulated (6, 15).

Motility is an important colonization factor that affects both the initiation of the symbiosis and the accumulation of V. fischeri cells within the E. scolopes host (6-8).

Furthermore, flagellation may be important in cells‟ ability to reach preferred sites within the E. scolopes light organ (13). Because the regulatory networks of luminescence and motility can overlap and because alterations in motility can result in colonization defects, we also examined the motility phenotypes of the twenty-eight luminescence-up mutants described in Chapter 3.

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Materials and Methods

Bacterial strains and media

All strains, media, and growth conditions are as described in Chapter 3.

Colonization of E. scolopes

Aposymbiotic E. scolopes hatchlings were inoculated with V. fischeri by placing them into 20-ml vials with 5 ml of Instant Ocean (Aquarium Systems, Mentor, Ohio) containing the bacterial strain of interest following a previously described procedure (9,

11). Briefly, to generate inocula, cultures were grown to an OD595 of between 0.4 and 0.7 in 5 ml of SWTO within 50 ml conical tubes at 28˚C without shaking. Cultures were diluted to ~2000 CFU ml-1 in Instant Ocean and hatchling squid were placed in these inocula for 12 to 14 hours before being rinsed in V. fischeri-free Instant Ocean. Although inocula ranged from 1000 to 3000 CFU ml-1, within each experiment, squid were exposed to similar concentrations of mutant or wild-type cells in the respective inocula. Infected squid were homogenized at 24 or 48 h post-inoculation, then serially diluted and plated onto LBS. Plates were incubated overnight at 28˚C and the colonies were counted to calculate CFU per squid. Relative luminescence was calculated as luminescence per

CFU with the luminescence of each individual squid measured using a LS 6500 Multi

Purpose Scintillation Counter (Beckman Coulter, Fullerton, Calif.).

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Motility Assays

Overnight cultures of V. fischeri strains were diluted 1:1000 in 3 ml of LBS in 18 mm tubes and then incubated at 28˚C with shaking until reaching an OD595 of ~0.5. 10-

μl samples were then spotted onto soft agar plates that contained, per liter of total volume, 5 g of tryptone, 3 g of yeast extract, 3 ml of glycerol, 700 ml of Instant Ocean mixed to 36 ppt, and 2.5 mg ml-1 agar. The swimming rate was determined by measuring the diameter (mm) of the visible spot of cells as the culture spread across the agar.

Results and Discussion

Symbiotic competence of luminescence-up mutants

The ability to colonize the squid host is enhanced by luminescence of V. fischeri

(2, 12), and we analyzed each luminescence-up mutant strain individually for its ability to colonize E. scolopes. Though all of the mutants were able to colonize the juvenile squid, certain mutants showed symbiotic phenotypes that deviated from wild-type, and these are discussed further in experiments reported below (Fig. A.1A-D).

Mutants with disruptions in pstA and pstC showed a severe delay in the onset of symbiotic luminescence relative to ES114 (Fig. A.1A), however, the mutants colonized at levels equivalent to ES114 by 48 h post-inoculation (data not shown). The topA mutants also had a delay in colonization (data not shown), and these mutants achieved colonization levels ~4-fold lower than that of ES114 at 48 h post-inoculation (Fig. A.1B).

Though mutants with disruptions in phoQ and guaB also colonized E. scolopes ~2-fold less than ES114 by 48 h post-inoculation (Figs. A.1C and D), these mutants were not delayed in the onset of symbiotic luminescence (data not shown). Mutants with

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insertions in hns showed the most severe colonization defect with colonization levels

(CFUs) ~10-fold less than ES114 at 48 h post-inoculation (Fig. A.1D) while also providing essentially no detectable symbiotic luminescence (data not shown).

Fig. A.1: Effects of transposon mutations on symbiosis. (A) Onset of symbiotic luminescence in aposymbiotic squid (white squares) and those colonized by strains ES114 (wild type) (black squares), SLV10 (pstA::mini-Tn5-ermR) (grey circles), and SLV30 (pstC::mini-Tn5-ermR) (grey triangles). (B - D) Ability of mutant strains to individually colonize the squid host relative to ES114 48 h post-inoculation. Error bars represent standard error for n≥9 animals. * Student‟s t-test indicates a significant difference between wild type and mutant colonization (p≤0.05). Data shown is from three representative experiments of transposon mutants with decreased colonization. One mutant representative of each locus is shown, including: EMH12 (topA), SLV15 (hns), SLV16 (phoQ), and EMH5 (guaB).

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Not all of these colonization phenotypes represented symbiosis-specific defects, however, as topA, pstA, pstC and hns mutants were attenuated for growth in culture in both LBS and SWTO. The topA, pstA and pstC mutants had doubling times 25% longer than ES114 and hns mutants had a doubling time twice that of ES114 (data not shown).

The slower growth rates of these mutants may also be apparent when the strains are within the host and lead to colonization levels lower than that of ES114.

We found it especially interesting that the insertion disrupting inosine-5‟- monophosphate dehydrogenase, guaB, caused a decrease in colonization (Fig. A.1D).

Though this mutant is a guanine auxotroph, it is able to colonize the host light organ, suggesting that the host provides guanine to symbiotic V. fischeri. The guanine auxotroph is the only mutant isolated in this study that maintains a luminescence-up phenotype in the host, with luminescence per CFU within the light organ ~9-fold higher than ES114 (data not shown) despite achieving ~2-fold lower colonization level (Fig.

A.1D). In contrast, the pstA, pstC, and phoQ mutants did not retain their luminescence- up phenotype when in symbiosis with E. scolopes, suggesting that these negative regulators of luminescence are inactive within the light organ. These results could

2+ indicate that both guanine and Mg are provided by the host, though Pi concentrations are low within the light organ. These studies give further insight into the host environment.

Co-culture competitive fitness of luminescence-up mutants

We also tested each mutant for its ability to compete with ES114 in a 1:1 mix for colonization of the host. All of the mutants showed varying degrees of competition

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defects at 48 h post-inoculation, but this is moderate in the arcA, arcB, tRNA, tfoX, phoQ, and guaB mutants, which had RCI per generation values of 0.95-0.99 (Table A.1).

Mutants with disruptions in hns were severely outcompeted, with any remaining hns mutant populations below the limit of detection (Table A.1).

Table A.1: Ability of mutant strains to compete with ES114 in co-cultures

Disrupted In Squida In Cultureb Strain Gene 48 Hour RCI RCI/Generationc RCI/Generation SLV41 arcA 0.49 0.98 0.90 NL3 arcB 0.25 0.95 0.93 EMH12 topA 0.03 0.89 <0.05 SLV32 lonA 0.02 0.88 0.96 SLV10 pstA 0.12 0.93 0.78 SLV30 pstC 0.04 0.90 0.78 SLV15 hns <0.05 <0.05 <0.05 EMH7 tRNA-MET 0.55 0.98 0.86 NL4 tRNA-THR 0.63 0.99 0.90 NL1 tfoX 0.48 0.98 1.02 SLV16 phoQ 0.30 0.96 0.94 EMH5 guaB 0.40 0.97 0.93 NL2 ainS NDd ND 0.95

aAll data calculated for n≥10 animals. bCo-culture data collected from cultures grown in SWTO and calculated from n=2 cAssuming 30 generations for 48 hours in squid host. d„ND‟ indicates experiment was not done due to completion in previous publication (5).

We considered the possibility that colonization defects of certain mutants may reflect general strain attenuation rather than a symbiosis-specific phenomenon. To test this we determined the growth rates and competitiveness of each strain co-cultured with

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ES114. Mutants with insertions in topA, pstA, pstC or hns were attenuated for growth in both LBS and SWTO (Table A.1 and data not shown). The topA, pstA, and pstC mutants had doubling times 25% longer than ES114 and hns mutants had a doubling time twice that of ES114. Moreover, most of the mutants showed RCI per generation values in culture similar to the values estimated from the host colonization experiments (Table

A.1). Mutations that show differing competition phenotypes in co-culture and in the squid host include the topA, pstA, pstC and lonA mutants. Strains with mutations in topA, pstA, and pstC have a larger competition defect per generation in LBS co-culture that in the host (Table A.1). This may suggest that the squid host is providing a necessary condition or nutrient that enables the survival of these mutants. In contrast, the lonA mutants are outcompeted to a greater extent in the host than in culture. Thus it appears that the lonA mutants have a symbiosis-specific defect whereas other mutants may be generally attenuated.

Motility of luminescence-up mutants

It was previously suggested that motility and luminescence may be co-regulated, and altered motility in V. fischeri can result in colonization defects (3, 6, 8). We therefore examined the motility of the luminescence-up mutants. Although most of the mutant strains were similar in motility to their ES114 parent, topA, hns, and tfoX mutants are 45-60% less motile than ES114 (data not shown). Consistent with a previous report

(4), the ainS mutants spread ~6% more quickly than ES114 in soft agar (data not shown).

Disruptions in topA, hns, and ainS resulted in both luminescence and motility phenotypes. Both topA and hns are global regulators and therefore it is not surprising that

176

these mutants have multiple phenotypes. Furthermore, the topA and hns mutants are deficient in colonization (Fig. A.1B and D), which may be explained by the growth and/or motility phenotypes of these strains. Taken together, these results support the notion that certain regulators may modulate both luminescence and motility, but also show that control of luminescence does not significantly affect motility in most cases.

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References

1. Bose, J. L., U. Kim, W. Bartkowski, R. P. Gunsalus, A. M. Overley, N. L.

Lyell, K. L. Visick, and E. V. Stabb. 2007. Bioluminescence in Vibrio fischeri is

controlled by the redox-responsive regulator ArcA. Mol. Microbiol. 65:538-553.

2. Bose, J. L., C. S. Rosenberg, and E. V. Stabb. 2008. Effects of luxCDABEG

induction in Vibrio fischeri: enhancement of symbiotic colonization and

conditional attenuation of growth in culture. Arch. Microbiol. 190:169-183.

3. Graf, J., P. V. Dunlap, and E. G. Ruby. 1994. Effect of transposon-induced

motility mutations on colonization of the host light organ by Vibrio fischeri. J.

Bacteriol. 176:6986-6991.

4. Lupp, C., and E. G. Ruby. 2005. Vibrio fischeri uses two quorum-sensing

systems for the regulation of early and late colonization factors. J. Bacteriol.

187:3620-3629.

5. Lupp, C., M. Urbanowski, E. P. Greenberg, and E. G. Ruby. 2003. The Vibrio

fischeri quorum-sensing systems ain and lux sequentially induce luminescence

gene expression and are important for persistence in the squid host. Mol.

Microbiol. 50:319-331.

6. Millikan, D. S., and E. G. Ruby. 2002. Alterations in Vibrio fischeri motility

correlate with a delay in symbiosis initiation and are associated with additional

symbiotic colonization defects. Appl. Environ. Microbiol. 68:2519-2528.

7. Millikan, D. S., and E. G. Ruby. 2003. FlrA, a sigma54-dependent

transcriptional activator in Vibrio fischeri, is required for motility and symbiotic

light-organ colonization. J. Bacteriol. 185:3547-3557.

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8. Millikan, D. S., and E. G. Ruby. 2004. Vibrio fischeri flagellin A is essential for

normal motility and for symbiotic competence during initial squid light organ

colonization. J. Bacteriol. 186:4315-4325.

9. Ruby, E. G., and L. M. Asato. 1993. Growth and flagellation of Vibrio fischeri

during initiation of the sepiolid squid light organ symbiosis. Arch. Microbiol.

159:160-167.

10. Stabb, E. V. (ed.). 2006. The Vibrio fischeri-Euprymna scolopes light organ

symbiosis. ASM Press, Washington, D. C.

11. Stabb, E. V., and E. G. Ruby. 2003. Contribution of pilA to competitive

colonization of the squid Euprymna scolopes by Vibrio fischeri. Appl. Environ.

Microbiol. 69:820-826.

12. Visick, K. L., J. Foster, J. Doino, M. McFall-Ngai, and E. G. Ruby. 2000.

Vibrio fischeri lux genes play an important role in colonization and development

of the host light organ. J. Bacteriol. 182:4578-4586.

13. Visick, K. L., and E. G. Ruby. 2006. Vibrio fischeri and its host: it takes two to

tango. Curr. Opin. Microbiol. 9:632-638.

14. Wei, S. L., and R. E. Young. 1989. Development of symbiotic bacterial

bioluminescence in a nearshore cephalopod, Euprymna scolopes. Mar. Biol.

103:541-546.

15. Whistler, C. A., and E. G. Ruby. 2003. GacA regulates symbiotic colonization

traits of Vibrio fischeri and facilitates a beneficial association with an animal host.

J. Bacteriol. 185:7202-7212.

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APPENDIX B

METHOD FOR EXTRACTING SYMBIOTIC VIBRIO FISCHERI CELLS

FROM THE EUPRYMNA SCOLOPES LIGHT ORGAN

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Introduction

The symbiosis between Vibrio fischeri and Euprymna scolopes is a model system used to study bacteria-animal interactions. Most studies in this field encompass inocula with wild-type or engineered mutant V. fischeri strains to initiate infection. In such experiments the luminescence of V. fischeri can be monitored, and fluorescence-based transcriptional fusions can be used to track expression of certain genes (2). However, studies that require the recovery and collection of V. fischeri cells from the host typically have used the much larger adult animals that are caught in the wild and infected with various non-isogenic symbiont strains (3, 4). Even with adult hosts, recovery of symbionts is either restricted to collection immediately after they are expelled in response to a light cue (3, 4), or requires dissection techniques that will include host cells and proteins while leaving behind some symbiont cells (1). Though the population size within the light organ can be determined by plating homogenized animals, there is no method for separating the V. fischeri population from host tissue debris following homogenation of the colonized animal.

In several investigations it would be useful to recover symbionts from the light organs of juvenile squid colonized by specific strains; however, this has not been possible. In this appendix, I present a method using V. fischeri antisera in combination with magnetic beads that extracts bacterial cells from the homogenate matrix. This technique employs antisera generated against V. fischeri using a rabbit model. IgGs present within the antisera associate with both symbiont cells and with magnetic beads linked to protein A. Thus, in the presence of V. fischeri, a complex is formed wherein the

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bacterial cells associate with the IgGs and the IgGs associate with protein A on the magnetic beads, which can then be separated from other tissue using a magnet.

Materials and Methods

Colonization of E. scolopes

Twenty-five aposymbiotic E. scolopes hatchlings were inoculated with V. fischeri

ES114 harboring a plasmid (pMTVI), which constitutively expresses gfp (see Table 4.1).

This was accomplished by placing the animals into 100 ml of Instant Ocean (Aquarium

Systems, Mentor, OH) that contained V. fischeri as previously described (5, 6). Briefly, the inoculum was prepared by growing the culture to an OD595 ~0.5 in 5 ml of SWTO within a 50-ml conical tube at 28ºC without shaking. The culture was diluted to ~2000

CFU ml-1 in Instant Ocean. Twenty-five hatchling squid were placed in the inoculum for

~14 hours then rinsed with V. fischeri-free Instant Ocean. At 48 h post-inoculation, the twenty-five infected squid were combined in 1 ml of V. fischeri-free Instant Ocean and homogenized, then this pooled squid homogenate was serially diluted and plated onto

LBS. These plates were incubated overnight at 28°C and the colonies were counted to calculate CFU ml-1 within the pooled squid homogenate.

Extraction of V. fischeri from squid homogenate

The pooled squid homogenate was combined with 100 μl of V. fischeri antisera

(kindly provided by Karen L. Visick) and 5 μl Protein A Magnetic Beads (New England

BioLabs, Ipswich, MA) and incubated at room temperature for 1 h with shaking (20 rpm). Following incubation, the magnetic beads were collected for 5 min from the squid

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homogenate pool using a magnetic centrifuge tube stand. The supernatant was removed and the magnetic beads were resuspended in 1 ml of V. fischeri-free Instant Ocean. Both the magnetic bead suspension and the supernatent were serially diluted and plated onto

LBS. These plates were incubated overnight at 28ºC and the colonies were counted to calculate CFU ml-1 of V. fischeri present in each sample.

Results and Discussion

To examine the efficiency of using magnetic beads and V. fischeri antisera to extract bacterial cells from animal homogenate, samples were serially diluted and plated at three key steps in the assay. First, the pooled squid homogenate was plated to determine the total V. fischeri CFU. Second, the magnetic bead resuspension fraction was plated to determine the CFU of bacteria cells that were extracted from the pooled squid homogenate. And third, the supernatant removed following bead extraction was plated to calculate the CFU that remained in the pooled squid homogenate. The results of these platings revealed that ~96% of the V. fischeri cells that were present within the pooled squid homogenate were extracted using the antisera and magnetic beads (Fig.

B.1).

3500000 3000000 2500000 2000000

1500000 CFU / ml/CFU 1000000 500000 0 Light Organ Bead Extraction Supernatent Homogenate

Fig. B.1: V. fischeri CFU at each step of the extraction assay. CFU was calculated within 1 mL using serially diluted and plated samples. Bars represent standard error for n=3.

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The interaction between the magnetic beads and V. fischeri cells was also directly examined. Because the cells used to infect E. scolopes harbored a constitutively active promoter-gfp fusion, the V. fischeri cells were easily visualized using fluorescence microscopy. Images were obtained following incubation with antisera and magnetic beads. The microscopic images observed indicated that V. fischeri cells were all within tight groupings (Fig. B.2), suggesting that the bacterial cells were associated with the

IgGs and magnetic beads added to the squid homogenate. These observations of V. fischeri within apparent IgG-bead complexes are consistent with the high recovery rate of

CFU in the bead extraction fraction (Fig. B.1).

A B

Fig. B.2: Microscopic images of V. fischeri-IgG-protein A magnetic bead complexes. (A) Phase contrast light microscopy and (B) fluorescence microscopy images of a V. fischeri grouping at 100x magnification.

Overall, these results suggest that using V. fischeri antisera in combination with magnetic beads linked to protein A is an effective method for extracting bacterial cells from squid homogenate debris. One limitation of this approach is the 1-hr incubation time used to allow formation of the symbiont-bead complexes. This incubation time is consistent with similar techniques used in other systems; however, shorter incubation times should be examined to limit changes in gene expression upon E. scolopes

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homogenation. Additional strategies such as high temperature, cross-linking, or phenol-

EtOH may also be used to stop V. fischeri cellular processes immediately following homogenation of colonized animals.

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References

1. Boettcher, K. J., and E. G. Ruby. 1995. Detection and quantification of Vibrio

fischeri autoinducer from symbiotic squid light organs. J. Bacteriol. 177:1053-

1058.

2. Dunn, A. K., D. S. Millikan, D. M. Adin, J. L. Bose, and E. V. Stabb. 2006.

New rfp- and pES213-derived tools for analyzing symbiotic Vibrio fischeri reveal

patterns of infection and lux expression in situ. Appl. Environ. Microbiol. 72:802-

810.

3. Graf, J., and E. G. Ruby. 1998. Host-derived amino acids support the

proliferation of symbiotic bacteria. Proc. Natl. Acad. Sci. U. S. A. 95:1818-1822.

4. Nyholm, S. V., and M. J. McFall-Ngai. 1998. Sampling the light-organ

microenvironment of Euprymna scolopes: Description of a population of host

cells in association with the bacterial symbiont Vibrio fischeri. Biol. Bull. 195:89-

97.

5. Ruby, E. G., and L. M. Asato. 1993. Growth and flagellation of Vibrio fischeri

during initiation of the sepiolid squid light organ symbiosis. Arch. Microbiol.

159:160-167.

6. Stabb, E. V., and E. G. Ruby. 2003. Contribution of pilA to competitive

colonization of the squid Euprymna scolopes by Vibrio fischeri. Appl. Environ.

Microbiol. 69:820-826.

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