The Aromatic Amino Acid Responsive Tyrr Transcription Factor of Enterobacter

The Aromatic Amino Acid Responsive TyrR Transcription Factor of Enterobacter

cloacae UW5: Its Role in Regulation of Indole-3-Acetic Acid Biosynthesis and the

Identification of an Expanded Regulon Using RNA-Sequencing

Thomas J.D. Coulson

B.Sc. University of New Brunswick 2008

A Dissertation Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

in the Graduate Academic Unit of Biology

Supervisor: Cheryl L. Patten, PhD, Department of Biology

Examining Board: Rene Malenfant, Ph.D, Biology Shawn MacLellan, Ph.D, Biology Ghislain Deslongchamps, Ph.D, Chemistry

External Examiner: Ji Yang, Ph.D, Microbiology and Immunology, University of Melbourne

This dissertation is accepted by the Dean of Graduate Studies

THE UNIVERSITY OF NEW BRUNSWICK

July, 2019

©Thomas J.D. Coulson, 2019 ABSTRACT

The control of transcription is an important process in all living cells. In the bacterial family Enterobacteriaceae, the transcription factor TyrR controls genes for aromatic amino acid uptake and biosynthesis. In this thesis, I explore the control of genes by TyrR in Enterobacter cloacae UW5, a soil bacterium commonly associated with plant roots that confer beneficial effects on its host and is also an inhabitant of human intestinal microflora and an opportunistic pathogen. Chapter 1 provides a general introduction to bacterial activities in the plant rhizosphere and transcriptional regulation, especially by

TyrR. In Chapter 2, I investigated the regulation of two divergently transcribed genes, ipdC and akr, by TyrR. The ipdC gene encodes indolepyruvate decarboxylase for the production of the plant growth hormone indole-3-acetic acid, which plays an important role in the plant beneficial behavior of E. cloacae. TyrR is required for activation of ipdC by binding a single DNA element upstream of the promoter. All three aromatic amino acids act as cofactors for TyrR to induce ipdC expression. Expression of akr, encoding a putative aldo-keto reductase, was repressed by TyrR independently of aromatic amino acids and involved TyrR binding an atypical DNA site within the promoter. In Chapter 3,

I assembled the E. cloacae UW5 genome sequence, which revealed genes and pathways that contribute to its plant-associated lifestyle and served as a reference for mapping

RNA-sequencing data. In Chapter 4, I delineated the TyrR regulon by comparing transcription profiles in wild-type and tyrR mutant strains of E. cloacae generated through RNA-sequencing. Broad changes in gene expression were identified and several new TyrR members confirmed, including dmpM encoding a methyltransferase that is highly upregulated by tyrosine and phenylalanine, and cpxP and cpxR, which encode ii

components of the envelope stress response. Additionally, pathways for aromatic metabolism, anaerobic respiration, and motility were altered in the tyrR mutant. Chapter

5 summarizes this research that suggests that the E. cloacae TyrR regulon has expanded from that of E. coli to include genes for survival in the diverse environments that this bacterium inhabits and illustrates the expansion and plasticity of transcription factor regulons.

iii

DEDICATION

I dedicate this thesis to my parents, Dana and Heather Coulson, who have supported me both emotionally and financially, allowing me to pursue my lifelong dream of becoming a scientist. I love you both.

ACKNOWLEDGEMENTS

I would first and foremost like to thank my supervisor Dr. Cheryl Patten. I am eternally grateful to have studied under such an amazing mentor, who has supported my efforts over these many years. It has been wonderful to work, study and play in the lab.

I would like to thank my supervisory committee members, Dr. Mike Duffy and

Dr. Bryan Crawford for their input, suggestions and many conversations which have helped shape the direction of my work. And Dr. René M. Malenfant for his input and assistance with analysis of my sequencing data.

Thank you to my family Dana, Heather, Caroline, Victoria, Charlotte, TJ, Shane,

Bell, Mocha and Ru for your love, encouragement and patience.

To my friends and fellow grad students who I have had the pleasure of knowing over the years I thank you for the insightful conversations, assistance in the lab and our times spent at the Grad House.

And finally a super special thank you goes to Nebai, who has been there for me both professionally and emotionally. I look forward to the many adventures we will continue to have together and promise to take you back to Mount Carleton J

Table of Contents

ABSTRACT ...... ii DEDICATION ...... iv ACKNOWLEDGEMENTS ...... v Table of Contents ...... vi List of Tables ...... xi List of Figures ...... xii Chapter 1 Introduction ...... 1 1.1 The Rhizosphere ...... 1

1.1.1 Plant root exudates ...... 3

1.1.2 Rhizobacteria ...... 4

1.1.3 Chemotaxis and motility in the rhizosphere ...... 6

1.1.4 Nutrient utilization by rhizobacteria ...... 11

1.1.5 Mechanisms of plant growth promotion by rhizobacteria ...... 14

1.2 Transcription in prokaryotes ...... 22

1.2.1 Transcription factors ...... 23

1.3 The TyrR transcription factor ...... 26

1.3.1 Structure of the TyrR protein ...... 26

1.3.2 Cofactor-mediated oligomerization of TyrR ...... 29

1.3.3 TyrR DNA recognition and binding ...... 31

1.3.4 TyrR repression of transcription ...... 32

1.3.5 TyrR activation of transcription ...... 37

1.3.6 PhhR: A TyrR homologue in Pseudomonas ...... 39

1.4 Enterobacter cloacae ...... 43

1.5 Objectives ...... 46

1.6 Organization of thesis ...... 47

1.7 References ...... 49

Chapter 2 The TyrR transcription factor regulates the divergent akr-ipdC operons of Enterobacter cloacae UW5a ...... 75 2.1 Abstract ...... 75

2.2 Introduction ...... 76

2.3 Materials and Methods ...... 80

2.3.1 Bacterial strains and culture conditions ...... 80

2.3.2 Transcription start site identification ...... 80

2.3.3 Construction of reporter gene fusions and promoter mutagenesis ...... 82

2.3.4 Reporter gene expression assays ...... 84

2.3.5 Electrophoretic mobility shift assays ...... 85

2.3.6 Statistical analysis ...... 87

2.4 Results ...... 87

2.4.1 Mapping of the akr and ipdC promoters ...... 87

2.4.2 ipdC is positively regulated by TyrR and the aromatic amino acids ...... 89

2.4.3 akr is negatively regulated by TyrR ...... 89

vii

2.4.4 ipdC activation requires the strong TyrR binding site ...... 90

2.4.5 Repression of akr is facilitated by the weak TyrR binding site occluding the

promoter ...... 91

2.4.6 TyrR binds to two sites in the akr-ipdC intergenic region ...... 92

2.5 Discussion ...... 94

2.6 Acknowledgements ...... 99

2.7 References ...... 99

2.9 Supplemental Information ...... 111

2.9.1 List of supplemental figures ...... 111

Chapter 3 Complete genome sequence of Enterobacter cloacae UW5, a rhizobacterium capable of high levels of indole-3-acetic acid productiona ...... 115 3.1 Abstract ...... 115

3.2 Genome announcement ...... 115

3.3 Nucleotide sequence and accession number...... 117

3.4 Acknowledgments ...... 117

3.5 References ...... 117

Chapter 4 Characterization of the TyrR regulon in the rhizosphere bacterium Enterobacter cloacae UW5 reveals overlap with the CpxR envelope stress responsea . 119 4.1 Abstract ...... 119

4.1.2 Importance ...... 120

4.2 Introduction ...... 120

4.3 Materials and Methods ...... 125 viii

4.3.1 Bacterial strains and growth conditions...... 125

4.3.2 Construction of a tyrR deletion mutant...... 125

4.3.3 IAA quantification assay...... 127

4.3.4 RNA isolation and sequencing...... 127

4.3.5 RNA-Seq data analysis...... 129

4.3.6 Pathway enrichment analysis...... 130

4.3.7 Reverse transcriptase-quantitative-PCR...... 130

4.3.8 Purification of His6x-TyrR and electrophoretic mobility shift assays...... 131

4.3.9 Data availability...... 133

4.4 Results ...... 134

4.4.1 RNA sequencing reveals a large number of TyrR-regulated genes...... 134

4.4.2 Orthologues of known E. coli TyrR regulated genes are similarly regulated in

E. cloacae...... 135

4.4.3 RNA sequencing results were validated by quantitative RT-PCR...... 136

4.4.4 The E. cloacae TyrR regulon has an expanded functional role...... 136

4.4.5 TyrR directly activates expression of a predicted SAM-dependent O-

methyltransferase...... 139

4.4.6 The Cpx regulon is perturbed in a tyrR mutant...... 142

4.5 Discussion ...... 145

4.5.1 The TyrR regulon overlaps the Cpx stress response regulon ...... 146 ix

4.5.2 TyrR activates expression of a predicted SAM-dependent O-methyltransferase.

...... 149

4.5.3 TyrR regulates rhizosphere colonization and survival traits ...... 150

4.5.4 Aromatic catabolism is regulated by TyrR...... 152

4.5.5 The TyrR regulon includes genes for anaerobic metabolism ...... 153

4.6 Conclusions ...... 156

4.7 References ...... 156

4.8 Supplementary information ...... 176

4.8.1 List of supplementary tables ...... 176

4.8.2 List of supplementary figures ...... 176

4.8.3 References ...... 191

Chapter 5 General Discussion and Conclusions ...... 192 5.1 TyrR: Beyond amino acid synthesis ...... 193

5.2 Assembly of the E. cloacae genome ...... 197

5.3 A more expansive role for TyrR? ...... 199

5.5 Concluding thoughts ...... 203

5.6 References ...... 206

Curriculum Vitae

List of Tables

Table 1.1. Genes of the TyrR regulon and its regulatory effects...... 71

Table 2.1. Bacterial strains and plasmids used in this study...... 104

Table 2.2. PCR primers used in this study...... 105

Table 4.1. TyrR-regulation of aromatic amino acid transport and biosynthesis genes is similar in E. cloacae and E. coli...... 166

Table 4.2 . Significantly enriched GO terms among the genes up and downregulated in the tyrR mutant...... 167

List of Figures

Figure 1.1. Major IAA biosynthetic pathways in bacteria...... 72

Figure 1.2. The common biosynthetic pathway for aromatic amino acids including the terminal pathways for phenylalanine, tyrosine and tryptophan...... 73

Figure 1.3. Diagram of the promoters regulated by TyrR in E. coli...... 74

Figure 2.1. The akr-ipdC regulatory region of E. cloacae UW5 ...... 106

Figure 2.2. The expression of akr and ipdC is modulated by TyrR...... 107

Figure 2.3. Role of the TyrR boxes in akr-ipdC gene regulation...... 108

Figure 2.4. Binding of TyrR to the TyrR boxes within the akr-ipdC intergenic region. 109

Figure 2.5. Conservation of the akr-ipdC genes and the two TyrR boxes across closely related bacteria...... 110

Figure 4.1. Volcano plot displaying differentially expressed genes in E. cloacae wild type and tyrR mutant strains...... 168

Figure 4.2. Verification of RNA-Seq results by quantitative RT-PCR...... 169

Figure 4.3. Activation of dmpM expression requires aromatic amino acids with TyrR. 170

Figure 4.4. Nucleotide sequence and regulatory elements of the dmpM promoter region.

...... 171

Figure 4.5. Sequence specific binding of TyrR to the dmpM promoter...... 172

Figure 4.6. Nucleotide sequence and predicted regulatory elements in the intergenic region of divergently transcribed cpxP and cpxR...... 173

Figure 4.7. TyrR downregulates expression of the cpx operons...... 174

Figure 4.8. Sequence specific binding of TyrR to the cpxR-cpxP promoter region...... 175

Figure 5.1. Phylogenetic tree of TyrR and its homologue PhhR (A)...... 209 xii

Figure 5.2. The tryptophan terminal biosynthetic pathway (A) and the triosephosphate isomerase bypass (B)...... 210

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Chapter 1 Introduction

1.1 The Rhizosphere

Bacteria have colonized almost every ecological niche on Earth, from subsurface rocks kilometres underground, to boiling black smokers on the ocean floor, sub-zero

Antarctic ice sheets and even as symbionts living within plant or animal hosts (1).

However, the most abundant sources of bacterial biomass and genetic diversity are found within the soil. Here, an estimated 2.6x1030 bacteria are responsible for mineralization, nutrient mobilization and the biological recycling that underlies soil fertility and provides sustenance for other terrestrial species to thrive (1). Multiple biotic and abiotic factors influence the abundance of bacteria found in different soil environments, primarily pH, but also soil texture (particle size), total organic carbon, nitrogen, elements (P, K, Mg) and land utilization strategies such as forestry or grassland (2-5). Total cell densities are

8 12 influenced by these factors, with reported measurements of 10 to 10 bacterial cells per

4 6 gram of soil, ranging from 10 to 10 different species comprising diverse taxonomic units (Proteobacteria, Actinobacteria, Acidobacteria, Bacteroidetes, Verrucomicrobia,

Planctomycetes, Chloroflexi, Gemmatimonadetes, and Cyanobacteria) (6-11). As such, the soil ‘metagenome’, the totality of all genetic material present in a soil sample, is a tremendous pool of genetic diversity, containing many unique genes encoding enzymes of novel biochemical functions (12, 13).

The evolution of plant roots occurred some 410-395 million years ago and symbiotic mutualistic associations between rhizobia and legumes arose roughly 100 million years ago (14, 15). The early proto-roots were likely less efficient in nutrient 1

acquisition and relied on the solubilisation and mineralization activities supplied by endogenous soil bacteria. As plants colonised new environments their association with bacteria coevolved. This concept is encompassed in the term ‘hologenome’, that is, the sum of genetic information of a host and its associated microbiome, as a selectable evolutionary unit (16, 17). The coevolution of plant host and microbiome has lead to a close, mutually-beneficial association, involving the exchange of essential nutrients and bioactive molecules for communication (16, 18). Much like the human microbiome, the phytomicrobiome constitutes an important biological association that affects plant growth and development. Therefore, to understand plant biology requires an understanding of their associated phytomicrobiome (17). In the modern genomics era a new push in the examination of this community is underway. Through the use of powerful new research tools such as high-throughput next generation sequencing and vast increases in available computational power we are beginning to better understand the complexity of plant- bacterial interactions at the level of genomics and gene expression (18). However, this knowledge is far from complete as the soil environment is one of the most complex and dynamic ecosystems in nature. Studies of the molecular mechanisms involved in this symbiosis allow greater insight into the mysteries of bacteria below our feet. The following sections will explore the physical makeup of the rhizosphere and the bacteria that inhabit it, including their colonization, nutrient utilization, and beneficial effects on host plants.

1.1.1 Plant root exudates

The term ‘rhizosphere’ was first coined by Lorenz Hiltner in 1904 to describe the region enriched with bacteria surrounding plant roots (19). The rhizosphere may be defined as the thin layer of soil immediately adjacent to, and surrounding, the plant root, although this definition may often be nebulous. The rhizosphere may also be expanded to include the root surface, the rhizoplane, or interior of the root tissues, the endorhizosphere. The chemical and physical makeup of this region is impacted by secretions of organic compounds from the roots, known as root exudates. This process of rhizodeposition changes soil chemistry. Major sources of rhizodeposition include the sloughing off of root cap and border cells (20), the death and lysis of root cells (21), the secretion of lubricating mucilage from the root cap (22), exudates leaked between the junctions of plant cells (23), active secretions from plant cells via ion transporters and pumps, the release of volatile organic carbon compounds, and the exchange of compounds with endosymbionts (23, 24). A significant portion of the photosynthetically fixed carbon of a plant may be released in this way, comprising upwards of 10-50% depending on the species and growth stage (23, 25). Exudates are typically rich with small organic molecules that are readily oxidizable by microorganisms, and these may include simple sugars, organic acids, and amino acids that serve as carbon or nitrogen sources for microbial growth (21). High molecular weight molecules such as proteins or carbohydrates are found in mucilage, which is secreted from behind the root cap as a lubricant to the growing root tip as it pushes through the soil. The released mucilage also improves soil aggregation and water retention, which aids in the recruitment of bacteria that travel through the water filled pores within the soil (25). The loss of nitrogen in the 3

form of proteins and amino acids, which are energetically costly to synthesize, from the plant reflects an important investment in the rhizosphere (26). While plants may recover some of this lost nitrogen, most is rapidly converted into microbial biomass. Many of the mechanisms of rhizodeposition are shared among different plant species; however, there is great variation in the composition of exudates which depends on the species, the developmental stage of the plant, and environmental conditions (23, 27). Rhizodeposition stimulates colonization by a number of organisms including arthropods, nematodes, fungi and bacteria (28). The half-life of organic molecules in the soil is short, ~3 days, due to their rapid utilization as nutrient sources by rhizobacteria which may respire 60% of the plant deposited carbon (29). Consequently, the qualitative and quantitative composition of the rhizosphere flora will vary considerably with changes in conditions that affect rhizodeposition.

1.1.2 Rhizobacteria

Rhizobacteria are attracted to the nutrients deposited in the rhizosphere by plants

(21). Only a small percentage of the root surface is colonized by rhizobacteria, typically at the junctions of root epidermal cells or emerging root hairs where leakage or secretion of exudates is common (30). Here, rhizobacteria form microcolonies and often biofilms once established on the root surface (31, 32). Bacterial biofilms are adherent mats of either a single or mixed species of cells typically embedded within an extracellular matrix of exopolysaccharides (33-35). Within the biofilm, bacteria may communicate via quorum sensing systems (36). This intercellular communication between bacteria

involves production of signalling molecules that coordinate gene expression and activities that promote community survival (37, 38).

Rhizobacteria metabolise nutrients found in exudates, and mutants deficient in utilization of these nutrients (organic acids, amino acids) have impaired colonization efficiency relative to their wild-type counterparts (39). The size and composition of rhizobacteria populations are dynamic, varying with the spatial-temporal changes in the quantity and composition of exudates from the plant (33). As such, the number of bacterial cells per gram of rhizosphere soil may vary considerably, and decrease sharply as distance from the plant root increases towards the bulk soil (100-1000 fold fewer cells per gram of bulk soil compared to rhizosphere soil) (40). Plant species, and possibly even cultivar, affects the composition of rhizosphere microbial communities (41, 42) and day- night cycles can even affect the bacterial communities due to shifts in carbon deposition

(43). By secreting specific metabolites, plants may select for and cultivate a specific rhizosphere microbiome (23).

Members of the Actinobacteria and Proteobacteria phyla are common rhizosphere inhabitants, but Bacteroides, Acidiobacteria, Firmicutes, Cyanobacteria and many other phyla may also be present (42). Gram-negative, rod-shaped, facultatively anaerobic bacteria predominate. Species of the genus Pseudomonas are overwhelmingly present in rhizosphere samples, as these bacteria have diverse metabolic capabilities that promote rhizosphere fitness (44, 45). Some rhizobacteria are beneficial to the host plant, and are collectively referred to as plant growth-promoting rhizobacteria (PGPR). Many species of the ubiquitous Pseudomonas are PGPR strains; however, beneficial bacteria also belong to other genera including Azotobacter (46), Arthrobacter (47, 48), Bacillius (49), 5

Clostriduim (50), Enterobacter (51, 52), Pantoea (53, 54), Serratia (55), and

Azospirillum (56), to name but a few. Stimulation of plant growth by PGPR may be direct by the solubilization of nutrients, such as phosphate, or by synthesis of plant growth hormones such as indole-3-acetic acid (IAA, discussed in more detail below). Indirectly,

PGPR may benefit host plants by acting as biocontrol agents, suppressing the growth or colonization of plant pathogenic bacteria and fungi. Production of antibiotics, including pyrrolnitrin or phenazine, inhibits pathogenic bacteria (30, 57), and anti-fungal metabolites suppress pathogenic fungi. To be effective, PGPR must first colonize the root and establish themselves in the rhizosphere. The following sections will discuss PGPR traits associated with rhizosphere colonization and fitness.

1.1.3 Chemotaxis and motility in the rhizosphere

Chemotaxis, the initial processes underlying rhizosphere colonisation, is a critical determinant of rhizosphere competence. Following movement toward the root, traits such as adhesion to the root surface via outer membrane bound structures

(lipopolysaccharides, pili), amino acid prototrophy, and catabolism of plant-derived energy sources (carbohydrates, sugars, organic acids) are important for persistence in the rhizosphere.

Chemotaxis is the process used by bacteria to monitor the chemical composition of the environment by detecting effector molecule gradients in the local environment, and to move towards or away from the source of the molecules by altering flagellar activity.

Organic acids and amino acids in root exudates are chemoattractants for rhizobacteria, and their concentration gradient in the soil stimulates the movement of bacteria towards

plant roots in the first step of colonization (58). The molecular basis of the chemotactic response has primarily been studied in Escherichia coli and Salmonella enterica, where protein interactions and the signal transduction are well understood (59). The chemotactic response of E. coli involves a signal relay between two protein complexes, the chemosensory complex located at one pole of the cell, and one or more flagellar motors

(60, 61). Signals are transmitted between the complexes by the CheY response regulator, which interacts with the flagellar motor switch causing flagella to shift from counterclockwise rotation (swimming) to clockwise rotation (tumbling) (61). The bacterial cells tumble briefly and then, in the presence of an attractant, resume forward swimming behaviour. This relay allows bacteria to alter their swimming direction in response to environmental signalling molecules.

Bacteria detect chemoattractants or repellants via methyl-accepting chemotaxis proteins (MCPs) which are specific for a single ligand or class of ligands (62, 63). These dimeric proteins span the cytoplasmic membrane and in the absence of signal, are methylated at a conserved glutamate residue of the cytoplasmic domain (64).

Conformational changes of the MCP induce transmembrane signalling causing autophosphorylation of the cytoplasmic CheA histidine kinase connected to the MCP via the scaffold protein, CheW (61, 64). Large groups of these proteins found in stoichiometries of ~6:2:2 MCP:CheW:CheA monomers are clustered together at the pole of the cell, forming the chemosensory complex (60, 65). CheA rapidly transfers the phosphoryl group to a conserved aspartate residue of the response regulator CheY, which then interacts with the FliM switch of the flagellar basal motor assembly (61, 66-68) causing the rotation of the flagellar rotor to reverse and induce tumbling of the cell. 7

Signal termination is required to alleviate tumbling. The CheZ phosphatase dephosphorylates CheY~P after a short 50-100 millisecond pause (69, 70). This allows the chemotactic signal to induce a period of reorientation, before the cell resumes swimming behaviour. Adaptation to increasing concentrations of signal molecules are mediated by the CheB methylesterase and CheR methyltransferase, which de-methylate and methylate MCPs, respectively. Phosphorylation of CheA also transphosphorylates

CheB, which stimulates its methylesterase activity, stripping the methyl group from the conserved glutamate residue of an MCP (71). This resets the receptor to pre-stimulus levels, allowing the cells to continue swimming and detect increasing levels of attractants.

A large number of MCPs enable rhizobacteria to integrate diverse environmental cues. E. coli with its five MCPs (Tar, Tsr, Trg, Tap and Aer) is simple compared to soil bacteria where the number of chemoreceptors is often far greater. The rhizobacterium

Pseudomonas putida KT2440 has 27 MCPs and Azospirillum strain B510 has 89 MCPs, while soil bacteria in general are predicted to have 13 MCPs on average (63, 72, 73).

Furthermore, rhizobacteria may have multiple loci with homologues of the chemosensory

Che proteins that have unique functions in motility or chemotaxis (63, 74, 75). The expansion of MCPs and chemotaxis systems in soil bacterial genomes is correlated with lifestyle complexity, and reflects the importance of environmental monitoring of local concentrations of specific soil and plant chemicals (73).

Studies with MCP mutants revealed chemosensing to be essential for rhizosphere colonization. The MCPs of Pseudomonas spp. detect a wide variety of amino acids, organic acids and other organic molecules (76) P. fluorescens Pf0-1 relies on three 8

carboxylic acid sensing MCPs for L-malic acid, succinate, and fumarate to colonize the roots of tomato (77, 78). The plant pathogen Ralstonia pseudosolanacearum also colonizes roots by detecting L-malic acid via the McpM receptor (79). Mutations to any of the three amino acid sensing MCPs of Bacillus subtilis abolishes chemotaxis towards

Arabidopsis thaliana root exudates (80). Possession of an MCP that detects a chemoattractant in root exudates recruits specific bacteria to a plant's rhizosphere. To stimulate early colonization by the symbiotic bacterium Sinorhizobium meliloti, alfalfa seeds specifically release high concentrations of proline which is detected by the McpU chemoreceptor (81, 82). Upon infection by the phytopathogen P. syringae DC3000, A. thaliana specifically increases secretion of L-malic acid through its roots to stimulate recruitment of the beneficial bacterium B. subtilis FB17 in an effort to reduce disease symptoms via a process known as induced systemic resistance (83). While L-malic acid is important for Bacillus chemotaxis towards roots, detection of fumarate and possibly succinate are required for the transition from mobile planktonic cells to sessile biofilms

(84).

Bacterial motility is primarily driven by flagella which consist of a basal body inserted into the cell wall, a long flagellar filament attached via a hook apparatus, and motor proteins through which proton translocation powers flagella rotation (85). By alternating the rotation of flagella between clockwise rotation (causing random tumbling) and counterclockwise (swimming) cells can reorient themselves to move in a new direction (85). Bacteria may possess only one, or several flagella, which may be distributed across the cell (peritrichous) or restricted to one (lophotrichous) or both

(amphitrichous) poles. The importance of motility and functional flagella for rhizosphere 9

colonization have been demonstrated with flagella mutants of a number of rhizobacteria.

Non-motile strains of P. fluorescens lacking flagella were impaired in their ability to colonize tomato and potato roots (86, 87). The nitrogen fixing rhizobacterium

Azospirillum brasilense requires its polar flagella for both motility and adhesion to the root cells of wheat seedling (88, 89). A genomic locus encoding a second flagellar apparatus in P. fluorescens F113 confers a hyper-motile phenotype, able to outcompete other strains for colonization of alfalfa roots (32, 90), and conditions in the rhizosphere select for the hyper-motile variants (32, 91). The colonization of lettuce by S. enterica required motility and flagellar chemotaxis towards root exudates (92), while Listeria monocytogenes flagella are required for rhizosphere competence and adhesion to root cells of alfalfa (93).

In addition to motility, flagella in E. coli, Salmonella, and Clostridium difficile are adhesions, where flagellin monomers comprising the filament, or the flagella cap protein are required for virulence (85). It is proposed that both the glycosylated flagellin monomers of the A. brasilense flagella, and its glycosylated flagellar cap are targets for plant lectins, mediating adhesion between the bacterium and the plant (56, 94-96).

Additional surface structures that are components of the outer membrane of gram negative bacteria may also be involved in adhesion to root cells. Lipopolysaccharides

(97-101), fimbria or pili (102-104), exopolysaccharides (39, 105), and major outer membrane porins (106, 107) are important for recognition of and adhesion to plant root cells (108 and references therein). Specific receptors were identified for some of these structures, notably plant lectins for attachment and subsequent nodulation by Rhizobium; however the precise mechanisms of interaction remain unknown for most (39, 109). 10

Species-specific discrimination between plant and bacteria indicate additional receptor requirements for bacterial adhesion. Multiple components of the bacterial envelope play important roles in rhizosphere fitness and colonization.

1.1.4 Nutrient utilization by rhizobacteria

The diverse composition of amino acids, organic acids, sugars and other nutrients found in root exudates provide many different energy and nutrient sources to be exploited by rhizobacteria. Exploitation of resources plays an important role in rhizosphere fitness.

Furthermore, the spatial and temporal availability of plant derived nutrients rapidly changes as the root grows through the soil, leading to the formation of ‘hot spots’ of carbon deposition via exudates, where microbial activity is consequently greater. Thus, bacteria will experience periods of boom and bust, switching from rapid growth to starvation and survival. To be successful, rhizobacteria must have the genetic capacity to scavenge organic nutrients when they are available.

Catabolism and prototrophy of amino acids are important for rhizosphere colonization and subsequent fitness, and plant-derived amino acids are rapidly depleted from the soil by rhizobacteria. Microbial turnover of amino acids ranges from 24-48 hours in soil (110). While amino acids are a common component of root exudates, they are often not available in sufficient quantities to support bacterial growth (111, 112).

Notably, tryptophan was found to be insufficient in alfalfa exudates for growth of

Salmonella (112). Yet in radish seedlings the available tryptophan content is sufficiently high to induce synthesis of IAA, which is only synthesized by some PGPR when exogenous tryptophan is in excess (27, 113, 114). A transposon mutagenesis screen

revealed a P. fluorescens mutant that was unable to catabolize aromatic amino acids was impaired in colonization, and exposure to maize root exudates triggered expression of an aminotransferase gene necessary for lysine catabolism in P. putida (115). The growth of

S. enterica on alfalfa seedlings induced the substantial expression of proteins for amino acid transport and biosynthesis, suggesting that de novo synthesis of amino acids is essential for bacterial growth, and generation of specific auxotrophic mutants confirmed this (112). Studies of auxotrophic mutants indicate that in addition to amino acid uptake, biosynthesis is important for survival of rhizobacteria such as R. etli, A. brasilense and P. fluorescens (111, 116). Rhizobacteria may also catabolize D-amino acids, the alpha- carbon enantiomers of L-amino acids, by producing amino acid racemases (117). D- amino acids are utilized by bacteria for peptidoglycan and teichoic acid synthesis, and are found in some plant root exudates (118).

Organic acids are often found in much greater amounts in root exudates than sugars

(119). Plants may release organic acids to increase the availability of phosphate in soils, which is often limiting (120). Many studies have demonstrated that uptake and catabolism of organic acids are associated with root colonization (121), and that microbial respiration of organic acids in the rhizosphere is rapid, undergoing depletion in as little as 2-3 hours (121). Furthermore, rhizobacteria may directly stimulate plant roots to enhance secretion of particular organic acids. Bacillus subtilis was found to induce the transcriptional activation of rice genes for malic acid biosynthesis (OsMS) and transport

(OsALMT), which correlated with increased levels of measurable malic acid in roots

(122).

1.1.4.3 Utilization of aromatic compounds

The ability to catabolize aromatic compounds is prevalent among soil bacteria where the breakdown of plant derived compounds leads to a large number of available aromatic molecules. Lignin is the most abundant phenolic polymer of the planet and its breakdown by fungi and bacteria (ex: Streptomyces viridosporus T7A, Sphigomonas paucimobilis SYK-6, Rhodococcus jostii RHA1, P. putida mt-2) yields a variety of aromatic compounds that may be further metabolized by rhizosphere bacteria (123).

Common molecules derived from lignin include benzoic acids, biphenyls, diphenyl ethers, cinnamic acids, benzaldehydes, ferulic acids, vannilate, protocatechuic acid, and syringic acid (123). Aromatic compounds are broken down to a limited number of common intermediates, which are fed into central metabolic pathways (124), these pathways are found to be widespread rhizobacteria (123). Some pathways for breakdown of aromatic compounds found in rhizobacteria, including the well-studied aromatic degrading P. putida KT2444, include the beta-ketoadipate, homogentisate, and phenylacetate pathways.

The beta-ketoadipate pathway is widespread among soil bacteria including P. putida, A. tumefaciens, Rhizobium spp. and S. meliloti (124-127). Peripheral pathways feeding the beta-ketoadipate pathway include those for quinate and shikimate, phenylpropanoid compounds (cinnamate, ferulate, coumarate) and other naturally occurring aromatic compounds derived from decaying plant matter (128). These may then be oxidatively cleaved to the TCA intermediates succinate and acetyl-CoA via beta- ketoadipate enol-lactone (128). Phenylacetate is a free intermediate formed in the anaerobic degradation of phenylalanine but can also be found in the environment as a 13

common carbon source. The hybrid aerobic-anaerobic phenylacetate pathway is found in

~16% of sequenced bacterial genomes (129). Peripheral degradation for phenylalanine, 2- phenylethylamine, phenylalkanoic acids, styrene, or ethylbenene may feed into the phenylacetate pathway (130, 131). The homogentisate pathway is the central route for the catabolism of the aromatic amino acids phenylalanine and tyrosine in P. putida (132).

Orthologues of these genes were identified in rhizobacteria Ralstonia, Bradyrhizobium,

Azotobacter, Xanthomonas and Mesorhizobium (132), and a functional homogentisate pathway was identified in Burkholderia xenovorans (133). The presence of multiple aromatic degradation pathways in rhizobacteria underlies the importance of aromatic compounds as exploitable source of carbon and energy in the rhizosphere (45).

Bacterial colonization and proliferation in the rhizosphere are important factors predicating plant health. The movement of bacteria towards roots, and subsequent colonization along their length is dependent on chemotaxis systems, mobility, and adhesins. Metabolic versatility is also an important determinant of rhizosphere fitness, and the ability to utilize diverse carbon sources for energy and biosynthesis of cellular macromolecules allows rhizobacteria to better exploit these environments. Next, we will cover mechanisms of plant growth promotion, and specifically focus on bacterial production of the phytohormone IAA.

1.1.5 Mechanisms of plant growth promotion by rhizobacteria

Once suitably established in the rhizosphere, PGPR can enhance the growth and health of important food crops; thus, there is great interest in their potential applications as biofertilizers. In practice however, the promising effects seen under laboratory

conditions, or small scale greenhouse studies, often do not translate into larger, healthier plants in field trials. More often this is due to the controlled conditions in these experiments, which do not include the additional complexities of the rhizosphere microbiome, or changing environmental conditions. The beneficial effects of PGPR may be either direct, indirect, or a combination of synergistic traits in a single PGPR strain or the entire plant microbiome.

1.1.5.1 Indirect mechanisms

Indirect mechanisms of plant beneficial effects may include detoxification of harmful compounds, removal of heavy metals, and protection against plant pathogens

(bacteria, fungi, nematodes, etc.). Many rhizobacteria are capable of degrading toxic compounds in soils such as polycyclic aromatic hydrocarbons, a process known as rhizoremediation (134). Such strains are of great interest as a mechanism to restore contaminated sites (135). A well-studied example of rhizoremediation is the degradation of naphthalene by naphthalene 1,2,dioxygenase producing P. putida (136-138). The use of rhizobacteria to confer protection of a host against a pathogen(s) is referred to as biocontrol. Biocontrol agents may prevent infection or disease symptoms on a host plant through direct antagonism of a pathogen, or by stimulating host immunity to future infection. Antibiotic synthesis is widespread among soil bacteria, including hydrogen cyanide (139-141), phenazine (142), pyrrolnitrin (143, 144), 2,3-butanediol (145, 146) and polyketide antibiotics (147). Biocontrol bacteria may also sequester iron via production of siderophores, thereby depriving uptake of this important nutrient by pathogenic bacteria (148, 149). Out-competing pathogenic species for nutrients and

available niches may furthermore prevent disease (30). The interaction of rhizobacteria and plant tissues may prime the plant immune system against future pathogenic attacks, this is known as induced systemic resistance. The host plant detects certain elicitors produced by a non-pathogenic strain (lipopolysaccharides, flagella, siderophores, acetyl homoserine lactones or antibiotics), which activate plant jasmonic acid and ethylene signalling pathways, priming the plant against later pathogen challenge (150, 151).

1.1.5.2 Direct mechanisms

Direct mechanisms of plant growth-promotion include nitrogen fixation, nutrient solubilization and the production of plant growth hormones. Nitrogen fixation by diazotrophic bacteria is perhaps the best known example of direct benefit by PGPR. The symbiotic association of Rhizobia species and legumes in root nodules allows the bacteria to fix atmospheric nitrogen and supply the plant with ammonia, and in return the bacteria are supplied by the plant with reduced carbon sources for energy (39, 152). Free living diazotrophic bacteria such as Azospirillum may also fix atmospheric nitrogen and make ammonia available for uptake by plants (153). Rhizobacteria may secrete phosphatases into the environment to increase the levels of available phosphate by solubilizing organic phosphate (154-156). Many PGPR are able to produce siderophores which are small, high affinity iron-chelating molecules that are secreted from the cell into the soil and are reabsorbed by the bacterium, but also plants, thereby supplying the roots with iron (157,

158).

Production of phytohormones by bacteria is a common mechanism of plant growth promotion, yet paradoxically it may also serve as a virulence mechanism for some

phytopathogens. Rhizobacteria may secrete gibberellins, cytokinin, and auxins, or reduce plant ethylene levels through production of 1-aminocyclopropane-1-carboxylate (ACC) deaminase (30). Ethylene is an important plant hormone that modulates growth and development; however, excess ethylene may reduce plant growth or exacerbate stress effects (159). Bacterial ACC deaminase degrades the ethylene precursor ACC to ammonia and alpha-ketobutyrate, thereby reducing ethylene levels (159).

1.1.5.3 Bacterial indole-3-acetic acid

The plant auxin IAA plays a crucial role in growth, development, and behaviour of plants and some algae. In plants, IAA is primarily synthesized at the apical meristems and then transported from or diffused to distal tissues (160). Root synthesis of IAA has been detected as well (161). The generation and maintenance of IAA concentration gradients within plants is crucial to proper plant growth and development of different tissues and organs. The production of IAA by bacteria is widely distributed across taxonomic divisions and environments (114). The application of bacteria capable of synthesizing

IAA improves the development of plant roots by increasing formation of lateral roots

(162), or root hairs (163) which increases nutrient uptake (164). Root development is regulated by the balance of IAA and cytokinin, and bacterial IAA may alter this balance leading to the effects noted above (165, 166). Bacterial IAA may improve host tolerance to abiotic factors such as salinity and drought stress (167-171). However, many plant pathogens also utilize IAA as a virulence mechanism during infection, which may similarly disrupt the auxin:cytokinin balance or otherwise impact the jasmonic acid signalling pathway of plant defence (172, 173). Notably, P. syringae pv. DC3000 (174),

Rhizobium radiobacter (formerly Agrobacterium tumefaciens) (175, 176), Pantoea agglomerans (formerly Erwinia herbicola) (177), and P. savastanoi (178) all rely on IAA biosynthesis as a part of their infection strategy. The human pathogen S. typhimurium also synthesizes IAA in the rhizosphere, and IAA production is implicated in virulence of mammalian hosts (179).

IAA also has a role in bacterial physiology. Exogenous IAA alters gene expression and protein profiles of rhizobacteria. Culturing of Pantoea sp. YR343 with IAA resulted in an increase in proteins responsible for carbohydrate and amino acid metabolism, in addition to exopolysaccharide biosynthesis (180). A similar experiment with

Bradyrhizobium japonicum, a free-living and endosymbiont that both synthesizes and catabolizes IAA, revealed the upregulation of genes for heat, cold, osmotic and desiccation stress, and exopolysaccharide synthesis, while repressing genes for amino acid biosynthesis, energy metabolism (respiratory proteins), translation, transport and binding proteins (181). Exposure of E. coli to IAA induced a generalized stress response by upregulating genes for cell envelope and stress adaptation, in addition to central carbon metabolism and amino acid biosynthesis genes (182, 183). Components of the

TCA cycle and stress tolerance were also activated in S. meliloti cells when exposed to

IAA (184). In A. brasilense, IAA induced changes in protein synthesis, energy production, transporters, and envelope components (185).

Bacterial IAA is primarily synthesized from the precursor tryptophan through one of five biochemical pathways (Figure 1.1), most commonly via the indoleacetamide or indolepyruvate pathways, although tryptophan-independent production is proposed to occur in both bacteria and plants. Tryptophan is produced from chorismate, which is a 18

product of the shikimate pathway and the precursor for all aromatic amino acids (Figure

1.2). Tryptophan is also the most metabolically expensive amino acid to be synthesized in the cell, consuming 74 high-energy phosphate bonds equivalents per molecule (186, 187).

It is perhaps for this reason that IAA production is typically induced in PGPR only when cells are supplied with an exogenous source of tryptophan. Tryptophan is found in plant root exudates where concentrations may vary depending on plant species (119) and in situ measurements have indicated that tryptophan is released at the emergence of secondary roots where bacteria colonize (188).

Synthesis of IAA via the intermediate indole-3-acetamide consists of two steps.

First, tryptophan monooxygenase (iaaM) converts tryptophan by oxidative decarboxylation to indoleacetamide, which is then subsequently hydrolysed by indole acetamide hydrolyse (iaaH) to IAA and ammonia (Figure 1.1). The genes for both of these enzymes have been identified and are predominantly found in phytopathogenic strains of bacteria, including P. syringae, P. savastanoi, Rhizobium radiobacter, and

Pantoea agglomerans. Notably, the iaaM and iaaH genes in each of these pathogens are found on plasmids, which may also encode additional virulence determinants (175). In R. radiobacter, these genes are located in the mobilization region of a tumor-inducing plasmid, which is transferred into plant cells whereupon the bacterial genes are constitutively expressed. Overproduction of IAA leads to tumor formation and opine synthesis (from genes on the transferred DNA) provides a bacterial energy source (189-

191).

A second major route to IAA synthesis is via the intermediate indole-3-pyruvate, which has been detected in both plants and bacteria. This three-step pathway begins with 19

the transamination of tryptophan with alpha-ketoglutarate by either a general or dedicated aromatic amino acid transaminase to yield the intermediate indolepyruvate, and glutamate (Figure 1.1) (192, 193). In the second, rate-limiting step, indolepyruvate is then non-oxidatively decarboxylated by the enzyme indolepyruvate decarboxylase to indole-3- acetaldehyde, which is finally oxidized to IAA. Indoleacetaldehyde is oxidized by indole-

3-acetaldehyde dehydrogenase in Bradyrhizobium amyloliquefaciens (194), and by an aldehyde dehydrogenase in A. brasilense (195). The decarboxylation step catalyzed by indolepyruvate decarboxylase (encoded by ipdC) is the key step to IAA production via this route. Mutational inactivation of the ipdC gene resulted in strains with reduced or abolished IAA production. Homologues of the ipdC gene have been identified in many groups of bacteria that are common inhabitants of both soil, plant and animal hosts. This includes several genera of enterobacteria such as Salmonella (179), Serratia, Klebsiella

(196), Citrobacter, Yersinia, Pantoea (196), Enterobacter (52) and Erwinia, but not E. coli, Shigella or Dickeya species, and in the actinobacteria Mycobacterium and

Corynebacterium. Note that some of these strains are also pathogens which may have the indoleacetamide pathway as well.

Indolepyruvate decarboxylase belongs to a family of thiamine-dependent decarboxylases whose catalytic capabilities include amino acid synthesis (acetolactate synthases) or degradation (indolepyruvate and phenylpyruvate decarboxylases), or production of alcohols (pyruvate decarboxylases). Decarboxylation results from binding of thiamine diphosphate to the keto-carbonyl group of indolepyruvate and, following the protonation of the activated intermediate, indoleacetaldehyde is released. In E. cloacae,

L-tryptophan aminotransferase has a higher affinity for indolepyruvate (Km = 24 µM) then 20

for tryptophan (Km = 3.3 mM); however, when ipdC is expressed, indolepyruvate

decarboxylase, which has a higher affinity for indolepyruvate (Km = 15-20 µM)(197,

198), drives the reaction toward IAA production. In general, the alpha-keto acid decarboxylases have a broad substrate specificity, utilizing both aromatic and branched chain alpha-keto acids as substrates. For example, an alpha-keto acid decarboxylase from

Lactococcus lactis (KdcA) catalyzes the decarboxylation of the alpha-keto acids derived from leucine, valine, phenylalanine and tryptophan to produce aldehydes that contribute to the flavour of cheese (199). The A. brasilense enzyme was originally characterized as an indolepyruvate decarboxylase (200) but was re-designated as phenylpyruvate decarboxylase based on its greater activity with phenylpyruvate as a substrate then with

indolepyruvate (201). Although the enzyme has a higher affinity for indolepyruvate (Km =

0.13 mM) then for phenylpyruvate (Km = 1.08 mM), the catalytic rate is almost 100-fold higher with phenylpyruvate as a substrate (201).

Tryptophan is not only a precursor for IAA synthesis but is known to induce expression of IAA biosynthetic genes. In the soil bacterium E. cloacae UW5, the ipdC gene is induced by tryptophan via the regulatory protein TyrR (52). Mutational inactivation of tyrR abolished ipdC expression and IAA production. Expression of ipdC also increased in the presence of phenylalanine and tyrosine in a TyrR-dependent manner

(52), which supports enzyme analyses that indicate that indolepyruvate decarboxylase catalyzes the metabolism of a broader range of substrates in some bacteria.

1.2 Transcription in prokaryotes

Transcription is the key step in gene expression whereby the coding sequence of a gene is transcribed into mRNA by RNA polymerase, and subsequently translated into a polypeptide by the ribosome. Transcription in prokaryotes is carried out by RNA

polymerase made up of five subunits (ββ’α2ω) which form the core enzyme, and a sixth dissociable subunit known as sigma (σ) which binds the core to form the functional holoenzyme (202, 203). While the core enzyme is capable of transcription elongation and termination, it requires σ to direct it to an appropriate promoter sequence and assist in

DNA melting and transcription initiation (202). A promoter is a short region of DNA found upstream of a gene transcription start site that contains particular elements that direct the expression of the gene. The dominant σ factor in E. coli is σ70 which recognizes

promoters with the canonical sequence TTGACA-N17-TATAAT centred -35 and -10 bp upstream of the transcription start site, and it is responsible for driving expression of most essential genes in the organism (203, 204). Other σ factors may control genes that often have a specialized function in bacterial physiology, such as σ38 which is expressed during late exponential and stationary phases of growth and regulates genes required for nutrient starvation or stress (205-207). The alternate sigma factor σ54 does not share amino acid sequence homology with σ70 and recognizes promoters with different elements centred at

-24 and -12. It primarily activates genes responsible for nitrogen utilization in E. coli

(208-210). While σ factors help direct RNA polymerase to drive transcription from promoters, bacteria rely on additional mechanisms to ensure genes are expressed or repressed in response to growth, environmental and nutritional cues.

1.2.1 Transcription factors

Transcription factors are DNA-binding proteins that recognize specific cis-acting sequences in or around the promoter of a target gene, and modulate its expression either positively or negatively through interactions with RNA polymerase. Transcriptional repressors may negatively regulate expression by binding to the promoter and directly interfering with RNA polymerase binding, or otherwise prevent later stages of transcriptional initiation. In contrast, transcriptional activators will bind the promoter and either by direct contact, or other indirect mechanisms, facilitate the recruitment or stabilization of RNA polymerase at the promoter, or assist in transcription initiation (211,

212). Transcription factors may bind DNA as monomeric proteins, or undergo oligomerization to form dimers, tetramers, hexamers or greater structures (213). The status of transcription factor oligomerization is unique to each protein and may be altered by binding of a target ligand or cofactor (214, 215). Similarly, the DNA sequence composition and length recognized by transcription factors is unique to each protein. The precise DNA sequence that may be recognized and bound is dependent on contacts made between the DNA-binding domain of the transcription factor, and regions of the major or minor grooves of DNA, or the phosphate backbone (216-218). Most operator sites recognized by the DNA-binding domain of a transcription factor are 4-5 base pairs long, which is roughly the length of contact a alpha-helix may make along the major groove of

DNA (219). Protein dimerization allows transcription factors to increase the number of

DNA contacts (8-10 base pairs) and increase the specificity of the interaction, therefore most binding sites are direct or inverted repeats found on the same face of the DNA helix

(219). Some transcription factors may only recognize the promoters of a limited number 23

of genes, such as a divergently transcribed pair of genes forming a bidirectional operon, and are termed ‘local’ acting transcription factors, or alternatively may impact the regulation of hundreds of genes and be considered ‘global’ acting transcription factors

(220). The collection of genes under the control of a single transcription factor is referred to as its ‘regulon’; however, transcription of genes is often influenced by the activities of more than one transcription factor (221). Global acting transcription factors are responsive to broad changes in bacterial physiology, such as stress, starvation, or utilization of alternate carbon sources, and regulate genes belonging to diverse processes and may interact with more then one σ factor, whereas local transcription factors are responsive to a individual metabolite or nutrient (220).

The primary function of transcription factors is to integrate changing environmental cues or molecular signals into appropriate alteration of gene expression. This is often detected via a small ligand (cofactor) binding the transcription factor which may induce a conformational change or otherwise alter its activity. Therefore, in order to function a transcription factor requires at minimum a sensory domain that is responsible for ligand- binding or protein-protein interaction, and a DNA-binding domain to recognize a unique conformation of nucleotide base pairs (222). The TrpR (tryptophan repressor) transcription factor is an example where both domains are present in a single protein. In

E. coli, TrpR regulates the tryptophan biosynthesis operon (223). Tryptophan is normally taken up by the cells via general or tryptophan-specific permeases. When tryptophan is abundant, TrpR is bound by two molecules of tryptophan which alters its conformation allowing it to bind DNA at a site overlapping the RNA polymerase binding site, thereby preventing transcription of the trp operon and endogenous tryptophan biosynthesis (224, 24

225). In the absence of tryptophan, TrpR can no longer bind DNA, and the enzymes for tryptophan biosynthesis are expressed (226, 227). In contrast to solo-functioning cytoplasmic transcription factors, two-component systems comprise two separate proteins: a membrane bound sensory histidine kinase and a cytoplasmic response regulator. The histidine kinase is autophosphorylated at a conserved histidine residue upon activation by an external stimulus, followed by transphosphorylation to a conserved aspartate residue within the response regulator. The activated response regulator may then activate, or repress, transcription of its target gene(s) (228). Two component systems are unique in that the target of the response regulator may either be DNA (a promoter) or another protein (CheY and the flagella basal motor) (68).

There are four described classes of transcriptional activation. In class I activation, a transcription factor is bound at a site upstream of the promoter and recruits RNA polymerase via direct protein-protein interactions between the transcription factor and the

C-terminal domain of the RNA polymerase alpha subunit (229). Such promoters often have sub-optimal elements that do not efficiently recruit RNA polymerase on their own.

This interaction requires the transcription factor to be bound on the same ‘face’ of the

DNA as the RNA polymerase to maintain efficient contacts which are lost if the binding site is rotated to the opposite face of the DNA (214, 219, 230). Class II activation involves the transcription factor binding at a site overlapping portions of the -35 element, and making direct contacts with the sigma factor, the N-terminal domain of the alpha subunit, or other components of the RNA polymerase enzyme (231). Both classes III and

IV involve protein-protein contacts between the transcription factor and RNA polymerase, independent of DNA binding (215). 25

1.3 The TyrR transcription factor

The tyrosine repressor (TyrR) transcription factor, in conjunction with the three aromatic amino acids, plays a major role in the regulation of aromatic amino acid biosynthesis and transport in E. coli, and other gammaproteobacteria (223). It controls a regulon of nine distinct transcriptional units: aroF-tyrA, aroLM, tyrP, tyrB, aroP, tyrR, mtr, aroG and folA (Table 1.1) (214). In most cases, TyrR with tyrosine and occasionally phenylalanine causes negative regulation, or repression, of its target promoters.

Expression of both mtr and folA expression are activated by TyrR with tyrosine or phenylalanine, and tyrP is both activated and repressed by phenylalanine or tyrosine, respectively (214). Each of the aromatic amino acids acts as a cofactor with TyrR, binding at two ligand binding sites and altering its oligomerization status and affinity for

DNA binding. Promoters recognized by TyrR contain one or more recognition motifs related to the consensus sequence TGTAAA-N6-TTTACA, referred to as a TyrR box

(Figure 1.3) (214). Strong TyrR boxes have greater agreement (>10/12 matching bases) and symmetry and are bound by TyrR in the absence of cofactors in vitro, while weak boxes (<10/12 matching bases) are only recognized by TyrR with tyrosine and ATP.

With the exception of aroP, genes repressed by TyrR contain a double box arrangement where the weaker TyrR box overlaps a portion of the RNA polymerase binding site

(214).

1.3.1 Structure of the TyrR protein

TyrR normally exists as a dimer in the cytoplasm, and each polypeptide of 513 amino acids has a molecular weight of 57 kDa (232). Limited proteolysis revealed that

the TyrR protein has three distinct domains (232). The amino-terminal domain ranges from amino acids 1-190 and contains domains and residues important for transcriptional activation, cofactor binding, and dimerization (233, 234). Deletions and point mutations within the N-terminal domain that do not affect the activity of TyrR as a repressor but abolish its ability to activate transcription have been identified, indicating the activation function is located in this domain. Both regions 2-19 and 92-103 contain residues that mediate protein-protein interactions (234-236). The charged side chains of Arg2, Asp9 and Arg10 interact with oppositely charged amino acids of the alpha subunit carboxy- terminal domain (α-CTD) of RNA polymerase (Asp258, Glu261), required for transcriptional activation (236). The crystal structure of E. coli TyrR N-terminal domain reveals conserved protein folds for a ligand binding ACT domain (residues 2-72), and a

PAS domain (residues 80-114) implicated in signal transduction (237).

A short linker which connects the N-terminal and central domains possibly relays signals to the C-terminal DNA binding domain necessary for activation (238). The central domain spans amino acids 191-467 and has sequence homology to the C-terminal domain of the nitrogen regulatory protein C (NtrC) transcription factor that controls nitrogen assimilation pathways (and others: NifA, DctD, XylR, FhlA) (239). NtrC is a member of the AAA+ (ATPases associated with various cellular activities) bacterial enhancer-binding proteins which facilitate σ-54 dependent transcription by hydrolyses of

ATP to cause open complex formation of the promoter (239). While TyrR shares structural and sequence similarity with these proteins, it is lacking a critical GAFTGA motif required for make direct contact with σ-54, and instead TyrR recognizes σ-70 promoters (239). Two ATP binding sites are located in the central domain, which are 27

necessary for ATP-dependent tyrosine binding and ATP hydrolysis (234, 238, 240, 241).

Mutations in the ATP-dependent tyrosine binding site impact its ability to repress transcription of aroF, aroL, tyrB, and tyrP in conjunction with tyrosine due to reduced tyrosine binding capabilities, while activation of mtr and tyrP is unaffected (234, 241).

Such tyrosine-binding mutations have no effect on phenylalanine-mediated repression, indicating that there are two distinct amino acid binding sites on the protein (241). The

ATPase activity of TryR is stimulated by each of the aromatic amino acids, and in the absence of effectors TyrR is incapable of ATP hydrolysis (241). While ATP binding is necessary for tyrosine binding and hexamerization, the role of ATP hydrolysis in repression is unclear. The hydrolysis of ATP may function to facilitate the transition from hexamers back to dimers, freeing up TyrR protein from high-affinity binding sites (223).

The central domain of TyrR is also reported to have both phosphatase and auto-kinase activities. TyrR has zinc-stimulated autophosphorylation activity (242) and it is conserved in the Haemophilus influenzae TyrR protein which lacks the N-terminal domain found in E. coli. Transphosphorylation of NtrC by its cognate histidine kinase

NtrB induces the transcription factor to undergo further oligomerization which transforms it into a functional activator, capable of interacting with σ54 and the RNA polymerase holoenzyme (243, 244). However the biological role of TyrR phosphorylation and autokinase activity in transcriptional control remains unknown. The inhibition of phosphatase activity by the N-terminal domain of TyrR suggests it may be the target of phosphorylation, or perhaps inhibit transphosphorylation of its biological substrate (242).

The carboxy-terminal domain of TyrR contains a Cro-type helix-turn-helix DNA binding motif situated 11 amino acids before the terminus of the protein (amino acids 28

482-502) (234). The crystal structure of the H. influenzae TyrR C-terminal DNA binding domain has been reported and provides evidence for the classic helix-turn-helix structure of the DNA-binding domain (245). The side chains of several residues were identified that may form a hydrophobic cluster important for stabilization of the whole TyrR protein, and similar residues are conserved in the E. coli protein (245). A second group of residues forms a hydrophilic cluster that would be solvent accessible, and could be involved in direct DNA interactions (245). A hinge helix was identified in the 3-D structure, which in the DNA binding domain of the LacI repressor is only fully formed once the protein undergoes oligomerization and is important for making contacts within the minor groove and inducing bends in operator DNA once bound (246). The remainder of helix-2 lies along the major grove of DNA and makes extended contacts with the medial portion of the palindromic arm (247). The final 11 amino acids following the

DNA-binding helix-turn-helix are also necessary for repression, either by maintaining

TyrR structure or to facilitate DNA binding (234).

1.3.2 Cofactor-mediated oligomerization of TyrR

Each of the three aromatic amino acids function as cofactors for TyrR, altering both its activity as a regulator and oligomerization status. Two independent amino acid binding sites have been identified, one ATP dependent, and one ATP independent. Both tryptophan (Kd = 1.2 mM) and phenylalanine (Kd = 260 µM) bind TyrR at the ATP independent site; however, tyrosine only weakly interacts at this location with a lower affinity then tryptophan (232, 248). The second ATP-dependent amino acid binding site is primarily responsible for tyrosine binding, and hexamerization. TyrR binds tyrosine

with different affinities depending on its status as a dimer (Kd = 330 µM) or hexamer

(Kd = 24 µM) (248). TyrR has a higher affinity for tyrosine over the other amino acids, but this may be overcome when tryptophan or phenylalanine are supplied in high concentrations (above those found naturally in the cell). Binding of amino acids by TyrR is mutually exclusive, meaning that TyrR monomers can bind only one amino acid at a time (232). Tyrosine binding of TyrR is ATP dependent, and under normal physiological conditions TyrR is predicted to be saturated by ATP (half maximal saturation of TyrR by

ATP (S0.5), 5-7 µM; cellular ATP, ~3mM) (232, 241). ATP and tyrosine binding is cooperative at a ratio of 1 mol ATP per 1 mol TyrR monomer (232), and this stimulates

ATPase activity (242). The H. influenzae TyrR protein lacks the first N-terminal 191 amino acids, yet binding of ATP to the central domain induces structural changes within the protein that may be translated to the helix-turn-helix DNA binding domain (249).

Subsequent tyrosine binding reverses these changes, and alters affinity for ATP (249).

Both tryptophan and phenylalanine also induced structural changes, but these were unrelated to ATP (249). The conformational changes due to effector binding are responsible for tyrosine-mediated repression and phenylalanine-mediated activation.

Under normal conditions, TyrR exists as a dimer in the cytoplasm, and will bind strong TyrR boxes in the absence of any cofactors (250). In the presence of ATP and tyrosine, TyrR undergoes further oligomerization from a dimer to hexamer, while remaining bound to DNA in situ (241, 250, 251). Cellular concentration of ATP is high enough to ensure constant saturation of TyrR, therefore the in vivo concentrations of tyrosine determine the hexamerization status of the protein. Yet even under saturating tyrosine conditions 35-50% of TyrR within the cell is predicted to remain in the dimeric 30

form, and the effects of molecular crowding in the cytoplasm are proposed to increase localised tyrosine concentrations and promote hexamer formation (hexamer Kd = 60 µM tyrosine) (232, 250). Formation of the hexamer stabilizes the protein-DNA interaction, allowing a TyrR hexamer to cooperatively bind to multiple boxes simultaneously, which may be facilitated by DNA flexing or looping depending on individual promoter architectures (250, 251). The in vivo effects of molecular crowding from proteins and solutes may increase localised TyrR protein concentrations, and thereby enhance its interaction with DNA at strong binding sites (252). The interaction of hexameric TyrR with multiple boxes requires the boxes to be located on the same face of the DNA molecule (251). This requirement was demonstrated by changing the spacing between adjacent boxes such that they were located on opposite faces of the DNA, which abolished TyrR repression of the tyrP and aroF promoters, respectively (253), and activation of tyrP was similarly abolished when the spacing between TyrR boxes and

RNA polymerase binding was altered. In the case of aroL, the promoter undergoes DNA looping when bound by TyrR with ATP and tyrosine, leading to the formation of hypersensitive sites along the DNA backbone every ~10 bp (~1 turn of the DNA) (254).

1.3.3 TyrR DNA recognition and binding

The C-terminal helix-turn-helix domain of TyrR recognizes DNA sequences related

to the palindrome TGTAAA-N6-TTTACA (214). While the core consensus sequence is

18 bp long, DNase protection studies indicate that TyrR covers an additional two bases flanking each ‘arm’ of the palindrome, although their conservation is not as critical (251,

255). Alanine scanning of this domain identified multiple residues that are important for

TyrR box recognition and DNA binding (234). The two residues Arg484 of helix 1 and

His494 of helix 2 make direct contact with the conserved G-N14-C bases of each half of the palindromic arm of the TyrR box (247). TyrR boxes are classified as either weak or strong based on the degree of sequence conservation and symmetry of the palindromic arms and affinity of TyrR binding. A strong TyrR box is bound by TyrR in the absence of any cofactors, and has a high degree of sequence symmetry within the palindromic arms along with an AT-rich spacer sequence (256); these sequences are proposed to be constitutively bound in vivo. Conversely a weak box has fewer then 10 base pairs agreement with the consensus sequence and requires an adjacent strong box bound by a

TyrR dimer, or hexamer, to be bound. Interestingly, when point mutations were introduced to the strong box of tyrP that resulted in its deviation further from the consensus sequence improved repressor binding was observed, indicating that sub- optimal binding sequences are sometimes required in vivo to provide appropriate levels of regulation (247). The ability of TyrR to function as both an activator or repressor of gene expression is largely dependent on the number and location of TyrR boxes in the promoter region, and their affinity for the various forms of the protein bound to each amino acid cofactor. With the exception of aroG, which contains a single TyrR box overlapping its promoter with an unusually GC-rich spacer, all known TyrR regulated genes contain more than one TyrR box.

1.3.4 TyrR repression of transcription

All genes that are repressed by TyrR contain a pair of adjacent weak and strong

TyrR boxes located within the promoter, where the weak box overlaps a portion of the

RNA polymerase binding site or otherwise interferes with RNA polymerase function

(Figure 1.3). In these promoters, the strong box is bound by TyrR in the absence of any cofactors. The adjacent weak box may only be bound by TyrR in the presence of ATP and tyrosine, as a hexamer, or with phenylalanine via dimer-dimer interactions (241, 248,

250). The hexamer may either increase the DNA binding affinity of the protein or situate

DNA binding domains optimally to overcome the less conserved DNA binding sequence of the weak box. The double TyrR boxes required for repression are found in the promoters of aroF-tyrA, aroL, aroP, tyrP and tyrB. A double TyrR box pair is found in the tyrP promoter, which encodes a tyrosine transporter, with the strong box centred 29 base pairs upstream of the -35 element and the weak box completely overlapping the -35 element (257, 258). Tyrosine-induced hexamerization of TyrR allows it to bind across both boxes and prevents RNA polymerase access to the promoter, repressing tyrP expression (258). The tyrB gene encodes an aminotransferase involved in the biosynthesis of both phenylalanine and tyrosine and has a double TyrR box located downstream of the transcription start site. In the presence of tyrosine, a TyrR hexamer binds across both boxes and acts as a roadblock for RNA polymerase, while phenylalanine allows two TyrR dimers to bind cooperatively and inhibit transcription initiation (259). For both tyrB and tyrP, the spacing between the two TyrR boxes is critical to ensure they remain on the same face of the DNA; repression was lost when bases were inserted between the two boxes such that they were on opposite faces of the

DNA (253), preventing the hexamer from binding both locations. aroF encodes the tyrosine-sensitive DAHP synthase isoenzyme, which along with aroG and aroH catalyse the condensation of phosphoenolpyruvate and D-erythrose-4-phosphate to 3-deoxy-D- 33

arabino-heptulosonate-7-phosphate (DAHP) in the first committed step in aromatic amino acid biosynthesis. Expression of aroF is primarily repressed by TyrR in the presence of tyrosine, which allows the hexamer to bind across the strong and weak boxes centered -52 and -29 base pair upstream of the transcription start site, respectively, and prevent access by RNA polymerase (260). Phenylalanine also represses aroF by binding across these boxes as a pair of interacting dimers. A third strong box located -104 base pairs upstream is also bound by the hexamer via DNA looping; however, its contribution to repression is minimal (260, 261). The third DAHP synthase isoenzyme encoded by aroH is primarily repressed by TrpR, and while TyrR was shown to negatively effect expression there is not yet evidence for direct promoter binding (262, 263). The gene aroL encodes shikimate kinase II, an enzyme in the common pathway of aromatic amino acid biosynthesis. A double weak-strong box arrangement is located downstream from the promoter with the weak box partially occluding the transcription start site with a third strong box found two base pairs upstream of the -35 element (264). Unlike most double

TyrR box arrangements, the weak box in the promoter region of aroL does not play an important role in repression (254). Repression of aroL is primarily dependent on TyrR binding of the two strong boxes, which in the presence of ATP and tyrosine allows the

TyrR hexamer to induce DNA looping to sequester the RNA polymerase binding site

(241, 254). The weak box is protected by TyrR with ATP and tyrosine; however, when mutated it does not impact levels of repression (254). In the absence of cofactors TyrR still binds the strong boxes, however no DNA looping is observed (254). It is postulated that the role of the weak box is to fine tune aroL regulation, possibly in conjunction with the nucleoid-associated protein integration host factor (IHF), which mediates TyrR 34

regulation of aroL, causing a slight alleviation of repression (254). Under these conditions, multiple TyrR dimers are bound across the promoter and, together with TrpR, mediate repression (254).

Both tyrR and aroG contain a strong box which overlaps the -35 portion of the promoter, and are repressed by TyrR independently of aromatic amino acid cofactors.

Aromatic amino acids had no impact on TyrR binding the strong box of aroG, and only by increasing tyrR dosage on a multicopy plasmid was aroG repressed (265). The atypical strong box of aroG contains a spacer sequence with a high GC content, which may reduce the affinity of TyrR for this site, despite the otherwise perfect agreement of the palindromic arms with the consensus sequence. Since the TyrR protein has been shown to make contacts with the spacer, preferentially recognizing AT regions, this is likely another fine tuning mechanism used by the protein (266). Similarly, repression of the tyrR promoter itself is due entirely to available TyrR concentrations. No cofactor- mediated repression is observed, despite the existence of a second weak TyrR box located upstream of the -35 element that is bound by TyrR in the presence of tyrosine (256, 267,

268). Only by increasing gene dosage via the introduction of a multicopy tyrR plasmid was repression increased by approximately five-fold (214, 267). Therefore, it is the available levels of TyrR protein that are responsible for repression at these promoters.

During cell growth the volume of the cytoplasm increases as the cell prepares to divide, while at the same time the chromosome copy number increases prior to division. This would dilute the concentration of available TyrR protein within the cell, while simultaneously increasing the number of available DNA binding sites. Tyrosine-induced

hexamerization would also increase TyrR concentrations at the aroF, tyrP and aroL promoters, drawing it away from tyrR and aroG, thus alleviating their repression.

TyrR has a much different mechanism of repression of the aroP gene, which encodes a general aromatic amino acid transporter. There are three distinct promoters associated with the aroP gene: P1 and P2 drive expression of aroP (P1 is the primary promoter), while P3 is located on the opposite strand of the DNA (269). The double TyrR boxes are located +22 and +5 bp downstream of the P1 and P2 transcription start sites, respectively, and comprise a promoter proximal strong box and a distal weak box (Figure

1.3). TyrR binds both boxes as a hexamer with tyrosine or the strong box alone as a dimer with phenylalanine or tryptophan, in all cases repressing expression from aroP P1 and P2 (270). TyrR recruits RNA polymerase to P3 forming a transcriptionally nonproductive complex (271). Because of the overlapping positions of the three alternate promoters, RNA polymerase may bind only one promoter at a time, precluding access to the other two (268-271). While P3 is active in vitro, no transcripts were detected in vivo, indicating the bound RNA polymerase complex is transcriptionally inactive (269). A

TyrR activation mutant failed to repress transcription of P1, as RNA polymerase was no longer recruited to P3 by TyrR-RNA polymerase interaction (270, 271). Both TyrR boxes must be on the same face of the DNA for P3 activation similar to tyrB and tyrP; insertion of 5 bp between the boxes repositions these sites on different faces of the helix and abolishes interaction with P3, and therefore P1 repression (270). Repression of aroP is therefore the result of TyrR preventing expression of the primary promoter by recruiting

RNA polymerase to, and stalling it at, an alternate promoter on the opposite strand (268).

For many promoters regulated by TyrR in response to cofactors, either the RNA polymerase or TyrR binding sites are imperfect. Sub-optimal DNA interactions with these proteins may be a fine-tuning mechanism to adjust expression, and thereby protein synthesis, to appropriate levels. Both P1 and P3 of aroP deviate considerably from the promoter consensus sequence, and both contain promoter elements known as UP elements (269). UP elements are AT rich sequences found -40 to -60 basepairs upstream of some promoters that increase transcription by binding the RNA polymerase alpha- subunit carboxy terminal domain (272, 273). However, the aroP UP elements are shorter than the rrnB UP element, which strongly binds RNA polymerase and greatly stimulates transcription rates (274), and therefore are less likely to exhibit strong effects (269). Such organisation favours TyrR binding to the divergent P3 promoter preferentially over P1, as strengthening of the P1 consensus completely abolished TyrR mediated repression (269,

270).

1.3.5 TyrR activation of transcription

TyrR is also an activator of gene expression by acting as a class I transcriptional activator, interacting with the RNA polymerase α-CTD to either recruit it to the promoter or stabilise its binding to DNA (236, 255). Unlike the aroP example, TyrR will directly enhance expression of the genes tyrP, mtr and folA in response to one or more aromatic amino acid cofactors. Activation involves contacts between the N-terminal domain of

TyrR and the RNA polymerase α-CTD. Mutations to the N-terminal domain have been isolated that abolish TyrR contact with RNA polymerase and its activating capability, which are unaltered in their capacity for repression (275). Complementary mutations to

residues of the RNA polymerase α-CTD predicted to make contact with TyrR were similarly unable to cause TyrR activation of genes (214). In order to function as a transcriptional activator, TyrR must be bound upstream of the -35 element such that it is positioned to interact with the α-CTD, but not interfere with RNA polymerase binding of the promoter. As mentioned above, TyrR represses tyrP in the presence of tyrosine; however, with phenylalanine TyrR will only bind the strong box alone allowing it to make contact with the RNA polymerase α-CTD thereby activating tyrP expression (233,

248, 255). TyrR mutants that are defective in tyrosine-induced hexamerization are unable to repress tyrP and instead activate transcription (241, 248, 255). The spacing of the strong box relative to the -35 element was shown to be sub-optimal for maximal transcriptional activation of tyrP, the strong box is centered 28 base pairs upstream from the -35, and by inserting three base pairs between these sequences such that they were centered 32 base pairs apart, maximal activation was achieved (253). This sub-optimal location likely reflects a fine-tuning mechanism by which TyrR modulates promoter activity to ensure proper mRNA levels, similar to aroG and aroL.

The mtr gene encodes a high affinity tryptophan transporter that is primarily repressed by the TrpR transcription factor binding downstream of the promoter in response to exogenous tryptophan (223), and is activated by TyrR with both tyrosine and phenylalanine. A strong and weak TyrR box are centered -39 and -69 base pairs, respectively, upstream from the -35 element. Maximal activation of mtr occurs with tyrosine, allowing TyrR to bind across both boxes as a hexamer and contact the RNA polymerase α-CTD (236, 241, 276). Phenylalanine may also activate mtr wherein TyrR binds both boxes via dimer-dimer interactions, although to a lesser extent than tyrosine 38

mediated activation (276). TrpR and tryptophan normally repress mtr; however, in the absence of functional TrpR, tryptophan and TyrR may induce activation of transcription

(276). Furthermore, TyrR binding is proposed to alleviate negative supercoiling of mtr and tyrP promoter DNA caused by the nucleoid-associated protein HU and integration host factor (IHF) that contribute to transcription repression in vivo (233, 236).

Screening of an E. coli genomic library for additional TyrR bindings sites via genomic systematic evolution of ligands by exponential enrichment (SELEX) identified the folA gene as a new TyrR target (277). folA encodes dihydrofolate reductase which catalyzes the reduction of dihydrofolic acid, derived from chorismate, to tetrahydrofolic acid. Tetrahydrofolate and its reduced derivatives are important cofactors in DNA and amino acid metabolism, providing one carbon transfers between molecules in a number of reactions (278). TyrR activates folA transcription primarily with tyrosine by binding the strong and weak boxes centred -75 and -114 base pairs upstream of the -35 element as a hexamer, and to a lesser extent with phenylalanine by binding as a dimer (277).

Activation also required IHF mediated DNA bending to bring TyrR and the RNA polymerase alpha subunit into contact, similar to tyrP and mtr (277).

1.3.6 PhhR: A TyrR homologue in Pseudomonas

A TyrR ortholog in Pseudomonas, PhhR, controls genes primarily involved in phenylalanine catabolism, in contrast to the aromatic amino acid biosynthesis and transport genes controlled by TyrR in E. coli. While both TyrR and PhhR share structural similarity at the amino acid level (45.1% identity, 62.7% similarity), and recognize identical DNA binding sequences, PhhR controls expression from both σ-54 and σ-70

dependent promoters, as opposed to TyrR which only recognizes σ-70 promoters in E. coli. A GAFTGA motif found in the central domain of PhhR, which is absent in TyrR

(279), is required to form critical contacts with σ-54 and induce open complex formation

(280, 281). However, a threonine to glutamic acid substitution in the PhhrR GAFTGA motif may explain the ability of this transcription factor to also recognize σ-70 promoters in some species of Pseudomonas. Furthermore, while PhhR can functionally substitute for TyrR in E. coli as a repressor of aroF-tyrA, it does not function as a transcriptional activator in the heterologous strain (279). Although PhhR shares structural homology with the NtrC family of transcription factors, the amino acid sequence of its N-terminal effector binding domain matches more closely to that of DmpR and XylR, two NtrC type transcription factors whose activity is regulated by effector molecule binding to control methyl-phenol and toluene-xylose catabolism, respectively (282, 283).

1.3.6.1 PhhR regulation of phenylalanine and aromatic catabolism

PhhR was initially identified as a regulator of the phhKLMNOPQB operon encoding a meta-cleavage pathway for growth on methylated phenolic compounds by

Pseudomonas putida P35X (282). Located upstream and divergently transcribed from the phh operon, phhR regulates its own expression via binding a TyrR box-like inverted repeat upstream of the σ-54 dependent promoter in conjunction with DNA looping mediated by an IHF-binding site (282). Activation of the main phh operon by PhhR is induced in response to phenol and methyl-phenol compounds and requires a functional rpoN gene (σ-54) (282).

In P. aeruginosa PA01, the phhR gene is transcribed divergently from a phenylalanine catabolic operon (phhABC) (279). Here, expression of the σ-70 phhR promoter is negatively auto-regulated, while expression of the divergent phhA σ-54 promoter is simultaneously enhanced (279). Activation is induced by phenylalanine

(279). PhhR shares many conserved residues with TyrR that are known to be required for tyrosine mediated repression and DNA binding (279). As a result, the P. aeruginosa

PhhR can functionally substitute for TyrR in E. coli to repress expression of aroF, but not to activate mtr (279). In contrast to TyrR, PhhR does not regulate the DAHP synthase genes required for erythrose 4-phosphate and phosphoenolpyruvate consolidation in the shikimate pathway of P. aeruginosa. Thus, the function of PhhR in P. aeruginosa has diverged from that of E. coli (279).

In P. putida KT2440, a model saprophytic soil bacterium, the phhAB operon is primarily involved in catabolism of phenylalanine as a nitrogen source (284). A divergently transcribed phhR also regulates this operon, however from a predicted σ-70- dependent promoter, as expression in a σ-54 mutant background was unaffected (284).

Both phenylalanine and tyrosine elicit PhhR dependent activation of phhA in P. putida.

PhhR is predicted to bind the promoter as a dimer at two sites upstream of the phhA promoter, the proximal site has stronger affinity for PhhR then the distal site (284). There is no evidence to suggest that PhhR functions as a hexamer similar to TyrR, as a conserved glutamate to glycine substitution in the Walker ATP hydrolysis domain is predicted to be necessary for oligomerization of NtrC (284).

1.3.6.2 Expansion of the PhhR regulon

The PhhR regulon of P. putida is primarily responsible for phenylalanine catabolism, homeostasis, and regulation of several genes of unknown function (285).

Microarray analysis in P. putida revealed that PhhR both activates and represses genes in conjunction with phenylalanine, but may also repress expression of some genes in the absence of phenylalanine (285). Genes activated by PhhR with phenylalanine include those for phenylalanine catabolism through the intermediates tyrosine (phhAB), p- hydroxyphenylpyruvate (hpd) and homogentisate (hmgABC) to yield TCA cycle intermediates. In addition, genes encoding an acetoacyl CoA-transferase, putative ABC transporter for phenylalanine uptake, and several others of hypothetical and diverse functions are also upregulated by PhhR with phenylalanine. PhhR also activates several genes independently of phenylalanine, including genes encoding a LysR-type transcription factor and a multi-drug efflux pump. Finally, several genes are repressed by

PhhR alone including phhR itself, DAPH synthase (aroF), shikimate dehydrogenase, and the phenylacetate-CoA thioesterase (paaY) (285). The paaGHIJ operon, responsible for catabolism of phenylalanine via phenylpyruvate-CoA is upregulated independently of

PhhR, and is instead activated under growth conditions that include phenylalanine as a nitrogen source or with phenylpyruvate as a carbon source (285).

In the human pathogen P. aeruginosa, PhhR controls a similar suite of phenylalanine catabolic genes when grown in medium that simulates the sputum of human lungs with cystic fibrosis (286). Under these conditions, aromatic amino acids are proposed to serve as carbon and energy sources for growth, and also stimulate production of Pseudomonas quinolone signal required for survival under adverse conditions (286). 42

Analysis of global gene expression by microarray identified 12 genes downregulated in the phhR mutant involved in phenylalanine/tyrosine degradation to fumarate and acetyl-

Coenzyme A (phhABC, hpd, hmgA, maiA, fahA, dhcAB, atoB). Other genes for aromatic amino acid transport (aroP2) and a predicted dipeptidase were also repressed (286).

PhhR was shown to directly bind σ-54 dependent promoters for phhA, hpd, and dchA, and σ-70 promoters for both hpd and hmgA (286). However, the preferred σ factor required for activation of each promoter was not examined experimentally in this study

(286).

1.4 Enterobacter cloacae

Enterobacter cloacae is a Gram-negative, facultative anaerobic, non-sporulating rod shaped bacterium belonging to the family Enterobacteracea, which also includes the genus Salmonella, Escherichia, Yersinia, Shigella, Citrobacter and Klebsiella, to which it is most closely related. The taxonomy of Enterobacter has undergone multiple revisions with improvements in metabolic and genomic testing (287-291), and includes 22 species

(E. aerogenes, amnigenus, arachidis, asburiae, cancerogenus, cloacae, cowanii, dissolvens, gergoviae, helveticus, hormaechei, kobei, ludwigii, mori, nimipressuralis, oryzae, pulveris, pyrinus, radicincitans, soli, taylorae and turicensis) and a E. cloacae complex comprised of closely related Enterobacter species that share 61-67% genetic identity (E. cloacae, asburiae, hormaechei, kobei, ludwigii and nimipressuralis) (292).

Members of the Enterobacter genus are widely encountered in nature, found as common inhabitants of soil, in association with plants, aquatic environments and as commensal inhabitants of the human gastrointestinal tract. Species of the E. cloacae complex have

important clinical relevance as they are a common cause of nosocomial acquired infections targeting immunocompromised patients facilitated by their high antibiotic resistance, ability to form resistant biofilms on prosthetic joints, and secretion of various cytotoxins (292-294). Members of the E. cloacae complex are also common saprophytic species, found in close association with plants as either beneficial bacterium or pathogens. E. cloacae has been isolated as a pathogen affecting food crops across the world (295-299). Yet many E. cloacae strains are also beneficial endophytes of roots

(300-303) and seeds (304), or rhizosphere and soil dwelling PGPR (305-308) that may possess multiple plant growth-promoting activities including phosphate potassium and zinc solubilization, acetoin production, IAA biosynthesis and siderophore production.

Furthermore, some E. cloacae strains exhibit potent biocontrol activities to control the fungal pathogen Pythium spp. of cucumber and corn (309-311).

E. cloacae UW5 was originally isolated from the rhizosphere of reeds (312, 313) and was demonstrated to significantly improve the length and number of lateral roots and overall length of primary roots of canola seedling (162). E. cloacae UW5 possesses plant growth-promoting traits including IAA biosynthesis (162) and is able to efficiently colonize and persist in the rhizosphere (314). In E. cloacae UW5 the ipdC gene, required for biosynthesis of IAA via the indolepyruvate pathway, is activated by the transcription factor TyrR (52). A strong TyrR box located 33 bp upstream of the -35 elements of the ipdC promoter is bound by TyrR and poised to interact with the RNA polymerase α-

CTD, enhancing transcription (52). Each of the aromatic amino acids induced expression of ipdC in a TyrR-dependent manner, with tyrosine having the strongest effect (52). TyrR regulates σ70 dependent promoters in E. coli, however ipdC appears to be regulated by σS. 44

The expression of ipdC is activated upon entry into stationary phase both in rich and minimal medium at a time when σS would be present, and IAA production is only measured in the culture supernatant after entry into stationary phase (52, 162).

Furthermore, when E. cloacae UW5 was grown with a plasmid inducible σS expression system, activation of the ipdC gene could be induced during logarithmic phase when it is not normally expressed (315). The ipdC promoter has several features that suggest it is bound by σS including an extended -13/-14 sequence, an A/T rich spacer between the transcription start site and -10 hexamer and a substitution in the -35 hexamer not normally tolerated by σ70 (323, 324). Some genes may be transcribed by both σ70 and σS displaying biphasic expression patterns and may be activated in conjunction with a single transcription factor, though not necessarily to the same level of expression in both phases of growth (325-327). The control of genes for the catabolism of aromatic amino acids by

TyrR suggests that this transcription factor impacts genes required for plant association in

E. cloacae UW5 and that the regulon may be more expansive in this bacterium than previously thought.

Following the completion of this thesis, the species designation of E. cloacae

UW5 was reclassified to E. ludwigii UW5. The NCBI average nucleotide identity (ANI) automated procedure (316, 317) compared the submitted genome against type strains and found the UW5 genome was 98.98% identical across 91.95% of its sequence to the type strain E. ludwigii EN-119 (318), compared to 88.17% against E. cloacae ATCC 13047.

Both E. cloacae and E. ludwigii are members of the E. cloacae complex and share a high degree of genome similarity and synteny (319). E. ludwigii also exhibits the same general growth characteristics as other members of the complex (318). Strains of E. ludwigii are 45

found in both clinical isolates and rhizosphere environments, where they may exhibit multiple plant growth beneficial effects and disease suppression (318, 320-322).

Classification of Enterobacter and the E. cloacae complex has undergone frequent revision as large scale sequencing projects supplant earlier relationships established by biochemical tests and DNA-hybridization, allowing for a truer interpretation of the relationship between these species. Despite the change to the species designation, UW5 remains a demonstrated PGPR and will be referred to as E. cloacae throughout this thesis for consistency.

1.5 Objectives

Rhizosphere-dwelling bacteria must contend with complex environmental conditions that undergo rapid changes in the availability of nutrients and stressors.

Survival depends on the ability of the bacterium to translate environmental signals into appropriate gene expression. Amino acids, including aromatic amino acids, are important signalling molecules and nutrient sources. The role of the TyrR transcription factor in the regulation of aromatic amino acids uptake and biosynthesis is well understood in the model bacterium E. coli. However, the TyrR regulon may be altered in other members of the gammaproteobacteria, especially those that can rapidly adapt to different environments such as the human gut and the rhizosphere. In addition, the overlap of the

TyrR regulon with those of other transcription factors is unknown.

The general objective of this thesis is to characterise the transcriptional control of genes in the enteric and rhizobacterium E. cloacae UW5 by TyrR. Three specific objectives are:

1. To characterize the binding and regulation of the ipdC and akr promoters by

TyrR and to determine the interaction of TyrR with a non-canonical DNA binding site

(Chapter 2).

2. To sequence and assemble the genome of E. cloacae to facilitate examination of the TyrR regulon in this bacterium (Chapter 3).

3. To delineate the TyrR regulon by comparing the transcriptomes in wild-type and tyrR mutant strains of E. cloacae, specifically to identify novel genes and biochemical pathways that are controlled either directly or indirectly by TyrR (Chapter 4).

1.6 Organization of thesis

This dissertation follows the format of an article-based thesis with a general introduction (Chapter 1), three data chapters (Chapters 2, 3, 4) and a general discussion

(Chapter 5). The first half of Chapter 1 serves as a general introduction to the rhizosphere and its inhabitants. The second half focuses on the TyrR transcription factor, summarizing the state of knowledge up to the start of this research. This section will serve as the basis for a future review article which will also draw from data presented in

Chapters 2 and 4, and ideas summarized in Chapter 5.

The experimental design of Chapter 2 was initially conceived by Dr. Cheryl

Patten with contributions from myself. The pJQSIPG and pJQSIAPG plasmids used in this study were generated by Julie Ryu. All experimental work described in this chapter

was performed by me with assistance from Dr. Cheryl Patten in statistical analysis of expression data and interpretation of results. Chapter 2 was published as a research article in Public Library of Science (PLOS) ONE in 2015 with me as lead author, and Dr. Cheryl

Patten as co-author and primary investigator.

Chapter 3 was conceived by me, with support from Dr. Cheryl Patten. All experimental work including culturing of bacteria and genomic DNA extraction was performed by me including data retrieval, quality control, assembly and annotation of the genome. Construction of the genomic library and sequencing was performed at the

McGill University and Genome Quebec Innovation Center. Chapter 3 was published in

Genome Announcements in 2015 with me as the lead author, and Dr. Cheryl Patten as co-author and primary investigator.

The final data Chapter 4 was conceived by me, with support from Dr. Cheryl

Patten. All experimental work including culturing, mutant strain construction, preparation of bacterial cultures, RNA isolation and purification, RT-qPCR and gel shift assays were performed by me. Construction of the RNA libraries and sequencing was done at the

McGill University and Genome Quebec Innovation Center. Bioinformatics analysis including retrieval of RNA-seq data, quality control, mapping, and calling differential gene expression was performed by me with assistance from Dr. René M. Malenfant. Dr.

Cheryl Patten assisted in interpretation of results. This Chapter has been prepared for submission to the Journal of Bacteriology with me as lead author, Dr. René M. Malefant as co-author and Dr. Cheryl Patten as primary investigator.

In Chapter 5, I summarize the general findings from each chapter, and propose additional interpretations of the data that may lead to further investigations in future research.

1.7 References

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Table 1.1. Genes of the TyrR regulon and its regulatory effects. Aromatic amino acid cofactors required by TyrR for transcriptional repression (R) or activation (A) are indicated. Adapted from Pittard et al. 2005. Mol Microbiol 55: 16-26. TyrR Regulation Control Cofactor Gene Protein Product Function aroF DAHP synthase tyrosine Chorismate biosynthesis R Tyr sensitive aroG DAHP synthase Chorismate biosynthesis R Phe phenylalanine sensitive aroL Shikimate kinase II Chorismate biosynthesis R Tyr aroP General aromatic amino Tryptophan, phenylalanine, tyrosine: R Trp, Phe, Tyr acid permease proton symport folA Dihydrofolate reducatase Tetrahydrofolate biosynthesis A Tyr mtr Tryptophan transporter Tryptophan:proton symport A Tyr, Phe tyrB Tyrosine aminotransferase Phenlyalanine, tyrosine biosynthesis R Tyr tyrP Tyrosine transporter Tyrosine:proton symport R Tyr A Phe tyrR Transcription factor Dual regulator of TyrR regulon R None

αKG AAT + PLP CO2 GLU L-Tryptophan

TMO + FAD

Indole-3-acetaldoxime Indole-3-pyruvate CO2 H2O

Oxd IPDC + TPP

CO2 Indole-3-acetamide H2O

H2O

NHase H2O

IAH

Indole-3-acetaldehyde Indole-3-acetonitrile NH3

O2 H2O 2 H2O

Aldh + NAD NIT

H2O2 NH3

Indole-3-acetate Figure 1.1. Major IAA biosynthetic pathways in bacteria. Broken arrow indicate reactions for which the enzymes have not yet been identified in bacteria. AAT, aromatic aminotransferase; FAD, coenzyme flavin adenine dinucleotide; GLU, glutamate; IAH, indoleacetamide hydrolase; IPDC, indolepyruvate decarboxylase; αKG, amino acceptor α-ketoglutarate; NIT, indoleacetonitrilase; NHase, indoleacetonitrile hydratase; Oxd, indoleacetaldoxime dehydratase; PLP, coenzyme pyridoxal 5’-phosphate; TMO, tryptophan 2-monooxygenase; TPP, coenzyme thiamine pyrophosphate. Adapted from Patten et al. 2013 Crit Rev Microbiol 39: 395-415

Glucose

Erythrose 4-phosphate Phosphoenol pyruvate

aroF H2O DAHPsyn(trp) aroG DAHPsyn(phe) DAHPsyn(tyr) aroH Pi 3-Deoxy-arabino-heptulonate 7-phosphate

aroB DHQ synthase Pi

3-Dehydroquinate

aroD DHQ Dehydrogenase H2O

3-Dehydroshikimate

NADPH aroE Shikimate Dehydrogenase

+ NADP Shikimate

aroK ATP aroL Shikimate Kinase I, II ADP Shikimate 3-phosphate

PEP aroA EPSP Synthase

Pi 5-Enolpyruvateshikimate-3-phosphate

aroC Chorismate Synthase Pi

trpE trpD Chorismate Chorismate Mutase Gln Glu Anthranilate pheA Synthase tyrA Prephenate Anthranilate Pyruvate PRPP Anthranilate trpD phosphoribosyl PPi transferase Phosphoribosyl anthranilate NAD+ pheA Prephenate Dehydrogenase tyrA Phosphoribosyl CO2 + H2O NADH + CO2 trpC anthranilate Isomerase Phenylpyruvate 4-hydroxyphenylpyruvate 1-(O-carboxyphenylamino) Glu Glu tyrB Tyrosine Aminotransferase tyrB

-1-deoxyribose phosphate αKG αKG

trpC Indolglycerol phosphate Phenylalanine Tyrosine Synthase CO2

Indoleglycerol phosphate

Serine trpA Tryptophan Synthase trpB D-GAP Tryptophan

Figure 1.2. The common biosynthetic pathway for aromatic amino acids including the terminal pathways for phenylalanine, tyrosine and tryptophan. Given gene names from E. coli are located on the left, and corresponding protein on the right. The final two steps of phenylalanine and tyrosine biosynthesis are catalyzed by a shared enzyme. Molecules from central carbon metabolism are boxed, pathway end products are in rounded boxes. αKG, alpha-ketoglutarate; DAHP, 3-deoxy-D-arabinoheptulosonate 7- phosphate; D-GAP, D-glyceraldehyde-3-phosphate; Gln, glutamine; Glu, glutamate; PEP, phosphoenol pyruvate; PRPP, phosphribosyl pyrophosphate; PPi, diphosphate. Adapted from Pittard et al. 2008. EcoSal Plus vol. 3

aroF

aroG

aroL

P1 P2 aroP

folA

mtr

tyrB

tyrP

tyrR

Strong TyrR Weak TyrR TrpR IHF box box

CpxR -35/-10 Transcription elements start site Figure 1.3. Diagram of the promoters regulated by TyrR in E. coli. The positions of strong and weak TyrR boxes relative to the core promoter elements are indicated. Additional DNA binding sites for other proteins are indicated. The three transcription start sites of aroP are sequentially numbered. Adapted from Pittard et al. 2005. Mol Microbiol 55: 16-26.

Chapter 2 The TyrR transcription factor regulates the divergent akr- ipdC operons of Enterobacter cloacae UW5a

Thomas J.D. Coulson1, Cheryl L. Patten1*

a Published in PLoS ONE, March, 2015, 10(3): e0121241. doi:10.1371/journal.pone.0121241

PMID: 25811953

1Department of Biology, University of New Brunswick, Fredericton New Brunswick,

Canada, E3B5A3

*Corresponding Author [email protected]

2.1 Abstract

The TyrR transcription factor regulates genes involved in the uptake and biosynthesis of aromatic amino acids in Enterobacteriaceae. Genes may be positively or negatively regulated depending on the presence or absence of each aromatic amino acid, all three of which function as cofactors for TyrR. In this report we detail the transcriptional control of two divergently transcribed genes, akr and ipdC, by TyrR, elucidated by promoter fusion expression assays and electrophoretic mobility shift assays to assess protein-DNA interactions. Expression of both genes was shown to be controlled by TyrR via interactions with two TyrR boxes located within the akr-ipdC intergenic region. Expression of ipdC required TyrR bound to the proximal strong box, and is strongly induced by phenylalanine, and to a lesser extent by tryptophan and tyrosine.

Down-regulation of akr was reliant on interactions with the weak box, and may also require a second, as yet unidentified protein for further repression. Tyrosine enhanced repression of akr. Electrophoretic mobility shift assays demonstrated that TyrR interacts with both the strong and weak boxes, and that binding of the weak box in vitro requires an intact adjacent strong box. While the strong box shows a high degree of conservation with the TyrR binding site consensus sequence, the weak box has atypical spacing of the two half sites comprising the palindromic arms. Site-directed mutagenesis demonstrated sequence-specific interaction between TyrR and the weak box. This is the first report of

TyrR-controlled expression of two divergent protein-coding genes, transcribed from independent promoters. Moreover, the identification of a predicted aldo-keto reductase as a member of the TyrR regulon further extends the function of the TyrR regulon.

2.2 Introduction

The TyrR transcription factor controls the expression of genes involved in the uptake and metabolism of aromatic amino acids. In Escherichia coli K12, the TyrR regulon consists of eight operons and nine genes (1, 2). Regulation of each transcriptional unit is either positive or negative, depending on the position of TyrR binding sites, known as TyrR boxes, in the promoter region. TyrR boxes that share ten or more nucleotide

matches with the consensus sequence (TGTAAA-N6-TTTACA) are defined as strong, while those with fewer than ten matches are referred to as weak boxes (3). An invariant feature of known and functional TyrR boxes are the G-C residues of each palindromic arm, spaced 14 base pairs apart, which are essential for TyrR binding (4). Strong boxes are bound by TyrR with high affinity in vivo, whereas weak boxes require an adjacent

strong box to bind TyrR (5). The three aromatic amino acids tryptophan, phenylalanine and tyrosine function as cofactors for TyrR, interacting with the protein dimer at two distinct sites and increasing its affinity for binding DNA (1, 6). In the cytosol, TyrR exists as a homo-dimer, comprised of two 57 kDa subunits. However, in the presence of tyrosine and ATP, TyrR undergoes further oligomerization to form a hexamer (7), allowing it to interact with multiple DNA binding sites (8).

TyrR typically represses transcription by preventing access of the RNA polymerase holoenzyme to the core promoter, or by inhibiting the later stages of open complex formation and elongation. A strong TyrR box overlapping the -35 element represses expression of the E. coli tyrP gene in the presence of tyrosine (4, 9). A similarly positioned strong TyrR box, whose sequence perfectly matches the consensus, represses aroG in the presence of phenylalanine and tryptophan (10). TyrR auto-regulates its own expression, repressing the tyrR gene by binding upstream of the -35 sequence (11, 12).

TyrR binding sites downstream of the core promoter may also repress transcription by occluding the transcription start site, as is the case for tyrB in the presence of tyrosine (9).

Expression of the aroP gene from its primary promoter P1 is inhibited when TyrR, bound to two sites downstream of the aroP transcription start site, recruits RNA polymerase to an alternate promoter P3 which overlaps the P1 promoter but is located on the opposite

DNA strand. This recruitment prevents open complex formation at P1, thereby inhibiting transcription (13-15). Repression of the mtr promoter by TyrR is also dependent on DNA structure; negative supercoiling stabilized by the DNA binding proteins IHF and HU is necessary for repression with tryptophan (16).

While the primary mechanism of TyrR transcriptional control is repression, the expression of several genes is enhanced by TyrR. The mtr gene of E. coli contains a pair of adjacent TyrR boxes, one strong (proximal to the promoter) and one weak (distal to the promoter), upstream of the -35 sequence, and separated from each other by eight base pairs (17, 18). In the presence of tyrosine, TyrR binds to the strong box only, thereby activating transcription of mtr. In the presence of phenylalanine, TyrR binds to both sites and further enhances mtr transcription above the level of induction by tyrosine (17, 19,

20). folA is activated via hexameric TyrR in response to tyrosine. Binding of IHF within the folA promoter induces DNA bending and facilitates TyrR interaction with RNA polymerase to enhance transcription (2). Upregulation of genes by TyrR is dependent on interactions between residues in the N-terminal domain of TyrR and the alpha subunit C- terminal domain (αCTD) of RNA polymerase (21).

In E. coli, experimental evidence has shown that the TyrR regulon is restricted to genes required for aromatic amino acid uptake and biosynthesis. However, bioinformatic and functional analysis suggests that TyrR homologues in other bacteria have expanded or different regulons. Microarray assays indicated that the TyrR homologue in pseudomonads, PhhR, is a much more global regulator, controlling 21 genes which, in addition to proteins for aromatic amino acid uptake and biosynthesis, encode enzymes for catabolism of phenylalanine and other aromatic compounds, ABC transporters, a LysR type transcription factor, and several proteins of unknown function (22, 23). Using sequence analysis and comparative genomics to identify transcription networks of

Shewanella spp., the orthologous TyrR was predicted to control a regulon that comprises

35 genes spanning 16 operons (24). The Shewanella TyrR orthologue has undergone 78

regulon expansion, and is proposed to regulate expression of genes for branched chain and aromatic amino acid degradation and components of the glyoxylate shunt of the tricarboxylic acid cycle (24).

Previously, we showed that in the rhizobacterium Enterobacter cloacae UW5, the

TyrR regulon includes the ipdC gene encoding the enzyme indolepyruvate decarboxylase which converts indole-3-pyruvate to indole-3-acetaldehyde, a key step in the indole-3- acetic acid (IAA) biosynthetic pathway (25). Binding of TyrR to a sequence with high similarity to the consensus binding sequence is required for activation of ipdC expression and IAA production. This strong TyrR box is centered directly between ipdC and a divergently transcribed gene, akr, encoding a putative aldo-keto reductase (Figure 2.1A).

In addition, a second sequence that resembles a TyrR box was found upstream of the akr coding sequence (Figure 2.1A and B). This second site contains palindromic arms with high identity to those of the consensus TyrR binding sequence but with non-canonical spacing between them (hereafter referred to as the weak TyrR box). In this study, we determined that akr is a member of the TyrR regulon and that TyrR binds to both predicted sites in the akr-ipdC intergenic region. Using reporter gene expression assays, site-directed mutagenesis and electrophoretic mobility shift assays (EMSAs), we characterize the roles of the strong and weak TyrR boxes in the activation and repression of ipdC and akr, respectively. Genes that are co-regulated are often functionally related and understanding how akr is controlled by TyrR is a first step towards elucidating its possible role in production of IAA, an important signaling molecule in bacterial-plant interactions (26).

2.3 Materials and Methods

2.3.1 Bacterial strains and culture conditions

The bacterial strains and plasmids used in this study are listed in Table 2.1. E. coli and E. cloacae were routinely grown at 37℃ and 30℃, respectively, in either Luria

Bertani (LB) broth or M9 minimal media (Difco). Media was supplemented with antibiotics when required at final concentrations of 100 µg/mL ampicillin, 25 µg/mL kanamycin, 20 µg/mL gentamicin or 5 µg/mL tetracycline. L-Aromatic amino acid supplements to the growth media were supplied at a final concentration of 1 mM when indicated.

2.3.2 Transcription start site identification

5’-rapid amplification of cDNA ends (RACE) was used to identify the transcription start sites of akr and ipdC. Overnight cultures of E. cloacae UW5 were inoculated 1:100 into fresh M9 media with or without 1 mM of each aromatic amino acid

(L-tryptophan, L-phenylalanine, L-tyrosine). Cultures were grown to mid-logarithmic

(OD600 = 0.6, 8 h) and stationary (OD600 = 2.5, 48 h) phase, at which point 1 mL of culture was transferred to a microfuge tube containing 0.5 mL RNAprotect (Qiagen), and vortexed for ~30 s. Cells were then centrifuged, supernatant discarded and the pellet stored at -20℃. Total bacterial RNA was isolated using the RNeasy Mini Kit (Qiagen).

RNA quality was assessed by running a sample on a 2% agarose gel, and RNA quantity and purity was measured spectrophotometrically at 260 nm using a SpectraMax M5 plate reader (Molecular Devices).

A total of 2 µg RNA was treated with amplification grade DNase I (Invitrogen) to remove any contaminating DNA. cDNA synthesis was performed using Superscript III reverse transcriptase (Invitrogen) according to the manufacturer’s recommendations with primers IPIF and IPDC-GSP1 (Table 2.2) for akr and ipdC, respectively. cDNA was purified using Qiagen’s PCR purification kit and end tailed with dCTP using terminal transferase (New England Biolabs) in a reaction containing 1X TdT buffer, 2.5 mM

CoCl2 and 10 mM dCTP. Reactions were incubated at 95℃ for two min and cooled to room temperature before addition of 20 U terminal transferase and further incubation at

37℃ for 20 min. Tailing reactions were stopped by heating to 65℃ for 10 min.

Ten microliters of tailed cDNA were then used in a 50 µL touchdown PCR with an abridged anchored primer (AAP) and gene-specific primers, AKR-GSP2 and IPDC-

GSP2 (Table 2.2) for akr and ipdC, respectively. Touchdown PCR cycling conditions were as follows: initial denaturation at 94℃ for 2 min; 20 cycles of 94℃ for 30 s, 58℃ -

0.5℃ per cycle for 30 s, 72℃ for 30 s; 25 cycles of 94℃ for 30 s, 48℃ for 30 s, 72℃ for 30 s; final elongation at 72℃ for 7 min. Ten microliters of PCR products were used as templates in a second round of PCR performed with a common AUAP primer (Table

2.2) and gene specific primers AKR-GSP2 or IPDC-GSP2 in 50 µL reaction volumes.

Cycling conditions were as follows: initial denaturation at 94℃ for 2 min; 35 cycles of

94℃ for 30 s, 57℃ for 30 s, 72℃ for 30 s; final elongation at 72℃ for 7 min. PCR products were gel purified before cloning.

Both akr and ipdC tailed cDNA fragments were ligated into pGEM-T Easy

(Promega), generating the plasmids pGEM-IP1 and pGEM-AP1 (Table 2.1), respectively, and subsequently transformed into E. coli JM109 cells (Promega). Inserts were then 81

sequenced using standard T7 and SP6 primers (Robarts Research Institute, London,

Ontario, Canada) to identify the transcription start site.

2.3.3 Construction of reporter gene fusions and promoter mutagenesis

The impact of the TyrR transcription factor on expression of akr and ipdC was examined using plasmid-based reporter gene fusions. The suicide plasmids pJQSIPG and pJQSIAPG contain the ipdC and akr promoters, respectively, fused to the uidA reporter gene encoding β-glucurionidase (25). The PipdC::uidA and Pakr::uidA fusions were excised as ApaI-XbaI fragments from pJQSIAPG and pJQSIAPG, respectively, and inserted into a similarly digested derivative of the pBBR1MCS-2 broad host range plasmid (30) lacking the lacZ promoter (pBBRL). The lacZ promoter was excised from pBBR1MCS-2 using the restriction enzymes KpnI and NsiI. Remaining 5’ overhangs were removed by Klenow digestion followed by re-circularisation by blunt end ligation with T4 DNA ligase (NEB) to generate pBBRL. Following insertion of the PipdC::uidA and Pakr::uidA fusions, the resulting plasmids pBBRLI and pBBRLA, respectively,

(Table 2.1) were transformed into E. coli JM109 (Promega), purified and introduced into

E. cloacae UW5 and J35 by electroporation.

The involvement of two predicted TyrR binding sites was examined by introducing point mutations into the binding sites using site-directed ligase-independent

(SLIM) PCR mutagenesis (31) with the pBBRLI and pBBRLA plasmids as templates.

SLIM-PCR utilizes inverse PCR amplification of the template plasmid with two pairs of primers, a tailed pair containing the mutation and a short pair flanking the mutation site.

Post-amplification denaturation and re-annealing produces a mixed population of

products, of which only those containing the targeted mutation are capable of hybridization and therefore re-circularization. Mutations were generated in 25 µL PCR reactions using Phusion High Fidelity DNA polymerase (NEB) and 8% DMSO with the following sets of primer pairs: weak box mutagenesis, WFT2 and WRT2; WFS1 and

WRS1; strong box mutagenesis, SFT1 and SRT1; SFS1 and SRS1; weak box conversion to strong box, WFT1 and WRT1; WFS1 and WRS1 (Table 2.2). A double TyrR box mutant was generated by introducing mutations in the wild-type strong box sequence of a previously generated weak box mutant plasmid. Cycling conditions for all SLIM-PCR reactions were as follows: 98℃ for 3 min; 35 cycles of 98℃ for 10 s; 63℃ for 15 s;

72℃ for 3 min; final elongation at 72℃ for 5 min. Following amplification, 25 µL of

PCR product was treated with 10 U DpnI at 37℃ for 60 min with 5 µL D-buffer (20 mM

MgCl2, 20 mM Tris-CL, pH 8.0, 5 mM DTT) to remove non-mutated template DNA. The reaction was stopped with 30 µL H-buffer (300 mM NaCl, 50 mM Tris-CL pH 9.0,

20 mM EDTA, pH 8.0) before SLIM-PCR hybridisation at 99℃ for 3 min; followed by two cycles of 65℃ for 5 min; 30℃ for 40 min. Plasmid DNA was purified and transformed into CaCl2 competent E. coli S17-1 (λpir) cells. Successful mutagenesis was confirmed by sequencing the plasmids (Robarts Research Institute). Following sequence confirmation, plasmids were introduced into wild-type E. cloacae UW5 and tyrR null mutant E. cloacae J35 by electroporation. The resulting plasmids are listed in Table 1, with their corresponding mutations in Figure 2.1C.

2.3.4 Reporter gene expression assays

ipdC and akr promoter-driven reporter gene expression was quantified using the method of Cowie et al. (2006) (32). Wild-type E. cloacae UW5 and tyrR null mutant E. cloacae J35 cells containing the reporter gene constructs described above were grown in triplicate from independent colonies overnight in LB broth with appropriate antibiotics, pelleted, and washed twice in 1X M9 salts. Cultures were then diluted 100-fold into 1 mL

M9 minimal media in 2.2 mL 96-deep well plates (Eppendorf, Canada), with or without each of the aromatic amino acids. Cultures were grown to mid-logarithmic (OD600 = 0.6,

8 h) and stationary (OD600 = 2.5, 48 h) phase, at which time they were assayed for β- glucuronidase activity. Briefly, 20 µL of cell culture were withdrawn to wells of a 96- well plate (Cellstar, Sigma) containing 80 µL of reaction buffer (50 mM sodium phosphate (pH 7), 50 mM dithiothreitol, 1 mM EDTA, 0.0125% sodium dodecyl sulfate) and 0.44 mg/mL p-nitrophenyl β-D-glucuronide (PNPG), and incubated at room temperature until development of a strong yellow precipitate, at which time the reaction

was stopped with the addition of 100 µL 1 M Na2CO3. Absorbance of the reaction products was measured at 405 nm using a SpectraMax M5 plate reader (Molecular

Devices) and used to calculate specific promoter activity in Miller units (1000 x

OD405nm)/(time x OD600nm x culture volume in reaction). Wild-type E. cloacae UW5 and uninoculated medium were included as controls to assess background levels of absorbance at 405 nm.

2.3.5 Electrophoretic mobility shift assays

The previously constructed expression vector pQEtyrR (Table 2.1) was used to

purify TyrR via an N-terminal His6 tag (25). E. coli M15 carrying plasmids pREP4, which constitutively produces high levels of lac repressor protein, and pQEtyrR were

grown at 37℃ in LB to mid logarithmic phase (OD600=0.6, 2.5 h) before induction with 1 mM isopropyl β-D-thiogalactopyranoside (IPTG) and allowed to incubate for an

additional four hours. His6-TyrR was purified under native conditions using the

QIAexpressionist Ni-NTA metal affinity chromatography kit (Qiagen). The native purification protocol was followed according to the manufacturer’s instructions with the exception of a modified wash step and elution buffer. Wash buffer 1 contained 50 mM

NaH2PO4, 300 mM NaCl, 10 mM imidazole; wash buffer 2 was as described for wash buffer 1 with 10% glycerol; the elution buffer contained 50 mM NaH2PO4, 300 mM

NaCl, 250 mM imidazole, 8% glycerol. Purified protein was quantified using the Quick

Start Bradford protein assay (Bio-Rad), and purity assessed on a 12% SDS-PAGE gel stained with Coomassie brilliant blue G-250.

DNA fragments encompassing the intergenic region of akr and ipdC were amplified with Phusion High Fidelity DNA polymerase (NEB) using the wild-type and mutant promoter fusion plasmids described above as templates. Primers CF1 and DIG-

CR1 (Table 2.2) amplified a 198 bp fragment spanning the full intergenic region (Figure

2.1B), while incorporating a digoxigenin moiety (DIG) at one end. Non-DIG labeled primer CR1 was used in conjunction with CF1 to amplify unlabelled wild-type intergenic

DNA to be used in competition assays. DNA fragments were separated on a 6% native-

PAGE gel and stained with ethidium bromide before target bands were cut out and purified using a PCR purification kit (Qiagen).

Protein-DNA binding reactions were carried out as described previously (25).

Purified His6-TyrR was incubated at specified concentrations (87 nM, 438 nM, 877 nM,

1750 nM) with 1 µg poly d(I-C), 0.1 mM ATP and 0.1 mM of either L-tryptophan, L- phenylalanine or L-tyrosine in binding buffer (20% glycerol, 250 mM NaCl, 50 mM Tris-

Cl (pH 7.5), 5 mM MgCl2, 2.5 mM dithiothreitol, 2.5 mM EDTA) for 5 min at room temperature. Following incubation, 50 ng of specific DIG-labeled DNA was added and incubated at room temperature for a further 30 min. Competition assays were performed by incubating His6-TyrR first with 100-fold molar excess unlabelled wild-type DNA (500 ng) for 5 min in reaction buffer at room temperature, followed by addition of 50 ng wild- type DIG-labeled DNA. Binding reactions were stopped with 5 µL loading buffer (0.25x

Tris-Borate-EDTA, 60% glycerol, 0.02% bromophenyl blue), and loaded onto a 6% native-PAGE gel (6% acrylamide:bis-acrylamide (37:1), 2.5% glycerol, 0.5X Tris-borate-

EDTA, 0.1 mM ATP, 0.1 mM either L-tryptophan, L-phenylalanine or L-tyrosine, 0.15% ammonium persulfate, 0.1% TEMED). Protein-DNA complexes were resolved at 80 volts for 2-3 h at 4℃.

DNA was transferred to a positively charged nylon membrane (Roche) using a

Trans-Blot SD Semi-Dry cell (Bio-Rad) and visualised using the anti-DIG-Detection system according to the manufacturer’s instructions (Roche). Developed membranes were imaged with a Bio-Rad ChemiDoc MP (Bio-Rad).

2.3.6 Statistical analysis

Statistically significant differences in gene expression from each promoter during logarithmic or stationary growth phases were detected by two-way analysis of variance

(ANOVA) with post-hoc comparisons by TukeyHSD test using JMP 7 (33). The genotype (i.e., strains UW5 and J35), promoter mutation, and amino acid treatments were treated as fixed effects, fully crossed with each other when indicated for each test.

Statistical differences were defined as p values of <0.05. Unless stated otherwise, all main effects and interactions were significant.

2.4 Results

2.4.1 Mapping of the akr and ipdC promoters

To understand how TyrR regulates akr and ipdC, the location of the promoters of both genes relative to the predicted TyrR boxes was first determined. 5’-RACE was used to identify the transcription start sites from which the core promoter sequences of the two genes were inferred. The transcription start site for ipdC was found to be a purine base located 26 nucleotides upstream of the ATG start codon (Figure 2.1B). Examination of the sequence upstream of the +1 site revealed candidate -10 and -35 hexamers with strong resemblance to the E. coli sigma-70 binding sequence, and separated by 16 base pairs.

The -10 sequence matches the E. coli sigma 70 consensus sequence at four of six nucleotides, while the -35 sequence matches at five out of six nucleotides. Within the 5’ untranslated region of ipdC is a strong candidate ribosome binding site. This positions the center of the strong TyrR box 33 base pairs upstream from the center of the ipdC -35 sequence, roughly three turns of the DNA helix. 87

5’-RACE identified a transcription start site for the akr gene 19 base pairs downstream of the proposed ATG start codon at an adenine residue. This was considered to be an artifact based on the following evidence: (i) the akr coding sequence is conserved among closely related bacteria with annotated genomes, including the start codon, and (ii) the identified adenine was located at the end of five contiguous adenines, and reverse transcriptase is known to pause or prematurely terminate activity when it encounters a poly-adenine tract (34). The genome wide transcription start sites of the closely-related enterobacterium Salmonella enterica sv. typhimurium SL1344 were identified by RNA sequencing, including those for the homologous ipdC and akr genes

(35). Alignment of the S. enterica SL1344 and E. cloacae UW5 akr-ipdC regions suggested that the E. cloacae UW5 akr transcription start site is a cytosine nucleotide 25 base pairs upstream of the akr start codon (Figure 2.1B). The S. enterica ipdC transcription start site identified by Kroger et al. (2012) is consistent with the E. cloacae

UW5 ipdC transcription start site identified in this study. Visual inspection of the sequence upstream of the predicted E. cloacae UW5 akr transcription start site revealed a possible -10 element that matches four out of six nucleotides of the sigma-70 consensus sequence (36). This positions the predicted TyrR weak box to overlap the -10 element which supports the proposal that TyrR functions as a repressor of akr. No definitive -35 hexamer was identified upstream of the -10, although the lack of a -35 element does not preclude RNA polymerase interaction (37). A candidate ribosome binding site sequence was found seven base pairs upstream of the ATG within the 5’-UTR of akr.

2.4.2 ipdC is positively regulated by TyrR and the aromatic amino acids

To study the regulatory effects of TyrR on the akr and ipdC genes, uidA transcriptional fusions were generated using a derivative of the low copy number plasmid pBBR1MCS-2 (30) which lacks the lacZ promoter. Consistent with previous results (25), when wild-type E. cloacae UW5 cells containing the ipdC::uidA fusions were grown and assayed for β-glucuronidase activity in stationary phase, expression from PipdC was 3.5- fold greater than that in the tyrR null mutant E. cloacae J35 strain which lacks a functional TyrR (Figure 2.2A). In the wild-type background, expression from ipdC increased significantly in response to each of the aromatic amino acids, relative to that in

M9 minimal medium (Figure 2.2A). Phenylalanine and tryptophan had the strongest effects, increasing ipdC expression four-fold and two-fold, respectively, above M9 minimal media alone in stationary phase. Tyrosine also induced expression 1.5-fold above that in minimal media; however, this was less then that previously observed (25).

Expression of ipdC was on average 3.6-fold greater in stationary phase relative to logarithmic phase. This is consistent with the observation that IAA, the product of the reaction catalyzed by IPDC, is not detectable in the culture media of E. cloacae UW5 until stationary phase (25, 29).

2.4.3 akr is negatively regulated by TyrR

In contrast to the positive regulatory effects of TyrR on the ipdC promoter, akr expression was negatively regulated by TyrR (Figure 2.2B). When expression from akr was measured in the tyrR null mutant E. cloacae J35, a 1.6-fold increase was observed (in both log and stationary phases) compared to expression levels in wild-type E. cloacae

UW5, indicating that TyrR normally represses akr. Inclusion of the aromatic amino acids tryptophan and phenylalanine did not result in significant down-regulation of akr; however, the inclusion of tyrosine in the medium resulted in a 1.5-fold decrease in akr expression relative to M9 medium alone during log phase. Transcription from the akr promoter remained at relatively constant levels during both logarithmic and stationary phases of growth when assayed in wild-type E. cloacae UW5.

2.4.4 ipdC activation requires the strong TyrR binding site

To test the involvement of the two predicted binding sites in TyrR-mediated activation of ipdC, four promoter variants were constructed via SLIM-PCR mutagenesis

(Figure 2.1C). These mutations were generated to abolish TyrR binding to the weak and/or strong boxes by changing nucleotides known to be essential for TyrR DNA binding in E. coli (1). The center of the strong TyrR box is located 33 base pairs upstream of the center of the predicted -35 element of the ipdC promoter, positioning the two sites three turns of the DNA helix apart and on the same face of the DNA. This would allow

TyrR to act as a strong type I transcriptional activator in a manner consistent with other genes that are positively regulated by TyrR, such as tyrP (38). Mutation of the strong box

(strains carrying pBBRLIS) resulted in the reduction of ipdC expression to a level similar to that in the tyrR null mutant E. cloacae J35 (Figure 2.3A). Similar results were observed from the double weak and strong box mutant promoter (pBBRLIWS). In both logarithmic and stationary phase, inactivation of the weak box alone (pBBRLIW) was found to have little effect on expression of ipdC (Figure 2.3A).

To examine the effect of TyrR binding at the weak box on the ipdC promoter, the weak box spacing was adjusted such that each arm of the palindromic TyrR box motif was separated by the ideal six nucleotides (corresponding to the same spacer nucleotides as in the adjacent strong box). The high identity of the weak box palindromic arms to the

TyrR consensus sequence, coupled with the optimal spacing would ensure that TyrR is bound with high affinity to both sites (1). The resulting double strong box promoter

(pBBRLISS) increased transcriptional activity 2.1-fold over levels of the wild-type promoter (pBBRLI) (Figure 2.3A). Similar expression trends were observed in the presence and absence of aromatic amino acids, although this effect was enhanced in the presence of aromatic amino acids, especially phenylalanine, as expected for a TyrR mediated response (Supplementary Figure S2.1).

2.4.5 Repression of akr is facilitated by the weak TyrR binding site occluding the promoter

Based on the predicted transcription start site for akr, the weak TyrR box is situated to overlap the predicted -10 sequence of the akr promoter, suggesting that binding of TyrR to the weak box competes with RNA polymerase for access to the promoter. Mutation of the weak box (pBBRLAW) resulted in a 3.5- and 5.3-fold higher level of expression from the akr promoter compared to the wild-type promoter

(pBBRLA) in logarithmic and stationary phases, respectively (Figure 2.3B). The effect of the weak box mutation was greater in the tyrR null mutant E. cloacae J35, particularly in stationary phase, which showed a 16-fold increase in akr expression over the wild-type

UW5 strain. Inactivation of the strong box alone (pBBRLAS) had no effect on akr

expression in either growth phase suggesting that the observed repression is mediated by the weak box (Figure 2.3B). When both TyrR boxes were mutated (pBBRLAWS), transcription from the akr promoter increased 7.3- and 11-fold over levels from the wild- type promoter in both wild-type E. cloacae UW5 and the tyrR null mutant E. cloacae J35, respectively, in stationary phase. Conversion of the weak box to a strong box

(pBBRLASS) yielded the same level of repression as from the wild-type promoter in wild-type E. cloacae UW5; however, in the tyrR null mutant E. cloacae J35, transcription increased 3.8- and 9.3-fold in log and stationary phases, respectively. The effect of the

TyrR box mutations on akr expression was similar in M9 medium with and without aromatic amino acid supplements (Supplementary Figure S2.2). These data indicate that both the weak box and TyrR are important for akr repression, but that TyrR interaction at the weak box does not sufficiently explain the observed repression.

2.4.6 TyrR binds to two sites in the akr-ipdC intergenic region

The ability of TyrR to bind to specific sites within the intergenic region of akr-

ipdC was demonstrated using purified His6-TyrR protein. When incubated with purified

His6-TyrR, a DIG-labeled DNA fragment encompassing the wild-type akr-ipdC intergenic region migrated to a position consistent with a higher molecular weight complex on a native-PAGE gel (complex I), compared to the DNA fragment alone

(Figure 2.4, lanes 1, 2). Migration of the DNA fragment was His6-TyrR concentration dependent, with levels of His6-TyrR greater than 87 nM inducing a second DNA shift to complex II (Figure 2.4, lane 3, and Supplementary Figure S3). These results suggested that TyrR binds to two sites in the akr-ipdC intergenic region. Pre-incubation of His6-

TyrR with a 10-fold molar excess of unlabeled DNA probe abolished the band at the complex II location, however complex I was still apparent in the gel (Supplementary

Figure S2.3, rightmost lane). There was no apparent increase or decrease in affinity for

DNA binding by His6-TyrR when EMSAs were repeated in the presence of aromatic amino acids (Supplementary Figure S2.3).

To determine if His6-TyrR binds to the akr-ipdC intergenic region at the two predicted TyrR boxes, EMSAs were performed using the mutated promoter sequences generated by SLIM-PCR as DNA binding templates for His6-TyrR. The results show that when the weak box was mutated, His6-TyrR still bound the DNA to yield complex I, however complex II was abolished (Figure 2.4, lanes 5, 6). Mutations in the strong box resulted in the loss of intense bands representing both complexes, although the weak bands may indicate some low affinity interactions with the intact weak box at higher

His6-TyrR concentrations (Figure 2.4, lanes 8, 9). All interactions were abolished with fragments carrying mutations in both TyrR boxes (Figure 2.4, lanes 11, 12). When the

DNA fragment with the double strong box was used as a template for EMSAs, all labeled

DNA migrated to complex II, even at the lowest concentration of His6-TyrR (Figure 2.4,

Lanes 14, 15). This indicates that the increased spacing between the palindromic arms of the weak box increased the affinity of TyrR for the weak box and supports strong protein- protein interactions between TyrR dimers bound at both locations on the DNA. The results shown in Figure 2.4 demonstrate that TyrR interacts with the two predicted TyrR binding sites within the akr-ipdC intergenic region. While interactions with the akr proximal box are weaker and require TyrR to be bound to the adjacent strong box in

vitro, this is not the case in vivo where the strong box has no discernible effect on akr expression. In the latter case, additional regulatory factors may be acting at the promoter

2.5 Discussion

In this report we detail the transcriptional control of two adjacent, divergently transcribed genes, akr and ipdC, by the transcription factor TyrR, elucidated by promoter fusion expression assays and EMSAs to assess protein-DNA interactions. Expression of akr and ipdC was controlled by TyrR interaction with a weak and strong TyrR box, respectively, located within the intergenic region. Direct protein-DNA interactions between TyrR and the strong and weak boxes were shown in vitro using purified TyrR protein and DNA fragments containing the akr-ipdC intergenic region; the interactions were abolished when point mutations were introduced in either or both TyrR boxes.

These interactions were strengthened when the sequence of the weak box was altered to match that of a strong box as demonstrated by the shift of the DNA fragment to a higher molecular weight complex when incubated with lower concentrations of TyrR and resolved by EMSA, and by the increased expression from ipdC reporter fusions.

Expression of ipdC is dependent on TyrR for activation; in the tyrR null mutant E. cloacae J35, expression of the PipdC::uidA fusion was abolished (this study; (25)), and production of IAA is also reduced. E. cloacae indolepyruvate decarboxylase, encoded by ipdC, catalyzes the production of IAA (25, 39). In plants, IAA is an important growth hormone, controlling many aspects of growth and morphogenesis such as gravitropism, cell elongation and tissue differentiation (40). E. cloacae UW5 inhabits the rhizosphere of plants, where it may produce high levels of IAA, and has been shown to positively

influence root development (29). Plant growth-promoting rhizobacteria have been applied in field studies and commercial agriculture to improve crop yields, although results are often inconsistent or improvements only marginal (41). Identifying the mechanisms underpinning regulation of IAA production in plant growth-promoting rhizobacteria will aid in their application as agricultural phytostimulants, especially considering that IAA is also a virulence factor produced by some plant pathogens and levels of this hormone may be an important factor in plant interactions (recently reviewed by (26)).

Inclusion of any of the aromatic amino acids in the growth media significantly increased ipdC expression in a TyrR-dependent manner, with phenylalanine as the strongest inducer. The center of the strong TyrR box is located 33 base pairs upstream from the center of the predicted ipdC -35 element. This would position a TyrR dimer bound at the strong box on the same face of the DNA as RNA polymerase bound to the ipdC promoter, and in an optimal location for interactions with the RNA polymerase ⍺-

CTD (1, 38). It is likely that the strong induction of ipdC is due to activation by TyrR complexed with aromatic amino acids, which increases the affinity of TyrR for the DNA and leads to strong activation in promoters with similarly positioned TyrR boxes (21, 42).

The promoter of the mtr gene in E. coli has a similarly orientated strong TyrR box centered 42 base pairs upstream of the -35 element. In the presence of phenylalanine, a

TyrR dimer bound to this strong box interacts with the RNA polymerase ⍺-CTD enhancing expression 2.8-fold (17, 21). A comparison of the TyrR activation of the E. coli mtr promoter and the E. cloacae UW5 ipdC promoter reveals very similar trends.

The mtr gene is activated 2-fold (tryptophan), 2.8-fold (phenylalanine) and 1.7-fold

(tyrosine); while ipdC is activated 2-fold (tryptophan), 3.7-fold (phenylalanine) and 1.4- fold (tyrosine) (20).

The newest member of the TyrR regulon, akr, identified in this study, was shown to be down-regulated by TyrR. Little is known regarding the function of the protein encoded by the akr gene. From the amino acid sequence it is predicted to be a member of the aldo-keto reductase (AKR) superfamily of proteins, a diverse group of enzymes found

in all domains of life that share a common (⍺/β)8 barrel fold and utilize the nicotinamide adenine dinucleotide (NAD(P)H) cofactor to reduce carbonyl compounds (43, 44). AKRs are further divided into families based on amino acid sequence identity (at least 40% identity for family designation). Phylogenetic analysis of the E. cloacae UW5 AKR groups it with E. coli YghZ (data not shown), which is homologous to the AKR6 family that comprises the β-subunit of voltage gated potassium channels (45). The yghZ gene encodes a methylglyoxal reductase, which has been shown to confer protection against the toxic electrophile methylglyoxal by reducing it to acetol (46), and has been designated ARK14A1 based on amino acid identity. Although the function of the E. cloacae UW5 AKR is unknown, the sequence and organization of the open reading frame is conserved among closely related bacteria that possess the ipdC gene: when ipdC is present in a sequenced enterobacterial genome, it is found adjacent to, and divergently transcribed from, a homologous akr (Figure 2.5A). Both the strong and weak TyrR binding sites are also conserved in these genomes (Figure 2.5B). Thus, the common regulation of these two genes by TyrR and their evolutionarily conserved pairing suggest a functional relationship between AKR and IAA biosynthesis.

Down-regulation of akr is dependent on the weak TyrR box which overlaps the predicted -10 element of the akr promoter; however, mutations in the strong TyrR box upstream of the akr promoter had no effect on expression. The observation that expression from the akr mutant weak box promoter was higher than that from the wild- type promoter in the tyrR null mutant E. cloacae J35, when the expectation was that they would be similar, may indicate that the nucleotide changes in the weak box increased the strength of the akr promoter. Alternatively, the increase in expression of the mutant weak box promoter may indicate the involvement of a second regulatory protein. This is supported by the similarly high levels of akr expression in the tyrR null mutant E. cloacae J35 containing the mutant weak or double strong box promoters. In both cases, nucleotide changes were introduced in the weak box sequence (in the case of the double strong box these would not be expected to enhance promoter strength) and may have abolished the binding site for a second regulator. In wild-type E. cloacae UW5 with a double strong box, the increased affinity of TyrR for the weak box maintained repression of akr at wild-type levels.

The observation that akr was down-regulated by TyrR, but not completely repressed may reflect the weak interactions between TyrR and the DNA binding site overlapping the -10 element. A TyrR dimer or hexamer may associate transiently with the weak box allowing RNA polymerase access to the promoter and a low level of expression. This is proposed to be the case with the aroG gene of E. coli, where the TyrR box overlaps the -10 element and competition between the two proteins for DNA binding results in a low but constant level of gene expression (1). Weak interactions between

TyrR and the weak box may be stabilized by protein-protein interactions between TyrR 97

bound to both boxes. These may be dimer-dimer interactions, as in the case of mtr (17) or the formation of a TyrR hexamer. TyrR is thought to polymerize from a dimer to a hexamer in the presence of tyrosine while bound to a strong box sequence which acts as a nucleation point (7). The resulting hexamer has a 2- to 5-fold increased affinity for DNA binding, allowing its interaction with adjacent, weaker binding sites (47). TyrR may polymerize when bound to the strong box in the akr-ipdC intergenic region, increasing its affinity for the adjacent weak box in the presence of tyrosine. This is supported by the requirement for the wild-type strong box for strong TyrR binding to the intergenic region in EMSAs. The observation that mutations in the strong box do not affect akr expression in the reporter gene assays may support the involvement of a second regulatory protein that stabilizes TyrR binding to the weak box in vivo.

The functionality of the weak TyrR box, demonstrated by its necessary role in repression of akr in vivo and by its binding to TyrR in vitro as shown by EMSAs, was one of the most interesting findings of this study. In E. coli, TyrR binds the consensus

sequence TGTAAA-N6-TTTACA and requires the conserved G-N14-C pair (1). The atypical spacing between the weak box arms in the akr promoter and the double tandem

5’ arms suggests two possibilities for the spacing of the G-C pair: G-N10-C or G-N16-C.

There is little information regarding protein binding to DNA sequences with atypical spacing between palindromic half-sites. The consensus binding sequence for PrrA, a global transcription factor controlling ~25% of the genome in Rhodobacter sphaeroides

(48), comprises two inverted half-sites separated by a spacer region of variable length, ranging from 0 to 10 base pairs (49). Site directed mutagenesis coupled with promoter

fusions and EMSAs revealed that while a five base pair spacer is optimal, PrrA also interacts with sites with an 8 base pair spacer (50).

Genome-wide studies of transcription factor interactions with DNA have revealed that interactions with non-canonical sites, which were previously considered artifacts, are more common than previously thought and reflect true protein-DNA interactions.

Chromatin immunoprecipitation coupled with microarray analysis (ChIP-chip) performed with the LexA regulator of E. coli demonstrated its interaction in vivo with several promoter regions lacking a discernible LexA DNA binding consensus sequence, while these sites were bound extremely poorly when studied in vitro (51). ChIP-chip performed with the nitric oxide responsive NsrR regulator of E. coli revealed that the protein can recognize and bind to two types of binding sites: one comprised of two 11 base pair inverted repeats separated by a single nucleotide and the other contains only a single 11 base pair arm, with no discernible second inverted arm (52). While in some cases, interactions between transcription factors and non-canonical sites have no discernible effect on gene transcription (51, 53), we have shown that interaction between TyrR and the weak box results in the down-regulation of akr expression.

2.6 Acknowledgements

We would like to thank Julie Ryu for construction of plasmids pJQSIAPG and pJQSIPG, and initial testing of akr expression from integrants generated from these.

2.7 References

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2. Yang J, Ogawa Y, Camakaris H, Shimada T, Ishihama A, Pittard AJ. 2007. folA, a new member of the TyrR regulon in Escherichia coli K-12. J Bacteriol 189: 6080–6084. 3. Pittard AJ, Davidson BE. 1991. TyrR protein of Escherichia coli and its role as repressor and activator. Mol Microbiol 5: 1585–1592. 4. Andrews AE, Lawley B, Pittard AJ. 1991. Mutational analysis of repression and activation of the tyrP gene in Escherichia coli. J Bacteriol 173: 5068–5078. 5. Argaet VP, Wilson TJ, Davidson BE. 1994. Purification of the Escherichia coli regulatory protein TyrR and analysis of its interactions with ATP, tyrosine, phenylalanine, and tryptophan. J Biol Chem 269: 5171–5178. 6. Wilson TJ, Argaet VP, Howlett GJ, Davidson BE. 1995. Evidence for two aromatic amino acid-binding sites, one ATP-dependent and the other ATP- independent, in the Escherichia coli regulatory protein TyrR. Mol Microbiol 17: 483–492. 7. Wilson TJ, Maroudas P, Howlett GJ, Davidson BE. 1994. Ligand-induced self- association of the Escherichia coli regulatory protein TyrR. J Mol Biol 238: 309– 318. 8. Bailey MF, Davidson BE, Minton AP, Sawyer WH, Howlett GJ. 1996. The effect of self-association on the interaction of the Escherichia coli regulatory protein TyrR with DNA. J Mol Biol 263: 671–684. 9. Yang J, Pittard AJ. 1987. Molecular analysis of the regulatory region of the Escherichia coli K-12 tyrB gene. J Bacteriol 169: 4710–4715. 10. Baseggio N, Davies WD, Davidson BE. 1990. Identification of the promoter, operator, and 5’ and 3’ ends of the mRNA of the Escherichia coli K-12 gene aroG. J Bacteriol 172: 2547–2557. 11. Camakaris H, Pittard AJ. 1982. Autoregulation of the tyrR gene. J Bacteriol 150: 70–75. 12. Cornish EC, Argyropoulos VP, Pittard J, Davidson BE. 1986. Structure of the Escherichia coli K12 regulatory gene tyrR. Nucleotide sequence and sites of initiation of transcription and translation. J Biol Chem 261: 403–410. 13. Wang P, Yang J, Lawley B, Pittard AJ. 1997. Repression of the aroP gene of Escherichia coli involves activation of a divergent promoter. J Bacteriol 179: 4213–4218. 14. Wang P, Yang J, Pittard AJ. 1997. Promoters and transcripts associated with the aroP gene of Escherichia coli. J Bacteriol 179: 4206–4212. 15. Yang J, Wang P, Pittard AJ. 1999. Mechanism of repression of the aroP P2 promoter by the TyrR protein of Escherichia coli. J Bacteriol 181: 6411–6418. 16. Yang J, Camakaris H, Pittard AJ. 1996. In vitro transcriptional analysis of TyrR- mediated activation of the mtr and tyrP+3 promoters of Escherichia coli. J Bacteriol 178: 6389–6393. 17. Sarsero JP, Pittard AJ. 1991. Molecular analysis of the TyrR protein-mediated activation of mtr gene expression in Escherichia coli K-12. J Bacteriol 173: 7701– 7704.

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18. Heatwole VM, Somerville RL. 1991. Cloning, nucleotide sequence, and characterization of mtr, the structural gene for a tryptophan-specific permease of Escherichia coli K-12. J Bacteriol 173: 108–115. 19. Heatwole VM, Somerville RL. 1991. The tryptophan-specific permease gene, mtr, is differentially regulated by the tryptophan and tyrosine repressors in Escherichia coli K-12. J Bacteriol 173: 3601–3604. 20. Sarsero JP, Wookey PJ, Pittard AJ. 1991. Regulation of expression of the Escherichia coli K-12 mtr gene by TyrR protein and Trp repressor. J Bacteriol 173: 4133–4143. 21. Yang J, Murakami K, Camakaris H, Fujita N, Ishihama A, Pittard AJ. 1997. Amino acid residues in the alpha-subunit C-terminal domain of Escherichia coli RNA polymerase involved in activation of transcription from the mtr promoter. J Bacteriol 179: 6187–6191. 22. Song J, Jensen RA. 1996. PhhR, a divergently transcribed activator of the phenylalanine hydroxylase gene cluster of Pseudomonas aeruginosa. Mol Microbiol 22: 497–507. 23. Herrera MC, Duque E, Rodríguez-Herva JJ, Fernández-Escamilla AM, Ramos JL. 2010. Identification and characterization of the PhhR regulon in Pseudomonas putida. Environ Microbiol 12: 1427–1438. 24. Rodionov DA, Novichkov PS, Stavrovskaya ED, Rodionova IA, Li X, Kazanov MD. 2011. Comparative genomic reconstruction of transcriptional networks controlling central metabolism in the Shewanella genus. BMC Genomics 12: S3. doi:10.1186/1471-2164-12-S1-S3. 25. Ryu RJ, Patten CL. 2008. Aromatic amino acid-dependent expression of indole-3- pyruvate decarboxylase is regulated by TyrR in Enterobacter cloacae UW5. J Bacteriol 190: 7200–7208. 26. Patten CL, Blakney AJC, Coulson TJD. 2013. Activity, distribution and function of indole-3-acetic acid biosynthetic pathways in bacteria. Crit Rev Microbiol 39: 395–415. 27. Yanish-Perron C, Vieira J, Messing J. 1985. Improved M13 phage cloning vectors and host strains: nucleotide sequences of the M13mp18 and pUC19 vectors. Gene 33: 103-119. 28. Simon R, Puhler U, Priefer A. 1983. A broad host range mobilization system for in vivo genetic engineering: transposon mutagenesis in gram negative bacteria. Nat. Biotech 1: 784-791 29. Patten CL, Glick BR. 2002. Role of Pseudomonas putida indoleacetic acid in development of the host plant root system. Appl Environ Microbiol 68: 3795–3801. 30. Kovach ME, Elzer PH, Hill DS, Robertson GT, Farris MA, Roop RM Peterson KM. 2004. Four new derivatives of the broad-host-range cloning vector pBBR1MCS, carrying different antibiotic-resistance cassettes. Gene 1995;166: 175–176. 31. Chiu J. Site-directed, Ligase-Independent Mutagenesis (SLIM): a single-tube methodology approaching 100% efficiency in 4 h. Nucleic Acids Res 32: e174– e174.

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32. Cowie A, Cheng J, Sibley CD, Fong Y, Zaheer R, Patten CL. 2006. An integrated approach to functional genomics: construction of a novel reporter gene fusion library for Sinorhizobium meliloti. Appl Environ Microbiol 72: 7156–7167. 33. JMP, Version 7. SAS Institute Inc., Cary, NC, 1989-2007. 34. Klasens BI, Huthoff HT, Das AT, Jeeninga RE, Berkhout B. 1999. The effect of template RNA structure on elongation by HIV-1 reverse transcriptase. Biochim Biophys Acta 1444: 355–370. 35. Kröger C, Dillon SC, Cameron ADS, Papenfort K, Sivasankaran SK, Hokamp K, et al. 2012. The transcriptional landscape and small RNAs of Salmonella enterica serovar Typhimurium. Proc Natl Acad Sci USA 109: E1277–E1286. 36. Djordjevic M. 2011. Redefining Escherichia coli σ70 promoter elements: -15 motif as a complement of the -10 motif. J Bacteriol 193: 6305–6314. 37. Niedziela-Majka A. 2005. Escherichia coli RNA polymerase contacts outside the - 10 promoter element are not essential for promoter melting. J Biol Chem 280: 38219–38227. 38. Lawley B, Fujita N, Ishihama A, Pittard AJ. 1995. The TyrR protein of Escherichia coli is a class I transcription activator. J Bacteriol 177: 238–241. 39. Koga J, Adachi T, Hidaka H. 1992. Purification and characterization of indolepyruvate decarboxylase. A novel enzyme for indole-3-acetic acid biosynthesis in Enterobacter cloacae. J Biol Chem 267: 15823–15828. 40. Peer WA, Blakeslee JJ, Yang H, Murphy AS. 2011. Seven things we think we know about auxin transport. Mol Plant 4: 487–504. 41. Cummings SP. 2009. The application of plant growth promoting rhizobacteria (PGPR) in low input and organic cultivation of graminaceous crops; potential and problems. Environ Biotechnol 5: 43–50. 42. Yang J, Hwang JS, Camakaris H, Irawaty W, Ishihama A, Pittard AJ. 2004. Mode of action of the TyrR protein: repression and activation of the tyrP promoter of Escherichia coli. Mol Microbiol 52: 243–256. 43. Jez JM, Bennett MJ, Schlegel BP, Lewis M, Penning TM. 1997. Comparative anatomy of the aldo-keto reductase superfamily. Biochem J 326: 625–636. 44. Mindnich RD, Penning TM. 2009. Aldo-keto reductase (AKR) superfamily: genomics and annotation. Hum Genomics 3: 362–370. 45. Kilfoil PJ, Tipparaju SM, Barski OA, Bhatnagar A. 2013. Regulation of ion channels by pyridine nucleotides. Circ Res 112: 721–741. 46. Grant AW, Steel G, Waugh H, Ellis EM. 2003. A novel aldo-keto reductase from Escherichia coli can increase resistance to methylglyoxal toxicity. FEMS Microbiol Lett 218: 93–99. 47. Sawyer WH, Chan RY, Eccleston JF, Davidson BE, Samat SA, Yan Y. 2000. Distances between DNA and ATP binding sites in the TyrR-DNA complex. Biochemistry 39: 5653–5661. 48. Eraso JM, Roh JH, Zeng X, Callister SJ, Lipton MS, Kaplan S. 2008. Role of the global transcriptional regulator PrrA in Rhodobacter sphaeroides 2.4.1: Combined transcriptome and proteome analysis. J Bacteriol 190: 4831–4848. 49. Mao L. 2005. Combining microarray and genomic data to predict DNA binding motifs. Microbiology 151: 3197–3213. 102

50. Eraso JM, Kaplan S. 2009. Half-Site DNA sequence and spacing length contributions to PrrA binding to PrrA site 2 of RSP3361 in Rhodobacter sphaeroides 2.4.1. J Bacteriol 191: 4353–4364. 51. Wade JT. 2005. Genomic analysis of LexA binding reveals the permissive nature of the Escherichia coli genome and identifies unconventional target sites. Genes Dev 19: 2619–2630. 52. Partridge JD, Bodenmiller DM, Humphrys MS, Spiro S. 2009. NsrR targets in the Escherichia coli genome: new insights into DNA sequence requirements for binding and a role for NsrR in the regulation of motility. Mol Microbiol 73: 680– 694. 53. Wade JT, Struhl K, Busby SJW, Grainger DC. 2007. Genomic analysis of protein-DNA interactions in bacteria: insights into transcription and chromosome organization. Mol Microbiol 65: 21–26.

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Table 2.1. Bacterial strains and plasmids used in this study.

Strains or Plasmids Relevant characteristics Reference E. coli JM109 Cloning host; endA1 recA1 gyrA96 thiA hsdR17 relA1 supE44 ∆lac- (27) proAB F’ traD36 proAB laqIq ∆lacZ M15 S17-λ pir Cloning host; pir recA thi pro hsdR M RP4:2 Tc:Mu:Km Tn7 Tpr, (28) Strs M15 Expression host; E. coli K12 ∆lacZ ∆mtl pREP4, Strs Qiagen E. cloacae UW5 Wild-type strain (29) J35 UW5 tyrR::Tcr (25) Plasmids pGEM-T Easy Cloning vector, Ampr Promega pGEM-IP1 420 bp 5’-RACE ipdC fragment cloned into pGEM-T Easy, Ampr This study pGEM-AP1 280 bp 5’-RACE akr fragment cloned into pGEM-T Easy, Ampr This study pQEtyrR 1542 bp fragment encoding the E. cloacae UW5 tyrR gene cloned (25) into pQE-30, Ampr pJQSIPG pJQ200SK derivative; PipdC::uidA, Gmr (25) pJQSIAPG pJQ200SK derivative; Pakr::uidA, Gmr (25) pBBR1MCS-2 Cloning vector, Kmr (28) pBBRL pBBR1MCS-2, PlacZ deletion, Kmr This study pBBRLI 2.3 kb PipdC::uidA fragment from pJQSIPG cloned into pBBRL, This study Kmr pBBRLIW pBBRLI derivative, mutation of the TyrR weak box, Kmr This study pBBRLIS pBBRLI, mutated strong box, Kmr This study pBBRLIWS pBBRLLI, mutated weak and strong box, Kmr This study pBBRLISS pBBRLI, weak to strong box, Kmr This study pBBRLA 2.3 kb Pakr::uidA fragment from pJQSIAPG cloned into pBBRL, This study Kmr pBBRLAW pBBRLA, mutated weak box, Kmr This study pBBRLAS pBBRLA, mutated strong box, Kmr This study pBBRLAWS pBBRLA, mutated weak and strong box, Kmr This study pBBRLASS pBBRLA, weak to strong box, Kmr This study

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Table 2.2. PCR primers used in this study.

Primer Sequence 5ʹ-3ʹ IP1F GCCATGGCAGGAAATCTTC IPA2R GACGGTCCAGCAGGTAATG U2R TTCCACAGTTTTCGCGATCC AAP GGCCACGCGTCGACTAGTACGGGIIGGGIIGGGIIG AUAP GGCCACGCGTCGACTAGTAC AKR-GSP2 CGAAATTACGTTCGGCTGAG IPDC-GSP1 ACATGCTCGGCATAGCTG IPDC-GSP2 AACGCCGAATGTGGTCAG CF1 CCCATTCTGATGCCCTCTG CR1 GGTTCGCATAACAGGTGTCC DIG-CR1 DIG-GGTTCGCATAACAGGTGTCC WFT1 TGTATACGTTTACATTTACATGAAAAAAAAGAGCATAGCG WFS1 TGAAAAAAAAGAGCATAGCGCAGCC WRT1 TGTAAATGTAAACGTATACACTGATTTTTCCTTTCAGC WRS1 CTGATTTTTCCTTTCAGCGCCAGAGG WFT2 TATATACGTTTATATTTATATGAAAAAAAAGAGCATAGCGCAGCC WRT2 TATAAATATAAACGTATATACTGATTTTTCCTTTCAGCGCCAGA SFT1 TATAAAGCATTCTTTCTATGCCCTTCTTACGACCAATT SFS1 TGCCCTTCTTACGACCAATTCTGGA SRT1 TAGAAAGAATGCTTTATAAAAAGGCTGCGCTATGC SRS1 AAAAAGGCTGCGCTATGCTCTTTTTTTTCA

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Figure 2.1. The akr-ipdC regulatory region of E. cloacae UW5. (A) Relative location of the two TyrR binding sites located between the akr and ipdC genes. (B) Nucleotide sequence of the akr-ipdC intergenic region. TyrR boxes are in bold and nucleotides proposed to be essential are indicated with asterisks. The predicted -10, -35 elements and ribosome binding sites for both genes are underlined. The identified ipdC transcription start site, and predicted akr transcription start site are in bold underline. Arrows above the sequence indicate the akr and ipdC start codons. Arrows below the sequence indicate primer binding sites for CF1 and DIG-CR2. (C) Reporter gene expression plasmids driven by the ipdC (I) or akr (A) promoters, with the corresponding mutation or insertions indicated in lowercase compared to the TyrR box consensus sequence.

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Figure 2.2. The expression of akr and ipdC is modulated by TyrR. Expression from (A) ipdC and (B) akr in the presence or absence of aromatic amino acids in wild-type E. cloacae UW5 (grey) and tyrR null mutant E. cloacae J35 (black). Cells were assayed for β-glucuronidase activity in both logarithmic and stationary phases of growth. Error bars represent the standard error of the means of three independent replicates. Statistically significant differences of p < 0.05 are indicated by lowercase letters in logarithmic phase, and uppercase letters in stationary phase.

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Figure 2.3. Role of the TyrR boxes in akr-ipdC gene regulation. Expression from (A) ipdC and (B) akr promoter mutants in wild-type E. cloacae UW5 (grey) and tyrR null mutant E. cloacae J35 (black) in tryptophan-supplemented M9 minimal media. Cells were assayed for β-glucuronidase activity in both logarithmic and stationary phases of growth. Error bars represent the standard error of means of three independent replicates. Statistically significant differences of p < 0.05 are indicated by lowercase letters in logarithmic phase, and uppercase letters in stationary phase.

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Figure 2.4. Binding of TyrR to the TyrR boxes within the akr-ipdC intergenic region. DIG-labeled DNA probes correspond to the akr-ipdC intergenic sequence of the template plasmid from which they were generated, wild-type sequence (Lanes 1-3); weak box mutant (Lanes 4-6); strong box mutant (Lanes 7-9); double weak and strong box mutant (Lanes 10-12) and double strong boxes (lanes 13-15). DNA probes were incubated with either no TyrR (Lanes 1, 4, 7, 10 and 13) or increasing concentrations of TyrR (87 nM: Lanes 2, 5, 8, 11 and 14; 877 nM: Lanes 3, 6, 9, 12 and 15). Arrows indicate the positions of free DNA (F) and the two resolved TyrR-DNA complexes (I, II).

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Figure 2.5. Conservation of the akr-ipdC genes and the two TyrR boxes across closely related bacteria. (A) Genomic context of the akr-ipdC genes in closely related Enterobacteriaceae, Homologous genes are indicated by the same colour, white indicates non-homologous genes. (B) Species with an akr-ipdC gene pair retain the weak and strong TyrR boxes. The second palindromic arm of the E. cloacae UW5 weak TyrR box is not conserved across species.

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2.9 Supplemental Information

2.9.1 List of supplemental figures

Supplemental Figure 2.1. Expression of ipdC in promoter mutants by TyrR in response to aromatic amino acids...... 112

Supplemental Figure 2.2. Expression of akr promoter mutants by TyrR in response to aromatic amino acids...... 113

Supplemental Figure 2.3. Binding of TyrR to the akr-ipdC intergenic region is concentration dependant...... 114

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Supplemental Figure 2.1. Expression of ipdC in promoter mutants by TyrR in response to aromatic amino acids. Expression from ipdC promoter mutants in M9 minimal medium without amino acid supplements (A) and in the presence of phenylalanine (B), or tyrosine (C) in wild-type E. cloacae UW5 (grey) and tyrR null mutant E. cloacae J35 (black). Cells were assayed for β-glucuronidase activity in both logarithmic and stationary phases of growth. Error bars represent the standard error of the means of three independent replicates. Statistically significant differences of p < 0.05 are indicated by lowercase letters in logarithmic phase, and uppercase letters in stationary phase.

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Supplemental Figure 2.2. Expression of akr promoter mutants by TyrR in response to aromatic amino acids. Expression from akr promoter mutants in M9 minimal medium without amino acid supplements (A) and in the presence of phenylalanine (B), or tyrosine (C) in wild-type E. cloacae UW5 (grey) and tyrR null mutant E. cloacae J35 (black). Cells were assayed for β-glucuronidase activity in both logarithmic and stationary phases of growth. Error bars represent the standard error of the means of three independent replicates. Statistically significant differences of p < 0.05 are indicated by lowercase letters in logarithmic phase, and uppercase letters in stationary phase.

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Supplemental Figure 2.3. Binding of TyrR to the akr-ipdC intergenic region is concentration dependant. Wild-type DIG-labelled DNA probe was incubated with increasing amounts of TyrR-His6 (0 mM, 87 nM, 438 nM, 877 nM 1750 nM) in the absence of aromatic amino acids (A), 1 mM tryptophan (B), phenylalanine (C) or tyrosine (D). Competition assays were performed with 10-fold molar excess of unlabeled- probe over DIG-labelled probe; rightmost lane. Positions of free DNA (F) and the two resolved TyrR-His6-DNA complexes (I, II) are indicated.

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Chapter 3 Complete genome sequence of Enterobacter cloacae UW5, a rhizobacterium capable of high levels of indole-3-acetic acid productiona

Thomas J.D. Coulson1, Cheryl L. Patten1*

a Published in Genome Announcements, August, 2015, 3(4): e00843-15. doi:10.1128/genomeA.00843-15

PMID: 26251488

1Department of Biology, University of New Brunswick, Fredericton New Brunswick,

Canada, E3B5A3

*Corresponding Author [email protected]

3.1 Abstract

We report the complete genome sequence of Enterobacter cloacae UW5, and indole-3-acetic acid producing rhizobacterium originally isolated from the rhizosphere of grass. The 4.9 Mbp genome has a G-C content of 54% and contains 4,496 protein coding sequences.

3.2 Genome announcement

Members of the Enterobacter genus are Gram-negative facultative anaerobic bacteria commonly found inhabiting the rhizosphere of plants, and may also be members of the intestinal microflora of humans. Enterobacter cloacae strains are often endophytes capable of biocontrol of plant infectious diseases, or production of plant growth-

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stimulating compounds (1). E. cloacae UW5 was originally isolated from the rhizosphere of reeds in Waterloo, Ontario (2,3), and has been shown to produce high levels of indole-

3-acetic acid (IAA)(4), a signalling molecule which acts as a bacterial stress response regulator, antimicrobial metabolite, and plant growth-promoting stimulant (5). Regulation of IAA by the global regulatory protein TyrR has been characterized in this strain (4,6).

The genome of E. cloacae UW5 was sequenced to provide further insight into the TyrR regulon and the function of IAA in bacterial physiology and bacterial-plant interactions.

Genomic DNA was extracted from an overnight culture grown at 30°C, using the

Wizard Genomic DNA Purification kit (Promega). A 100 bp paired end TruSeq library was prepared with an average fragment size of 444 bp and sequenced in a single Illumina

HiSeq 2500 lane (McGill University and Génome Québec Innovation Center, Montreal,

Canada). A total of 251,715,967 reads were generated totalling over 51 billion bases.

Seqtk (https://github.com/lh3/seqtk) was used to randomly extract a subset of 10 million forward and reverse paired reads which were then trimmed using Trimmomatic (7) to remove adapters and low quality sequences. The resulting reads were assessed for sequence quality using FastQC v.0.11.2 (8) before assembly. Genome assembly was performed using a combination of de novo assembly and reference guided read mapping to a closely related reference genome. An optimal kmer of 71 was chosen using

VelvetOptimizer, which was used for de novo assembly with Velvet v.1.2.10 (9). A total of 83 contigs were generated, with an average of 330-fold coverage and an N50 length of

2,836,112 bp (largest contig measuring 2,836,112 bp). The same read data were then mapped to the E. cloacae WSU1 reference genome (NC_016514) (10) using CLC

Genomics Workbench v.8.0.1 (Qiagen) at both low (80% identity and 50% length 116

fraction) and high (90% identity and 90% length fraction) specificity, and the consensus sequences extracted to produce the reference guided contigs. Contigs from both de novo and reference guided assemblies were then joined using the CLC Microbial Genome

Finishing module to merge overlapping sequences into a single contig. Genome annotation was performed using the NCBI Prokaryotic Genome Annotation Pipeline

(PGAP) (http://www.ncbi.nlm.nih.gov/genome/annotation_prok/).

The E. cloacae UW5 genome consists of a single chromosome that is 4,904,981 bp in length with a G+C% of 54.46%. The genome contains 4,496 genes assigned with protein encoding function, 77 tRNA genes and 25 rRNA genes organized into eight rRNA operons.

3.3 Nucleotide sequence and accession number.

The complete E. cloacae UW5 genome has been deposited in Genbank under the accession number CP011798. The version reported in this paper is the first version.

3.4 Acknowledgments

This work was supported by a Discovery grant to Cheryl L. Patten and a Post- graduate Scholarship to Thomas J.D. Coulson from the National Sciences and

Engineering Research Council of Canada (NSERC).

3.5 References

1. Nguyen MT, Ranamukhaarachchi SL. 2010. Soil-borne antagonists for biological control of bacterial wilt disease caused by Ralstonia solanacearum in tomato and pepper. J. Plant. Pathol. 92:395-406. 2. Glick BR, Karaturovic DM, Newell, PC. 1995. A novel procedure for rapid isolation of plant growth promoting pseudomonads. Can. J. Microbiol. 41:533-536.

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3. Shah S, Li J, Moffatt BM, Glick BR. 1997. ACC deaminase genes from plant growth-promoting bacteria, p 320-324. In Ogoshi A, Kobayashi K, Homma Y, Kodama F, Kondo N, Akino S (ed), Plant growth promoting bacteria: present status and future prospects. Organization for Economic Cooperation and Development, Paris. 4. Ryu RJ, Patten CL. 2008. Aromatic amino acid-dependent expression of indole-3- pyruvate decarboxylase is regulated by TyrR in Enterobacter cloacae UW5. J. Bacteriol. 190:7200-7208. 5. Patten CL, Blakney AJC, Coulson TJD. 2013. Activity, distribution and function of indole-3-acetic acid biosynthetic pathways in bacteria. Crit. Rev. Microbiol. 39:395-415. 6. Coulson TJD, Patten CL. 2015. The TyrR transcription factor regulates the divergent akr-ipdC operons of Enterobacter cloacae UW5. PLoS One. 26:e0121241. doi:10.1371/journal.pone.0121241. 7. Bolger AM, Lohse M, Usadel B. 2014. Trimmomatic: A flexible trimmer for Illumina sequence data. Bioinformatics. Btu170. 8. Andrews S. 2010. FastQC: A quality control tool for high throughput sequence data. Available online at: http://www.bioinformatics.babraham.ac.uk/projects/fastqc 9. Zerbino DR, Birney E. 2008. Velvet: Algorithms for de novo short read assembly using de Bruijn graphs. Genome Res. 18:821-829. 10. Humann JL, Wildung M, Cheng CH, Lee T, Stewart JE, Drew JC, Triplett EW, Main D, Schroeder BK. 2011. Complete genome of the onion pathogen Enterobacter cloacae EcWSU1. Stand. Genomic Sci. 5:279-286.

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Chapter 4 Characterization of the TyrR regulon in the rhizosphere bacterium Enterobacter cloacae UW5 reveals overlap with the CpxR envelope stress responsea

Thomas J.D. Coulson1, René M. Malenfant1 and Cheryl L. Patten1*

a Submitted, November, 2018

1Department of Biology, University of New Brunswick, Fredericton New Brunswick,

Canada, E3B5A3

*Corresponding Author [email protected]

4.1 Abstract

The TyrR transcription factor controls the expression of genes for the uptake and biosynthesis of aromatic amino acids in Escherichia coli. In the plant-associated and clinically significant proteobacterium Enterobacter cloacae UW5, the TyrR orthologue regulates genes that encode enzymes for synthesis of the plant hormone indole-3-acetic acid and gluconeogenesis, indicating a broader function for the transcription factor. This study aimed to delineate the TyrR regulon of E. cloacae by comparing the transcriptomes of the wild-type and a tyrR deletion strain. Our data show that TyrR regulates the expression of over 150 genes both positively and negatively. TyrR downregulates expression of envelope stress response regulators CpxR and CpxP through interaction with an atypical DNA binding site in the intergenic region between divergently 119

transcribed cpxP and cpxR. Repression is independent of aromatic amino acids, although repression of cpxP is alleviated by tyrosine. The methyltransferase gene dmpM, possibly involved in tyrosine metabolism, is strongly activated in the presence of tyrosine and phenylalanine by TyrR binding to its promoter region. TyrR also regulates expression of genes for aromatic catabolism, motility, and anaerobic respiration. Our findings suggest that the E. cloacae TyrR regulon has diverged from that of E. coli to include genes for survival in the diverse environments that this bacterium inhabits, and illustrate the expansion and plasticity of transcription factor regulons.

4.1.2 Importance

Genome-wide RNA sequencing revealed a broader regulatory role for the TyrR transcription factor in the ecologically versatile bacterium Enterobacter cloacae beyond aromatic amino acid synthesis and transport that constitute the TyrR regulon of E. coli. In

E. cloacae, a plant symbiont and human gut commensal, the TyrR regulon is expanded to include genes that are beneficial for plant interactions and response to stresses.

Identification of the genes regulated by TyrR provides insight into the mechanisms by which the bacterium adapts to its environment.

4.2 Introduction

Expression of genes for aromatic amino acid biosynthesis and uptake in E. coli is regulated by the transcription factor TyrR, which represses or activates nine separate transcriptional units encompassing 12 genes (1, 2). TyrR controls the expression of genes of the shikimate pathway (aroF, aroG, aroL) and subsequent production of the aromatic amino acids phenylalanine and tyrosine via chorismate (tyrA, tyrB), in addition to genes 120

encoding several aromatic amino acid transporters (aroP, mtr, tyrP) and itself (tyrR) (1).

TyrR also activates the folA gene for biosynthesis of folic acid, which is derived from chorismate (2). TyrR recognizes and binds to DNA at sites related to the consensus sequence TGTAAA-N6-TTTACA that overlap or are adjacent to sigma-70 promoters;

DNA footprinting studies indicate that TyrR protection extends an additional two base pairs on either side of the consensus sequence (3-5). These DNA binding sites, known as

TyrR boxes, have varying degrees of agreement with the consensus sequence, which affects the affinity of TyrR binding to the sequence (3, 5). Generally, strong TyrR boxes have at least ten nucleotide matches to the consensus, a high degree of sequence symmetry, and an AT-rich spacer sequence, while weak boxes have fewer than ten matching nucleotides and less symmetry (6, 7). A key feature of functional TyrR boxes is the invariable G-N14-C bases; changes to either nucleotide result in a loss of TyrR binding (1). Each of the three aromatic amino acids, tryptophan, phenylalanine and tyrosine, function as cofactors for TyrR, modulating its regulatory action such that each regulon member is uniquely controlled. The aromatic amino acids bind TyrR at two sites: one ATP independent binding site for tryptophan and phenylalanine, and a second ATP- dependent tyrosine binding site (1, 5). TyrR exists as a dimer in the absence of cofactors

(8), but in the presence of tyrosine and ATP it undergoes further polymerization to form a hexamer, which may then interact with multiple TyrR boxes simultaneously (9, 10).

Phenylalanine can facilitate the cooperative binding of TyrR dimers across multiple boxes through dimer dimer interactions (11, 12).

TyrR functions as both a transcriptional repressor and activator, in some cases exhibiting both capacities at the same promoter. Promoters repressed by TyrR in the 121

presence of tyrosine contain at least two adjacent TyrR boxes, one strong and one weak

(7). The weak box is typically closer to, or overlapping with, the core promoter (-35 or -

10 elements), and cofactor-mediated binding at this low-affinity site requires an adjacent strong box. Repression may involve the exclusion of RNA polymerase from the promoter, as is the case for tyrP, or the inhibition of subsequent steps of transcription such as open complex formation or promoter escape, as for tyrB (13-16). A strong box located 42 base pairs upstream from the -35 sequence of the alternate P3 promoter of aroP recruits RNA polymerase; however, the resulting transcriptional complex is nonproductive and inhibits the activation of the primary aroP P1 promoter located on the opposite DNA strand (17-19). TyrR activation of transcription occurs through direct interactions with the RNA polymerase alpha subunit carboxy terminal domain, whereby it recruits RNA polymerase to the promoter or otherwise stabilizes its interaction with the

DNA (1). This requires TyrR to be positioned upstream of the -35 promoter element, and on the same face of DNA as to allow direct contact between the N-terminal domain of

TyrR and RNA polymerase (20, 21). TyrR activates expression of mtr when bound to two adjacent boxes, one strong and one weak, upstream of the promoter as a pair of dimers with phenylalanine, or to the strong box only as a hexamer with tyrosine (22). The gene folA, encoding dihydrofolAte reductase, is activated by TyrR binding two sites centered

76 and 155 base pairs upstream of the -35 sequence, and requires integration host factor

(IHF)-mediated DNA bending to facilitate contact with RNA polymerase (2). A further requirement for TyrR-mediated gene activation is an imperfect promoter, as seen with the

GC rich -10 elements of the tyrP, mtr and aroPP3 genes (1, 23). In the presence of tyrosine, hexameric TyrR binds to the weak box that overlaps the -35 element in the tyrP 122

promoter and thereby represses expression; however, with phenylalanine TyrR binds the upstream strong box alone as a dimer to activate tyrP expression. In all instances of TyrR regulation, the location of the TyrR boxes, and the spacing between them and the promoter is a crucial determinant for the mode of transcriptional control exhibited by

TyrR, which is further modulated by the oligomerization status of the protein influenced by the presence of aromatic amino acids (12, 14).

The TyrR regulon has been extensively studied in E. coli; however, in other gammaproteobacteria additional TyrR-regulated genes have been identified that encode functions accessory to or beyond aromatic amino acid biosynthesis and uptake. The tpl gene of Citrobacter freunii and Erwinia herbicola, encoding tyrosine phenol lyase for tyrosine catabolism, is upregulated by cooperative TyrR binding across three boxes in conjunction with IHF- and CRP-mediated DNA bending (24-26). In the entomopathogenic bacterium Photorhabdus luminescens expression of the phenylalanine- ammonium lyase gene sltA is controlled by TyrR, and required for production of a stilbene antibiotic (27). The acid-inducible genes aniC and hyaB of Salmonella typhimurium, encoding a arginine:agmatine symporter and hydrogenlyase subunit, respectively, are activated by TyrR with tyrosine under acidic growth conditions (28).

The Pseudomonas TyrR homologue, PhhR, controls genes for metabolism of aromatic compounds and amino acid catabolism, in addition to aromatic amino acid biosynthesis

(29-31). Rodionov et al. (2011) inferred from reconstruction of transcriptional networks that TyrR has undergone regulon expansion in the Shewanella genus, controlling degradation pathways for phenylalanine (phhAB), tyrosine (fahA-maiA), branched chain amino acids (ldh, brnQ, liu, ivd, blk), proline (putA), and oligopeptides, in addition to the 123

glyoxylate cycle for carbohydrate synthesis (aceBA) (32). Furthermore, there is evidence from genomic SELEX and bioinformatic analysis that there may be more TyrR regulon targets in E. coli, however direct regulation of these putative genes is yet to be demonstrated (1, 2). The differences in TyrR regulated genes among these bacteria may reflect genus- and species-specific adaptations required to inhabit different environments.

Enterobacter cloacae is a gram-negative, plant growth-promoting rhizobacterium, as well as a clinically relevant opportunistic pathogen (33, 34). Previous work has shown that E. cloacae UW5 synthesizes high levels of the plant growth hormone indole-3-acetic acid (IAA) when tryptophan is supplied exogenously (33). In this bacterium, IAA is produced from tryptophan via the indolepyruvate pathway, which is dependent on the rate limiting decarboxylation of indole-3-pyruvate to indole-3-acetaldehyde by indolepyruvate decarboxylase, encoded by ipdC. The expression of ipdC is wholly dependent on TyrR, and is upregulated by aromatic amino acids, especially phenylalanine

(33, 35). Transcriptional activation of ipdC occurs via TyrR binding to a promoter- proximal strong box centered 33 bp upstream from the -35 element. A second TyrR box further upstream overlaps the -10 element of the divergently transcribed akr gene promoter, and represses its expression (35). This second weak box has atypical spacing of the left and right palindromic arms, which are separated by two or eight base pairs rather than the canonical six, yet binding by TyrR has been demonstrated in vitro (35). The homologous gene in E. coli, gpr, encodes an L-glyceraldehyde-3-phosphate reductase, which along with L-glyceraldehyde dehydrogenase generates dihydroxyacetone phosphate (DHAP) for gluconeogenesis (36). The novel finding that in E. cloacae TyrR controls expression of an enzyme for tryptophan catabolism, in addition to another 124

involved in gluconeogenesis, indicates that its function is more diverse compared to E. coli TyrR. In this study we aimed to delineate the TyrR regulon in E. cloacae UW5 by comparing the genome wide transcriptional profiles in the wild type and a tyrR mutant strain. We found that TyrR controls an expanded regulon in E. cloacae, significantly altering expression of genes and operons for chemotaxis, anaerobic respiration and aromatic catabolism, and directly activating transcription of a methyltransferase.

Moreover, TyrR mediates changes in the Cpx stress envelope response pathway by directly repressing transcription of the cpx operon through interactions with a non- canonical TyrR box.

4.3 Materials and Methods

4.3.1 Bacterial strains and growth conditions.

The bacterial strains and plasmids used in this study are listed in Supplementary

Table S4.1. Bacteria were routinely cultured in Luria-Bertani (LB) broth, with shaking at

250 rpm and incubated at 37℃ for E. coli or 30℃ for E. cloacae. To measure TyrR- responsive gene expression, E. cloacae was grown in M9 minimal media supplemented with 1 mM of L-tryptophan, L-phenylalanine, or L-tyrosine. Antibiotics were used at the following concentrations: 100 µg/mL ampicillin, 25 µg/mL kanamycin, 25 µg/mL gentamicin or 25 µg/mL chloramphenicol.

4.3.2 Construction of a tyrR deletion mutant.

The entire tyrR coding sequence was removed from the E. cloacae chromosome using flippase (Flp)-Flp recognition target (FRT) recombination thereby avoiding the

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introduction of antibiotic resistance genes (37). The tyrR sequence was initially replaced with a kanamycin resistance gene (KmR) flanked by FRT sites via homologues recombination, before its subsequent removal with Flp. The upstream and downstream regions of the genome flanking the tyrR sequence were amplified from wild-type E. cloacae genomic DNA using primers TUF-PstI and TUR-XbaI, and TDF-XbaI and TDR-

SacI, respectively (Supplementary Table S4.2). Purified PCR products were used as templates for splicing by overlap extension PCR with primers TUF-PstI and TDR-SacI to give a 1.5-kb fragment which was subsequently ligated into pGEM®-T Easy (Promega) generating plasmid pTUD. Plasmid pTUD was transformed into E. coli JM109 and transformants selected on ampicillin. The FRT-flanked kanamycin resistance gene (KmR) was amplified from plasmid pKD4 using primers PKD1 and PKD2, and ligated into

XbaI-digested pTUD, resulting in plasmid pTUD-Km which was transformed into E. coli

JM109. The 3-kb TUD-Km cassette was excised from pTUD-Km via NotI digestion and ligated into similarly digested pJQ200SK, creating pJQTUD-Km. pJQTUD-Km was transformed into E. coli S17-1λpir and subsequently conjugated into wild-type E. cloacae. Single recombinants were identified by gentamicin and kanamycin resistance, sucrose sensitivity, and PCR amplification of the KmR gene with primers PKDP1 and

PKDP2. Subsequently, the pJQ200SK backbone was selected against by growing single recombinants overnight in LB broth with 10% sucrose, and plating on LB kanamycin with 10% sucrose. Double recombinants were screened for gentamicin sensitivity and kanamycin resistance, and verified by PCR amplification with primers TUF and TUR, and TDF and TDR, which amplify outside the region used for homologous recombination. The KmR gene was removed by transforming calcium chloride competent 126

E. cloacae tyrR::KmR cells with plasmid pCP20 encoding Flp recombinase, and selecting on chloramphenicol-containing medium. Positive transformants were restreaked and grown overnight in LB broth at 37℃ to induce expression of the Flp recombinase and inhibit pCP20 replication. Loss of the KmR gene was verified by testing for kanamycin sensitivity, PCR amplification with primers TUF and TDR, and sequencing of the amplified product. Verified tyrR mutants were stored at -80℃.

4.3.3 IAA quantification assay.

Because IAA production in E. cloacae is wholly dependent on TyrR mediated expression of the ipdC gene, colorimetric quantification of IAA was used to validate the loss of TyrR activity. E. cloacae wild-type and tyrR mutant strains were grown overnight in LB broth, pelleted, and washed twice with 1x M9 salts (Difco), before inoculating

1:100 into M9 minimal media or M9 supplemented with 1 mM tryptophan. IAA production was quantified after 24 hours of incubation at 30℃ by the method of Gordon and Weber (1951) (38). Culture turbidity at OD600nm was recorded before pelleting 1 ml of cells and transferring 40 µl of culture supernatant to a 96-well plate containing 160 µl

Salkowski’s reagent (36% H2SO4, 10 mM FeCl3). Reactions were incubated at room temperature for 20 minutes before quantifying absorbance at OD535nm on a SpectraMax

M5 plate reader (Molecular Devices). Total IAA was determined by extrapolating from a standard curve of known concentration and normalized to culture turbidity.

4.3.4 RNA isolation and sequencing.

Overnight cultures from single colonies of wild-type and tyrR mutant E. cloacae were inoculated 1:100 into 3 ml fresh LB in quadruplicate and incubated at 30℃, 127

250 rpm for ~2.5 hrs until reaching mid logarithmic phase of growth (OD600 = 0.6-0.9). A total of 0.5 ml of cell suspension were then transferred to a 2 ml tube containing 1 ml

RNAprotect Bacterial Reagent (Qiagen), vortexed, and incubated 5 minutes at room temperature before pelleting by centrifugation. Cell pellets were stored at -20℃ until further use. Frozen cell pellets were thawed on ice and resuspended with 15 mg/ml lysozyme and 15 mg/ml Proteinase K in 200 µl TE buffer to lyse cells for 10 min at 4℃ with periodic vortexing. Total RNA was then extracted using the RNeasy Mini Kit

(Qiagen, Hilden, Germany) according to the manufacturer’s instructions. RNA quality and quantity were measured on a SpectraMax M5 plate reader (Molecular Devices) and visualized by agarose gel electrophoresis to ensure sample integrity. Contaminating genomic DNA was removed by treating 5 µg total RNA in a 50 µl reaction containing

5 units Amplification grade DNaseI (Invitrogen) and 5 µl 10x DNaseI digestion buffer and incubating at 37℃ for 60 minutes. The reaction was terminated by addition of 1 mM final concentration ethylenediaminetetraacetic acid (EDTA) and heating at 70℃ for

10 minutes. The DNase-treated RNA was purified using the Qiagen MinElute Cleanup

Kit (Qiagen) and eluted into 30 µl RNase-free water. DNA removal was verified by performing quantitative real-time PCR of the ipdC gene on each DNase-treated RNA sample using primers ICRTIF and ICRTIR (Supplementary Table S4.2). Samples that yielded no amplicons or Cp values greater than 30 were considered acceptable for library preparation. The final quality and quantity of RNA samples were measured on an Agilent

Bioanalyzer-RNA 6000 Pico chip (Agilent Technologies, CA, USA). Construction of

RNA-seq libraries and sequencing were performed by the Genome Quebec Innovation

Centre (McGill, Montreal Quebec). Ribosomal RNA was removed using the Ribo-Zero 128

rRNA Removal Kit for Gram Negative Bacteria (Illumina, CA, USA). The resulting enriched mRNA was used to construct a 100 bp paired-end stranded Illumina library using the TruSeq RNA Library Prep Kit v2 (Illumina) following manufacturer’s instructions (Illumina). The libraries were individually barcoded and sequenced together on a single Illumina HiSeq 2000 lane.

4.3.5 RNA-Seq data analysis.

Sequence data was retrieved from Genome Quebec servers and inspected for overall quality using FastQC v0.11.2 (S. Andrews, http:// www.bioinformatics.babraham.ac.uk/projects/fastqc/). Low-quality reads and contaminating Illumina TruSeq adapters were removed using Trimmomatic v0.3 (39) with a sliding window size of 4 bp, cutting sequences when the average Phred score dropped below 15. Surviving paired reads were mapped to the E. cloacae UW5 reference genome (Genbank Accession number CP011798 (40)) using bwa v0.7.9-r789 (41) as single fragments with a maximum mismatch of 1%. Read alignments were sorted, indexed and converted to BAM format with SAMtools v0.1.19-44428cd (42). Sample read alignments were visualised using the Integrative Genomics Viewer v2.3.32 (43) to ensure correct mapping of reads to genomic features. The number of reads mapped to each genomic feature were calculated using the default parameters of HTSeq-count v

0.6.1.p1 (44). Genes with significantly different levels of expression between wild-type and tyrR mutant strains were called using DESeq2 v1.12.4 (45). Genes were considered to be differentially expressed when there was at least a four-fold change (i.e., |log2-fold

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change| > 2) in expression with a false discovery rate threshold (FDR) adjusted p-value

(padj) of ≤ 0.001 as determined by the Benjamini-Hochberg correction method (45).

4.3.6 Pathway enrichment analysis.

Gene ontology (GO) terms were mapped to the annotated E. cloacae UW5 genome using Blast2GO (46-48). R version 3.4.3 (49) was used for statistical testing and plotting of data. The Bioconductor package goseq v1.30.0 (50) was used to identify over and under-represented GO terms present in the differentially-expressed dataset applying the Wallenius distribution approximation, and significantly enriched terms were determined by applying an FDR cutoff of 0.05 using the Benjamini-Hochberg adjustment. REVIGO was used to visualize the distribution of GO terms according to the major GO classifications (biological processes, cellular components, molecular function).

Coverage plots for mapped RNA-seq data were generated with the Bioconductor package

Gviz 1.22.3 (51).

4.3.7 Reverse transcriptase-quantitative-PCR.

To quantify target mRNAs by PCR, independent cultures of E. cloacae wild-type and tyrR mutant strains were grown (in triplicate) in LB broth or M9 minimal media supplemented without and with 1 mM of each aromatic amino acid. RNA was purified, quantified and treated with DNaseI to remove genomic DNA as described above for

RNA-seq sample preparation. cDNA was generated by mixing 100 ng DNaseI treated

RNA, 20 pmol of gene specific reverse primer (Supplementary Table S4.2) and 10 nmol dNTPs in a final volume of 13 µl. The reaction was incubated at 65℃ for 5 min before placing on ice for 1 min. To each cDNA reaction were added 4 µl First-Strand Synthesis 130

buffer (Invitrogen), 1 µl 0.1 M dithiothreitol, 1 µl RNaseOUT (Invitrogen), and 1 µl

SuperScript III Reverse Transcriptase (RT) (Invitrogen), and incubated at 55℃ for 1 hr.

Controls without reverse transcriptase were prepared in identical reactions. cDNA synthesis was stopped by heating at 70℃ for 10 min, and samples were stored at -20℃.

Duplicate real-time quantitative PCR reactions were performed with each cDNA in 25 µl reactions containing the following components: 200 nM each forward and reverse primer (Supplementary Table S4.2), 12.5 µl Platinum Taq SYBR Green Master

Mix (Invitrogen) and 4 µl cDNA corresponding to 100 ng total RNA. RT-qPCR amplification was performed on a Rotor-Gene 6000 thermocycler (Qiagen) with the following cycling conditions: initial denaturation at 98℃ for 2 minutes, followed by 40 cycles of denaturation at 98℃ for 10 seconds, primer annealing at 51℃ (aroG, proP);

57℃ (recA, cpxP, cpxR); or 61℃ (yagU, eco, dmpM) for 15 seconds, and elongation at

72℃ for 20 seconds with acquisition of the fluorescent signal. A melt-curve analysis was performed at the completion of each run, by raising temperature from 60℃ to 95℃ in

1℃ increments every 5 seconds with acquisition of fluorescent signal. The raw data were exported into LinRegPCR to determine individual reaction efficiencies and calculate corresponding Cp values (52). These values were imported into REST (Relative

Expression Software Tool) to determine fold expression against a normalized reference gene (recA) (53) using the ΔΔCt method (54, 55).

4.3.8 Purification of His6x-TyrR and electrophoretic mobility shift assays.

Purification of His6x-TyrR was performed as previously described (35). Briefly, overnight cultures of E. coli A-118 were subcultured 1:100 into 50 ml fresh LB medium

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and grown at 37℃, 250 rpm, for 4 hrs until reaching a cell density of OD600 = 0.4.

Expression of His6x-TyrR from plasmid pQEtyrR was induced with 1 mM of isopropyl β-

D-1-thiogalactopyranoside (IPTG) and cells were incubated a further 4 hours before harvesting by centrifugation. Cell pellets were stored at -20℃ until further use. Cell pellets were thawed on ice and resuspended in lysis buffer (50 mM NaH2PO4, 300 mM

NaCl, 10 mM imidazole, pH 8.0) with 1 mg/ml lysozyme (Sigma-Aldrich) for

30 minutes. Afterwards, cells were fully lysed while on ice with six 10-second bursts at

26 watts using a Sonic dismembrator model 100 (Fisher). His6x-TyrR was purified using

Ni-NTA resin (Qiagen) according to the manufacturers’ protocol for protein purification under native conditions, with the following exception: wash buffer 1: 50 mM NaH2PO4,

300 mM NaCl, 20 mM imidazole [pH 8.0]; wash buffer 2: 50 mM NaH2PO4, 300 mM

NaCl, 20 mM imidazole, 10% glycerol [pH 8.0]. Purified protein was eluted four times with 0.5 ml EB (50 mM NaH2PO4, 300 mM NaCl, 250 mM imidazole, 8% glycerol [pH

8.0]) and stored in aliquots at -80℃.

The cpx and dmpM promoters bearing either the wild-type sequence or containing specific point mutations within putative TyrR boxes were synthesised by Integrated DNA

Technologies (IDT, Coralville, IA) custom gene synthesis and provided on sequence- verified plasmids (carried on plasmid pUCIDT-AMP) (Supplementary Table S4.3).

These plasmids were used as direct PCR templates with primers cpxP-F and cpx-R-FAM or dmpM-F and dmpM-R-FAM (Supplementary Table S4.2) to amplify the cpxP, cpxR and dmpM promoters, respectively, while incorporating the fluorescent molecule 6- carboxyfluorescein (6-FAM) into the DNA product. Labelled PCR products were purified after running on a 4% native polyacrylamide gel using the Qiagen PCR 132

Purification Kit (Qiagen). All steps were performed while minimizing exposure to direct light and probes were stored at -20℃. Binding reactions were performed as described previously (35). Briefly, His6x-TyrR was added to 5 µl 5X binding buffer (50 mM Tris-Cl

[pH 7.5], 250 mM NaCl, 5 mM MgCl2, 20% glycerol, 2.5 mM DTT, 2.5 mM EDTA),

1 µg poly[d(I-C)] (non-specific competitor) and 0.1 mM aromatic amino acid or 0.1 mM

ATP when specified, and was incubated at room temperature for 5 minutes. Afterwards,

100 ng 6-FAM labelled DNA was added to the reaction and incubate 30 min at room temperature. Reactions were stopped with 5 µl loading buffer (0.25x TBE, 60% glycerol,

0.001% bromophenol blue) and loaded onto pre-run 6% native polyacrylamide gels containing 1 mM of the respective aromatic amino acid or 0.1 mM ATP for tyrosine reactions. Protein-DNA complexes were resolved at 100V, at 4℃ for ~120 min and visualised on a ChemiDoc MP Imager (Bio-Rad, CA, USA).

4.3.9 Data availability.

The E. cloacae UW5 genome used in this study was previously sequenced and annotated as described (40) and is available from the NCBI GenBank database under the accession number CP011798. The raw RNA sequence data are available in the Gene

Expression Omnibus database under the accession number GSE122440.

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4.4 Results

4.4.1 RNA sequencing reveals a large number of TyrR-regulated genes.

The scope of gene regulation by TyrR was assessed by comparing the global transcription profiles of wild-type E. cloacae and a markerless tyrR deletion mutant by

RNA sequencing. Four independent biological replicates of each strain were grown in LB media to mid log phase for purification of RNA and subsequent sequencing. Each of the aromatic amino acids that are cofactors for TyrR is present in LB media at concentrations

(≥1 mM) that are typically used for gene expression studies with TyrR (56). Extracted

RNA was of high quality, with an average RNA integrity number (RIN) of 7.95

(maximum RIN of 10 indicates fully intact RNA) (57), and OD260/280nm ratios of 1.99. On average, each sample library yielded 16.5 million reads and 3.3 billion bases, providing a good level of coverage for coding sequences and other genomic features. Deletion of tyrR resulted in 155 genes (3.47% of all genes) showing differential gene expression compared to the wild-type strain (|log2FC| > 2, padj < 0.001) (Figure 4.1, and

Supplementary Tables S4.4, S4.5). Transcription was lower for 75 of these genes in the tyrR mutant (Supplementary Table S4.4), of which the most downregulated was dmpM

(ABT55_07800, log2FC = -5.36), encoding a predicted S-adenosylmethionine (SAM) O- methyltransferase. Eighty genes were derepressed (Supplementary Table S4.5), with the highest fold increase observed for yebE (ABT55_15985, log2FC = 6.16) which encodes a predicted inner membrane protein of unknown function. The reliability of the RNA-seq data was supported by the relative expression levels of the ipdC locus (Supplementary

Figure S4.1A) which is known to require TyrR for activation (33, 35). The RNA-seq data

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confirm previous findings, showing strong downregulation of ipdC (log2FC = -4.28) and weak, but significant, upregulation of akr (log2FC = 1.11, padj < 0.001) in the tyrR mutant strain. Consistent with reduced ipdC expression, production of the phytohormone IAA in the tyrR mutant was abolished (Supplementary Figure S4.1B).

4.4.2 Orthologues of known E. coli TyrR regulated genes are similarly regulated in

E. cloacae.

TyrR regulon genes previously characterized in E. coli were similarly regulated by TyrR in E. cloacae. In E. coli, the TyrR regulon was experimentally shown to consist of nine transcriptional units (1), and orthologues for each are present in E. cloacae. In the

E. cloacae tyrR mutant, strong derepression of transcription was observed for aroF-tyrA, aroG, aroP and tyrP (Table 4.1, and Supplementary Table S4.5), consistent with repression by TyrR in the presence of either phenylalanine or tyrosine in E. coli (1). aroL, which is also repressed by TyrR in E. coli (58), displayed increased levels of transcription in the E. cloacae tyrR mutant (log2FC = 1.78, padj < 0.001); however, this increase was below the four-fold cutoff value for differential expression in this study. folA, which is activated by TyrR in E. coli, was not appreciably regulated by TyrR in E. cloacae. An examination of the sequence upstream of folA revealed that the two TyrR boxes found in the E. coli folA promoter are absent in E. cloacae, while the IHF binding site and sigma-

70 promoter are conserved (Supplemental Figure S4.2). Therefore, the folA gene is not likely to be a TyrR regulon member in E. cloacae. The mtr and tyrB genes, which are weakly regulated by TyrR in E. coli, were not differentially expressed more than four- fold in the E. cloacae tyrR mutant (Table 4.1) (58). Together, our results are in good

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agreement with previous measurements of TyrR-regulated genes, with regulon members displaying up or down regulation consistent with expectations for a tyrR mutant.

4.4.3 RNA sequencing results were validated by quantitative RT-PCR.

To validate the RNA-seq data and investigate the dynamics of TyrR-regulated gene expression, several genes with a range of expression levels were analyzed by RT- qPCR. Genes were chosen that increased or decreased in expression in the tyrR mutant and a known TyrR-regulated gene, aroG, was included as a control. The quantitative RT-

PCR measurements of expression are in strong agreement with the values determined by

RNA sequencing (Figure 4.2). Two of the genes, dmpM and yagU, were consistently expressed at lower levels in the tyrR mutant compared to the wild type (Figure 4.2).

Genes proP, eco and aroG were highly expressed in the tyrR mutant as measured by both methods. Expression of cpxP was upregulated in the RT-qPCR data (21.8-fold), but to a lesser extent than in the RNA-Seq results (65.3-fold). While the cpxR gene was not differentially expressed according to the conservative criteria used in this study, both methods indicated a consistent 3.5-fold increase in expression in a tyrR mutant background. Overall, there was a strong, significant correlation between expression levels measured by RNA-Seq and quantitative RT-PCR (Pearson’s r = 0.975, p = 1.82 x 10-4,

Supplementary Figure S4.3).

4.4.4 The E. cloacae TyrR regulon has an expanded functional role.

Initial assessment of the RNA-Seq data revealed many genes with significant differential expression that were seemingly unrelated to aromatic amino acid metabolic processes, including genes encoding membrane-related functions such as transport, 136

anaerobic respiration, and chemotaxis. To understand the broader function of TyrR, GO terms were assigned to the E. cloacae genome and an enrichment analysis was performed with GOSeq using the Wallenius non-central hypergeometric distribution for both the up and downregulated genes to determine which terms, if any, were significantly more frequent than the genome-wide background for those categories (50). GO terms that are enriched in the downregulated and derepressed genes for each of the three main functional classes (48) are clustered together based on semantic similarity. For all differentially expressed genes, enriched GO terms were primarily identified in the biological processes and metabolic functions classifications, with fewer GO terms were related to cellular components (Supplementary Figure S4.4).

Analysis of significantly enriched GO terms among genes normally activated by

TyrR and downregulated in a tyrR mutant included anaerobic respiration, the glyceraldehyde-3-phosphate dehydrogenase complex, formate oxidation, chemotaxis, and transmembrane signalling (Table 4.2). The expression of neighbouring genes was often similarly upregulated, although in some cases at levels below the four-fold cutoff, suggesting that the genes may be components of polycistronic operons (Supplementary

Table S4.6). Operons that displayed downregulation included the arginine fermentation pathway arcABC (ABT55_04505-04495), formate dehydrogenase fdnGHI

(ABT55_14090-14080), anaerobic glycerol-3-phosphate dehydrogenase glpABC

(ABT55_18110-18120), an inactive iron permease efeUBC (ABT55_10470-10480), and threonine dehydratase tdcABC (ABT55_22195-22185) (Supplementary Table S4.4).

Expression of some respiratory operons was lower including cytochrome d ubiquinol oxidase cydABXybgE (ABT55_08730-08745) and dimethyl sulfoxide reductase dmsAB 137

(ABT55_09730-09735). Of the respiratory nitrate reductase A operon narGHJI

(ABT55_15245-15230), only the narH gene was more than four-fold lower in the tyrR mutant (log2FC = -2.29), while the remaining genes were slightly downregulated (log2FC

= -1.26 to -1.70, padj <0.001). The fdhF gene (ABT55_03610), encoding formate dehydrogenase-H, was also downregulated, as were two proteins of a hydrogenase complex (ABT55_20230-20235). Operons for fimbria, flagella, and chemotaxis systems were downregulated in the tyrR mutant. The fimbria structural operon fimAICDHF

(ABT55_07295-07320) was downregulated (log2FC -2.06 to -1.31; padj <0.001), however only expression of the leader gene fimA was more than four-fold lower in the tyrR mutant. The fimZ transcriptional repressor (ABT55_07325), located downstream of the main fim operon, was also significantly downregulated (log2FC = -2.55). A cluster of genes encoding methyl-accepting chemotaxis proteins (tap, tar) and chemotaxis signal transduction protein (cheRBYZ) (ABT55_16140-16165) was downregulated (log2FC = -

1.15 to -2.36, padj <0.001), but only the first two methyl accepting chemotaxis genes were more then four-fold lower. Several genes of a flagella biosynthesis operon

(ABT55_16495-16510) were downregulated, but only one, the fliD flagellar cap

(ABT55_16500), met the criteria of four-fold change (log2FC = -2.05). Genes that are part of a larger colanic acid biosynthetic operon were also downregulated (wcaE, gmd;

ABT55_17390, 17380).

Significantly enriched GO terms among the genes whose expression was enhanced in the tyrR mutant, and therefore normally downregulated by TyrR, included those that function in xenobiotic metabolism, aromatic catabolism, and the breakdown of organic carboxylic acids (Table 4.2). Genes for central carbon metabolism, solute 138

transport, multiple ABC transporters, and several transcription factors were upregulated in the mutant (Supplementary Table S4.5). Upregulated operons in the tyrR mutant included lldPRD (ABT55_02790-02780) for aerobic L-lactaldehyde metabolism, paaCDEFGHI (ABT55_12990-13330) for phenylacetate catabolism, tctCBA

(ABT55_18145-18155) encoding a tripartite citrate transporter and its associated two component response regulator tctDE (ABT55_18135-18140), and acs-act

(ABT55_02780-02790) encoding an acetate/glycolate symporter and acetyl-CoA synthetase (Supplementary Table S4.5). Members of a putative myo-inositol catabolic operon (ABT55_19015, 19020) were also expressed at higher levels, as was a 14-gene operon predicted to encode an aromatic dioxygenase and degradation pathway

(ABT55_21175-21220). Taken together, these results show that TyrR impacts the expression of diverse cellular and metabolic processes beyond those related to aromatic amino acids.

4.4.5 TyrR directly activates expression of a predicted SAM-dependent O- methyltransferase.

The strongest downregulated gene in the tyrR mutant background was dmpM, predicted to encode a SAM dependent O-methyltransferase (Supplementary Table S4.4), suggesting that TyrR is required for its expression. While originally annotated as a methyltransferase involved in the biosynthesis of the tyrosine-derived antibiotic puromycin, a BLAST search of the Streptomyces alboniger puromycin biosynthesis genes against the E. cloacae genome indicated that E. cloacae lacks homologous genes to produce this antibiotic. Despite sharing a conserved structural fold, SAM-dependent

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methyltransferases share little sequence identity and recognize a wide range of substances, making it difficult to assign physiological function to these enzymes (59).

According to the structural classifications of SAM-dependent methyltransferases, the predicted DmpM polypeptide shares structural features with plant small molecule O- methyltransferases based on the presence of an extended N-terminus comprised of a putative dimerization domain and an additional alpha helix inserted between beta sheet-5 and alpha helix-D, and the absence of the ‘C’ alpha helix in the main structural fold (59-

61). The plant methyltransferases are involved in the secondary metabolism of a number of small molecules including phenylpropanoids derived from phenylalanine and tyrosine.

A search of the Protein Data Bank using HHpred with the DmpM amino acid sequence as a query returned several top hits for bacterial methyltransferases involved in antibiotic production and plant O-methyltransferases (60, 62) (Supplementary Table S4.7).

In wild-type E. cloacae, expression of dmpM was significantly enhanced in the presence of tyrosine, and to a lesser extent phenylalanine, increasing 102.7- and 6.9-fold, respectively, compared to in M9 medium without aromatic amino acids (Figure 4.3A).

Expression in tryptophan was not significantly different from that in M9 minimal medium alone. The dmpM transcript levels were 100-fold lower in the tyrR mutant relative to the wild-type strain in the presence of tyrosine (Figure 4.3B). These findings indicate that expression of dmpM is dependent on TyrR primarily in conjunction with tyrosine.

Within the dmpM promoter region are three predicted TyrR boxes, each with varying degrees of identity to the TyrR box consensus sequence (Figure 4.4). Box 1 is centered 32 bp, or 3 turns of the DNA helix, upstream from the predicted -35 sequence 140

and matches the consensus sequence at 9/12 base pairs with a A/T rich spacer. The second predicted TyrR box is centered 63 bp upstream from the -35 sequence, and also matches the consensus at 9/12 base pairs with a A/T rich spacer sequence, consistent with the criteria for a strong TyrR binding site (1, 20). Thus, these binding sites are on the same face of the DNA, three and six helical turns upstream of the -35 site, respectively. A third predicted TyrR box matching at 10/12 bp to the consensus sequence is located further upstream, 159 bp from the -35 sequence and 7.4 turns of the DNA helix upstream from box 2. The location of boxes 1 and 2 relative to the predicted RNA polymerase binding site, and to each other, strongly suggests their function as TyrR boxes for the positive regulation of dmpM, while the distal box 3, which lacks a highly conserved G, may have limited involvement in TyrR regulation of the gene. EMSAs, using a 6-FAM labeled DNA probe spanning the three boxes, revealed a distinct shift of the wild-type dmpM DNA probe incubated with purified TyrR, and a decrease in the amount of free

DNA with increasing protein concentration (Figure 4.5). The amount of protein required to shift half the wild-type DNA was between 0.44 µM and 0.87 µM, and at 2.19 µM almost all free DNA was shifted. The migration of TyrR-bound DNA probe was further retarded as the concentration of protein increased, which may indicate multiple dimers of bound TyrR. There was no noticeable effect on the migration of TyrR-DNA complexes when EMSAs were performed in the presence of aromatic amino acids (data not shown).

In contrast, a mutated dmpM probe that contained key base pair substitutions in each of the three TyrR boxes failed to bind TyrR and shifted DNA was abolished (Figure 4.5).

Therefore, TyrR directly binds to DNA bearing the dmpM promoter at one or more TyrR

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boxes. The mode of interaction of TyrR with the individual boxes is unknown, as is the influence, if any, of aromatic amino acid cofactors on DNA recognition.

4.4.6 The Cpx regulon is perturbed in a tyrR mutant.

An interesting observation in the RNA-seq data was the identification of multiple differentially expressed genes that are also members of the regulon of the two-component regulator CpxRA that controls the envelope stress response. For example, the highest upregulated gene in the tyrR mutant, yebE, an inner membrane protein of unknown function, is positively regulated by CpxR in E. coli. (63). Expression of several other

CpxR activated genes was increased in the tyrR mutant, including ytfK, degP, aroG, yccA, yceJ, spy, htpX, yebE, acrD and yqaE (Supplementary Table S4.5) (63-70). CpxR repressed genes efeU and glpABC were highly repressed (log2FC = -2.91 to -4.98) in the tyrR mutant (Supplementary Table S4.4). Genes controlled by CpxR in S. typhimurium were among the differentially expressed genes including ybbA, ybiJ, mdtJ, eco, deoA, fimA and dcuB (71). The cpxP and cpxA genes, which are both components of the Cpx response and encode the periplasmic chaperone CpxP and transmembrane histidine kinase CpxA that together with the CpxR response regulator mediate the Cpx stress response, were significantly upregulated in the tyrR mutant (log2FC = 6.03 cpxP, log2FC

= 2.15 cpxA) (Supplementary Table S4.5) indicating that TyrR normally represses their expression. Expression of the cpxR gene (ABT55_01655) was also higher in the mutant

(log2FC =1.81, padj <0.001) (Supplementary Table S4.6), although it did not meet the conservative criteria for differential expression in this study. The divergently transcribed cpxP and cpxRA operons in E. coli are activated by CpxR binding at three CpxR boxes in

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the cpxP-cpxR intergenic region (73, 74). The DNA binding motifs recognized by CpxR and TyrR share sequence similarity, and in the E. coli aroG promoter the binding site for these transcription factors overlap (63, 72). The enrichment of CpxR-regulated genes in the RNA-seq data, including cpxP and cpxA, and the presence of orthologous CpxR boxes and promoter elements in the E. cloacae cpxP-cpxR intergenic region (Figure 4.6) suggested that TyrR may repress the cpxP-cpxRA operon by co-opting the CpxR binding sites as weak TyrR box.

The possibility of TyrR recognizing and interacting with the cpxP and cpxR promoter sequences was examined by RT-qPCR in the presence and absence of aromatic amino acids and by EMSAs to assess specific protein-DNA interactions. Transcription from the cpxP promoter in wild-type E. cloacae was significantly downregulated by tryptophan and phenylalanine (1.5- and 1.9-fold, respectively) compared to M9 minimal media alone (Figure 4.7A), while growth in tyrosine induced a much greater 24.9-fold increase in expression relative to M9 medium (Figure 4.7A). In contrast, wild-type expression of cpxR was not different when grown in minimal medium with and without any of the aromatic amino acids (Figure 4.7A). Therefore, the downregulation of cpxP by tryptophan and phenylalanine and upregulation by tyrosine in the wild-type strain is not due to altered levels of CpxR. Relative to expression in the wild-type background, transcription of cpxP was significantly greater in the tyrR mutant in M9 medium alone

(25.5-fold), or media supplemented with tryptophan (45.5-fold) or phenylalanine (50.1 fold) (Figure 4.7B). Transcription of cpxP in tyrosine was not significantly different in the tyrR mutant relative to the wild-type. Expression of cpxR was consistently higher under all conditions in the tyrR mutant strain (3.9- to 4.8-fold increase), with aromatic 143

amino acids having no effect (Figure 4.7B). Taken together, these results indicate that transcription of cpxR and cpxP is repressed by TyrR, and that cpxP is derepressed by tyrosine.

While canonical TyrR boxes were not apparent in the cpxP or cpxR promoter, we proposed that TyrR may recognize and bind to one or more of the three CpxR DNA binding sites (Figure 4.6). This hypothesis was predicated on the observation that: (i)

TyrR has previously been shown to bind a non-canonical site in E. cloacae (35), (ii) TyrR may bind sequences lacking the conserved G-C pair, although at a much reduced affinity

(75), and (iii) both TyrR and CpxR have been experimentally demonstrated to bind the same DNA sequence in the aroG promoter (63, 72), although no study has looked at the concurrent binding of both proteins. Incubation of a 6-FAM labelled DNA probe corresponding to the wild-type cpxP-cpxR intergenic region with increasing concentrations of TyrR protein resulted in the migration of the probe in EMSAs (Figure

4.8). In the presence of 2.19 µM of purified TyrR, about 50% of the probe had shifted, and with 4.39 µM TyrR a strong shifted band was present. Point mutations were introduced to each of the CpxR boxes at nucleotides predicted to play similar roles to the

G-N14-C pair within a TyrR box (Figure 4.6) (1). Mutations to CpxR boxes 2 and 3 had no discernible effect on the ability of TyrR to bind to the DNA, as probes containing these sequences shifted similarly to the wild-type sequence. Mutations to CpxR box 1 resulted in a weak reduction in shifted DNA probe (Figure 4.8). Among the three predicted CpxR boxes, box 1 has the closest agreement to the TyrR consensus sequence with 8/12 base pair matches, and its proximity to the -35 elements of both the cpxR and cpxP promoters may enable a bound TyrR dimer to interfere with RNA polymerase 144

binding. In all EMSA reactions, TyrR was pre-incubated with a 10-fold excess of nonspecific competitor DNA to ensure sequence specific DNA binding. Inclusion of aromatic amino acids in the binding reactions did not have any effect on TyrR-DNA migration (data not shown). These findings were reproducible in multiple experiments

(data not shown), indicating that CpxR box 1 contributes to TyrR binding to the cpxR- cpxP intergenic region. Together, the results show that TyrR represses expression of the cpxP-cpxRA operons through direct binding of the promoter regions at a non-canonical site that overlaps a predicted CpxR binding site, and thereby alters expression of the

CpxAR regulon.

4.5 Discussion

The mechanism by which TyrR exerts control over gene expression is complex, and its ability to either activate or repress transcription of a target gene is dependent on a number of interconnected factors including DNA binding site affinity, the number and location of the binding sites, concentration of available TyrR protein in the cell, relative promoter strength of target genes, DNA conformation, nucleoid binding proteins, and the presence of aromatic amino acid cofactors (1, 76, 77). Thus, TyrR is a versatile transcription factor that integrates multiple signals to exert appropriate transcriptional control over each member of its regulon in response to physiological and environmental factors. While the regulon has been extensively studied in E. coli, there is growing evidence that it is more complex in other gammaproteobacteria that produce TyrR (27,

28, 32, 78), possibly reflecting the lifestyles differences among these bacteria. In this

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study, the results of high-throughput RNA sequencing revealed that in E. cloacae TyrR significantly impacts the expression of 155 genes, including repression of the CpxR envelope stress response regulon through direct interaction with a non-canonical TyrR box overlapping a CpxR binding site in the cpx promoters, and direct activation of a

SAM-dependent menthyltransferase in response to tyrosine. Additionally, genes for motility, aromatic catabolism, and respiration are regulated by TyrR. Regulon expansion is not unexpected for a bacterium such as E. cloacae that inhabits the diverse environments of the human intestinal tract and the plant rhizosphere.

4.5.1 The TyrR regulon overlaps the Cpx stress response regulon

A surprising finding of this study was the overlap between the TyrR and CpxR regulons, and the responsiveness of the cpx operon to aromatic amino acids. The cpxP- cpxRA operon encodes a two-component signal-transduction pathway, comprised of a sensory kinase CpxA, cytoplasmic response regulator CpxR, and a periplasmic accessory protein CpxP. CpxA, an integral cytoplasmic membrane protein, uses ATP to autophosphorylate a conserved histidine residue in response to diverse signals, including alkaline pH, altered membrane lipid composition, copper, misfolded pilus subunits, detergents, indole, osmolarity, EDTA, hydrophobic surfaces, or surface attachment via the outer-membrane lipoprotein NlpE (74, 79-85). Phosphorylated CpxA activates CpxR by transphosphorylation at a conserved aspartate residue (79). CpxR~P controls a regulon that encompasses genes for the alleviation of envelope stress, type III secretion, motility, chemotaxis, adherence, biofilm formation, virulence, cell wall integrity, and responses to antibiotics (85-91). Chaperone CpxP binds misfolded periplasmic proteins and also

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interacts with the periplasmic domain of CpxA, inhibiting its autokinase function (79,

92). The primary function of CpxP is thought to be fine-tuning the Cpx response (79, 92,

93). Binding of misfolded proteins to CpxP titrates it away from CpxA, thereby increasing CpxA autophosphorylation and activation of CpxR (93, 94). Transcription of the cpxP gene is strongly upregulated by CpxR~P, providing a rapid negative-feedback mechanism to prevent constitutive activation of the Cpx response (95, 96).

Downregulation of the cpx operon by TyrR depends, at least in part, on the interaction of TyrR with a non-canonical binding site situated immediately upstream of the -35 element of both the cpxR and cpxP promoters and overlapping a CpxR binding site. Previously, we have shown that such atypical TyrR-DNA interactions modulate expression of the akr gene in E. cloacae (35). TyrR bound to a weak box that overlaps the -10 element of the akr promoter independently of aromatic amino acid cofactors and downregulated transcription of akr (35). Repression of the cpx operons by TyrR may be due to steric occlusion of RNA polymerase and/or CpxR~P binding sites. When bound to promoter DNA E. coli RNA polymerase has a footprint extending from roughly -55 to -5 base pairs relative to the transcription start site (97, 98), which overlaps the TyrR binding site upstream of both cpxR and cpxP. DNase I foot printing studies with TyrR indicate that the region of DNA protected by the protein extends 2 base pairs beyond the 18 base pair consensus motif required for DNA binding (72). This coverage would theoretically prevent both RNA polymerase and CpxR~P from binding to the cpx promoters, thereby repressing expression.

Repression of cpxP is alleviated by tyrosine, which is unusual among TyrR- regulated genes. Tyrosine, with ATP, induces hexamerization of TyrR and normally 147

increases its DNA binding affinity and binding to promoter regions with multiple boxes

(1). However, the presence of aromatic amino acid cofactors that alter the oligomerization status of TyrR and its relative affinity for a DNA binding site also influence the concentration of available TyrR. In the presence of tyrosine or phenylalanine, TyrR hexamers and dimers, respectively, are recruited to promoters with multiple TyrR boxes reducing available TyrR concentrations in the cell. Tyrosine- induced hexamerization may titrate TyrR away from cpx promoters, which are predicted to be weakly bound, to other TyrR regulated promoters that have a greater affinity for the protein, thereby derepressing cpxP expression.

In addition to repression of the cpxP-RA operons, TyrR may directly modulate expression of other genes in the Cpx regulon. In E. coli MG1655 CpxR activates expression of aroG, which encodes the phenylalanine-sensitive DAHP synthase that catalyzes the initial step in synthesis of the aromatic amino acid precursor chorismate; aroG is also repressed by TyrR (63). CpxR also weakly activates expression of aroK, encoding shikimate kinase I of the chorismate pathway, which leads to the biosynthesis of the aromatic amino acids (63, 99). TyrR represses aroL encoding shikimate kinase II, which is the dominant isoenzyme (100), and aroF, a tyrosine-sensitive DAHP synthase

(1). A current model suggests that CpxR upregulates the chorismate pathway to increase production of aromatic metabolites such as indole that enhances Cpx-mediated resistance to antimicrobial compounds (101). This may be a fine-tuning mechanism to provide aromatic metabolites to activate antimicrobial resistance when intracellular pools of chorismate are low due to TyrR mediated repression of the chorismate biosynthetic pathway. 148

4.5.2 TyrR activates expression of a predicted SAM-dependent O- methyltransferase.

Despite the initial annotation of TyrR-activated dmpM as a SAM-dependent methyltransferase gene involved in puromycin biosynthesis, the function of the encoded protein is unknown. These enzymes catalyze the transfer of a methyl group from SAM to a wide variety of molecular targets, and are classified based on structure, substrate specificity, and the target atom for methylation (59). The strong upregulation of dmpM in the presence of tyrosine, and to a lesser extent phenylalanine, suggests that the encoded protein may utilize these amino acids or a metabolic derivative, as substrates. The antibiotic puromycin is synthesised from tyrosine; however, E. cloacae UW5 lacks the additional genes for synthesis of this antibiotic, which seem to be restricted to

Streptomyces alboniger (102). Several other bacterial SAM-methyltransferases involved in antibiotic production share sequence identity and predicted structural elements with the

DmpM polypeptide (Supplemental Table S4.4). While SAM-dependent methyltransferases share a common structural fold, there is little conservation at the level of sequence identify, making sequence based identification difficult.

The activation of dmpM transcription by TyrR fits known modes of upregulation.

Activation by TyrR requires a strong box positioned upstream of the -35 sequence of a promoter such that it may interact with the carboxyl-terminal domain of the alpha-subunit of RNA polymerase (1). There are two TyrR boxes upstream of dmpM, centered 32 and

63 base pairs upstream from the -35 element of a predicted sigma-70 promoter, placing them 3 and 6 full turns of the DNA helix, respectively, from the -35 sequence. The strong box of the tyrP promoter of E. coli is centered 28 base pairs upstream of the -35 element 149

allowing a bound TyrR dimer to interact with RNA polymerase and activate transcription in the presence of phenylalanine, however it is not situated optimally to allow maximal transcription activation (103). Mutation of the tyrP promoter that alter the spacing between the strong box and -35 sequence revealed that 31-32 base pairs between centres was optimal for transcription in the presence of phenylalanine and tyrosine (103).

Therefore, the TyrR box upstream of dmpM is positioned to activate high levels of expression, as was confirmed by quantitative RT-PCR in the presence of tyrosine. TyrR activated genes folA and mtr also require the hexameric form of TyrR for activation with tyrosine (2, 21). Together with the location of the promoter regulatory elements, the strong (>100 fold) activation of dmpM in response to tyrosine suggests that this gene is activated by the hexameric form of TyrR in vivo.

4.5.3 TyrR regulates rhizosphere colonization and survival traits

Many of the E. cloacae TyrR-regulated genes are common among rhizobacteria, and may reflect a role for TyrR in rhizosphere competence (104, 105). Previously, we found that TyrR controlled expression of the ipdC gene for IAA biosynthesis in E. cloacae (33, 35). IAA is produced by many plant-associated bacteria, and it positively affects the growth and health of host plant root systems (106). Other major requirements for effective and competitive rhizosphere colonization include motility and chemotaxis

(105), and several methyl accepting chemotaxis proteins (MCP) and flagella biosynthesis genes are upregulated by TyrR. MCPs are transmembrane sensors that allow bacteria to respond to changing concentrations of small molecule attractants or repellants. These signals are transmitted to the flagella motor via the chemotaxis signal transduction

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proteins (encoded by che genes), which alter flagellar rotation to propel the bacterium toward a favorable environment (107). While E. coli has comparatively few MCPs and che genes, reflecting its modest sensory requirements, free-living soil and plant- associated bacteria often have an expanded repertoire of chemotaxis genes, which reflects a greater need for signalling pathways in complex environments (108, 109). MCPs are necessary to mediate movement towards root or seed exudates by sensing amino acids, organic acids and other nutrients (110, 111). Rhizobacteria Azospirillum brasilense and

Pseudomonas fluorescens with mutations in components of the flagella or chemotaxis system, were unable to colonize the rhizospheres of wheat and tomato roots, respectively

(112, 113). TyrR also upregulates the fliD gene, which encodes the flagellar cap protein.

This protein lies at the tip of the flagellin filament and plays a critical role in its assembly

(114).

Adhesion structures, such as fimbria, are important for initial attachment to root and other surfaces (115, 116). The fimA and fimZ genes encoding the type I fimbrial subunit and transcriptional activator, respectively, were downregulated in the tyrR mutant. Several genes of the colanic acid biosynthesis locus were also downregulated.

Colanic acid is an important exopolysaccharide involved in biofilm formation in E. coli

(117), and its enhanced production has been detected in the lettuce rhizosphere (118).

Catabolism of sugar and organic acids that are present in the rhizosphere was altered in the tyrR mutant. Specifically pathways for citrate, succinate, fructose and myo-inositol were perturbed in the tyrR mutant. Myo-inositol catabolism is important for root nodule formation by rhizobia, and the ability to catabolize inositol, and its derivatives known as rhizopines, confers a competitive advantage in the rhizosphere (119, 120). 151

4.5.4 Aromatic catabolism is regulated by TyrR.

A number of aromatic catabolic genes, including the paa locus encoding the phenylacetate degradation pathway (121), were repressed by TyrR in E. cloacae. The fourteen paa genes share genome synteny with E. coli and are arranged in three transcriptional units comprised of paaZ, paaABCDEFGHIJK, and paaXY (Supplementary

Figure S4.6A); expression of each is driven by a PaaX controlled promoter in E. coli.

Only the main paa operon, specifically genes paaCDEFGHI, was more than four-fold upregulated in the tyrR mutant (Supplementary Figure S4.6B, Supplementary Table

S4.5). Expression of the PaaX repressor was not significantly different between the strains (log2FC = 0.1) and there were no apparent TyrR boxes in the paaA promoter

(Supplemental Figure S4.6C); therefore, derepression of the operon in the tyrR mutant is likely due to inhibition of PaaX by phenylacetyl-CoA, the first intermediate of the pathway (122). One or more peripheral pathways for phenylalanine catabolism may be upregulated in the tyrR mutant, yielding phenylacetate, which is then ligated with CoA by

PaaK (not significantly altered in the mutant strain). These results suggest that phenylacetic acid catabolism is normally downregulated by TyrR. Although the mechanism of repression is not known, it does not appear to be due to direct interaction of TyrR with the paaA promoter region or increased expression of repressor PaaX.

A second cluster of eight genes (ABT55_21185-21220), possibly comprising a single transcriptional unit involved in aromatic metabolism, was also significantly upregulated in the tyrR mutant indicating that this locus is normally repressed by TyrR.

The predicted functions of the genes in this cluster include an aromatic ring hydroxylating dioxygenase, a regulatory gene, a transporter and several hydrolytic 152

cleavage enzymes. Aromatic ring hydroxylating dioxygenases, such as the naphthalene dioxygenase of Pseudomonas, are important for the metabolism of aromatic compounds in the environment and catalyze the insertion of molecular oxygen into the benzene ring followed by the ortho- or meta-cleavage of the aromatic ring (123). These enzyme complexes function in catabolic pathways that yield, among other products, phenylacetic acid, which is further degraded by the paa gene products as described above. Aromatic dioxygenases have relaxed substrate specificity, and may oxidize a number of similar molecules, making assignment of function difficult without biochemical investigations

(123). The presence of genes encoding transporters and regulators in some aromatic catabolic clusters may indicate that the substrates brought into the cell by the permease act as effectors for their respective regulators (124). A putative TyrR-binding site is located upstream of this operon, and directly overlaps with the -10 element of a possible sigma-70 promoter. However, this box only matches the TyrR consensus at 8/12 base pairs and presumably would require an adjacent strong TyrR box to facilitate its binding.

Taken together, the results here indicate that TyrR contributes to the repression of aromatic catabolism in E. cloacae. This is in contrast with the role of the orthologous regulator PhhR in Pseudomonas, which is primarily responsible for activating aromatic catabolism (31).

4.5.5 The TyrR regulon includes genes for anaerobic metabolism

Significant dysregulation of genes for anaerobic metabolism is observed in the tyrR mutant. Expression of respiratory nitrate reduction pathways and several components of anaerobic electron transport chains including dehydrogenases and

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terminal reductases was reduced in the mutant compared to the wild-type strain.

Respiratory nitrate reduction pathways convert nitrate to ammonium for nitrogen assimilation or are associated with denitrification. The initial reduction of nitrate to nitrite is catalyzed by the membrane-bound respiratory nitrate reductase complex A (NarGHJI), nitrite is then reduced to ammonium by the cytoplasmic NirBD NADH-nitrite reductase

(125); expression of both complexes is downregulated in the E. cloacae tyrR mutant. In

E. coli, nirBD are induced under anaerobic conditions and required specifically for the detoxification of nitrite from nitrate respiration (126). Nitrate reduction draws electrons from the membrane quinol pool, coupling reduction of nitrate to generation of the proton motive force (127). The nitrate-inducible formate dehydrogenase complex (fdhIHG), and pyruvate formate lyase (pflAB) are also downregulated in the mutant, suggesting that

TyrR directly or indirectly upregulates both the quinone-reducing formate dehydrogenase and the quinol-oxidizing nitrate reductase that form a proton motive redox loop. Formate, which is oxidized by formate dehydrogenase, is produced from anaerobic catabolism of organic substrates by pyruvate formate lyase (127). Nitrate reductase may also accept electrons from both anaerobic (glpABC) and aerobic (glpD) glycerol-3-phosphate dehydrogenase, both downregulated in the mutant. The nitric oxide tolerant cytochrome bd-1 terminal oxidase encoded by cydAB-cydXybgE, along with Hcp/Hcr, is part of the gram-negative response to bactericidal nitric oxide and enhances tolerance to nitrosative stress (128, 129); genes encoding several of these proteins were expressed at lower levels in the tyrR mutant. Expression of this oxygen reductase is induced by multiple stressors including oxygen limited conditions and may facilitate bacterial colonization of oxygen poor environments (128). Other terminal reductases downregulated in the tyrR mutant 154

include fumarate and DMSO reductases. Fumarate reductase may form electron transport chains with either the NADH dehydrogenase complex or glycerol-3-phosphate dehydrogenase (130). The DMSO terminal reductase generates a proton motive force in concert with the proton pumping activity of NADH dehydrogenase under anaerobic conditions (130, 133). The membrane potential generated by the activity of these protein complexes may drive the uptake of amino acids (131, 132).

In support of the role of TyrR in anaerobic metabolism, the lldPRD operon for aerobic lactate metabolism (134) is derepressed in the tyrR mutant. Lactate is taken up by a permease (encoded by lldP) and converted to pyruvate via lactate dehydrogenase

(encoded by lldD); lldR encodes a regulator (134). Expression of the operon is repressed under anaerobic conditions by ArcA (135).

Although significant changes to respiratory complexes were observed in the tyrR mutant strain, it is not known whether the role of TyrR is direct or a consequence of changes in the physiological state of the mutant brought on by loss of TyrR. Many of these pathways are regulated by specific transcription factors or more broadly impacted by global transcription factors. ArcA is a global regulator of several hundred E. coli genes, responding to changes in the oxidation state of the quinone pool and the transition to anaerobic growth. Fumarate nitrate reductase (FNR) is another global transcriptional regulator that controls anaerobic respiratory pathways (136, 137). Reduction of oxygen availability occurs rapidly in the rhizosphere and in the animal intestinal tract, as nutrients are rapidly oxidized and E. cloacae must respond by expressing enzymes to utilize alternate electron donors and acceptors to continue to growth under anoxic conditions. 155

4.6 Conclusions

In addition to confirming TyrR regulation of aromatic amino acid biosynthetic and transport genes in E. cloacae, transcriptome analysis demonstrated that TyrR has broad control over the physiology and ecology of this bacterium including the envelope stress response, aromatic catabolism, anaerobiosis, and colonization of a host environment. The results demonstrate that the TyrR regulon is more expansive in

Enterobacteraceae than previously thought. However, the specific role of TyrR in many of these processes has yet to be investigated, and may be direct through interaction with gene promoters or indirect through intricate regulatory networks with other transcription factors, such as CpxR. Nonetheless, the work presented here provides further insight into the complexity of TyrR regulatory interactions and its role beyond aromatic amino acid biosynthesis and uptake in the commensal rhizobacterium and opportunistic human pathogen E. cloacae.

4.7 References

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Table 4.1. TyrR-regulation of aromatic amino acid transport and biosynthesis genes is similar in E. cloacae and E. coli.

a Expression (log2FC) E. cloacae locus tag Gene Product E. cloacae E. colib ABT55_19500 aroF-tyrA DAHP synthase, tyrosine-repressible 6.04 4.06 ABT55_06645 aroL Shikimate kinase II 1.78 1.18 ABT55_16305 tyrP Tyrosine transporter 4.31 1.74 ABT55_22355 mtr High affinity tryptophan transporter -1.49 -0.58 ABT55_05725 aroP General aromatic amino acid transporter 5.09 0.64 ABT55_03500 tyrB Aminotransferase -0.21 0.32 ABT55_08825 aroG DAHP synthase, phenylalanine-repressible 2.75 0.79 ABT55_05420 folA DihydrofolAte reductase 1.50 -1.30c a Expression in tyrR mutant relative to wild-type strain b Data derived from (1) for strains grown in minimal media c Data derived from (2) for strains grown in tyrosine

166

Table 4.2 . Significantly enriched GO terms among the genes up and downregulated in the tyrR mutant. GOSeq called terms as significantly enriched at padj<0.01 applying the Benjamini and Hochberg correction method. Expression Gene ontology term Process

UP GO:0006805, GO:0071466, GO:0042178, GO:0009410 Xenobiotic metabolism GO:0042537, GO:0010124, GO:0019439, GO:1901361 Aromatic catabolic processes GO:0072329, GO:0016054, GO:0046395 Organic carboxylic acid breakdown DOWN GO:0009061, GO:0009055 Anaerobic respiration GO:0009331, GO:0046168 Glyceraldehyde-3-phosphate dehydrogenase GO:0015944 Formate oxidation GO:0006935 Chemotaxis GO:0004888 Transmembrane signalling

167

cvpA tyrA yebE tyrR 300 ydcI ybbP aroF aroP cpxP

ycdO

250 spy

dmpM yagU ycfS ipdC ybbP 200 eco ycdB ybiJ rbsD 150

ABT55_04870 yqaE tyrP -log10(padj) 100

MsI ABT55_07875 yﬂP 50 0dT thcD ABT55_05730 nirD glpA ydcI glpC glpB yhjX tctD arcC 0 ABT55_16350

-6 -4 -2 0 2 4 6

log2FoldChange

Figure 4.1. Volcano plot displaying differentially expressed genes in E. cloacae wild type and tyrR mutant strains. The green dots indicate genes whose transcripts increased >2 log2FC (false discovery rate (FDR)-adjusted p value < 0.001) in the tyrR mutant compared to the wild type strain (n=75); the red dots indicate genes whose transcripts were decreased < -2 log2FC (FDR-adjusted p value <0.001)(n=80). Genes whose expression was |log2FC| > 3 are labelled. The y-axis corresponds to the negative base-10 logarithm of the adjusted p-values, and the x-axis displays the log2 fold-change (log2FC) value. The horizontal and vertical dotted blue lines indicate the threshold criteria of Benjamini-Hochberg corrected p -value = 0.001 and |log2FC| = 2, respectively.

168

A 1 B 100

0.1

0.01

RelativeExpression RelativeExpression

0.001 1 dmpM yagU proP eco aroG cpxP cpxR

Figure 4.2. Verification of RNA-Seq results by quantitative RT-PCR. Differential expression of genes that were (A) downregulated or (B) derepressed in the tyrR mutant as measured by RNA-Seq (white bars) and quantitative RT-PCR (grey bars). Relative expression was determined using the ΔΔCt method for RT-qPCR by comparing against the respective wild type transcripts. The results are normalized against recA expression levels. Data represents the average of three biological replicates, error bars represent the standard error of the mean.

169

A 1000 B 1 * * * 100 0.1 10 * 0.01 1 * RelaSve Expression RelaSve Expression

0.1 0.001 TRP PHE TYR M9 TRP PHE TYR

Figure 4.3. Activation of dmpM expression requires aromatic amino acids with TyrR. Relative expression of dmpM was determined in E. cloacae wild-type (A) and the isogenic tyrR mutant (B) grown in M9 minimal media supplemented with or without 1 mM of each aromatic amino acid by quantitative RT-PCR as described in the Materials and Methods. Relative expression was determined using the ΔΔCt method by comparing to the respective wild-type transcripts in M9 minimal media alone (A) or to the wild type transcripts in each growth condition (B). Expression was normalized to recA expression levels. Data represents the average of three biological replicates, error bars represent the standard error of the mean. The asterisks indicate significant differences (p<0.05) from the control treatment.

170

dmpM-F TyrR Box 3 CCGTAAACAACCGTTAATTTCGCATTCAGATATATATAACAATATTTACAGGCCACACCCGGAAATGAATTGAAAAA t TyrR Box 2 CGCATGATCTTTAACGGTATTAGTTACCAGCATTAAAAAACGGTGACTTAATGTCCGGTTTTATTTACAGGAGTCTG a t TyrR Box 1 dmpM-R-FAM TCTTATGTAACTTAAATGTTACTCCTGTTTTCTGGGAGGGGGCGTTACATATGAAGTAATAAAGGTTTAACTGTACC a t -35 -10 dmpM CGGTAGCTGTAAATAAGAAATTCACATTTTTTAATCCTTCAGGAGCTAATATGACTGACAAACATATATTTTCAGCG

Figure 4.4. Nucleotide sequence and regulatory elements of the dmpM promoter region. The three predicted TyrR boxes are boxed and point mutations made to each site are indicated in bold below the sequence. The putative -35 and -10 elements of a sigma- 70 promoter are indicated in bold text, along with the dmpM start codon. The primer binding sites used for EMSAs are marked with arrows.

171

Probe WT dmpM mutant dmpM TyrR (µM) - 0.08 0.44 0.87 2.19 4.39 - 0.08 0.44 0.87 2.19 4.39

Figure 4.5. Sequence specific binding of TyrR to the dmpM promoter. Electrophoretic mobility shift assays (EMSAs) were performed with 100 ng of 6-FAM labeled DNA fragments spanning the wild-type dmpM promoter region or the same region containing point mutations in the three TyrR boxes, and increasing concentrations (0-4.39 µM) of purified TyrR. The location of free unbound DNA (F) is indicated. The mutations made to key base pairs predicted to be required for TyrR DNA binding abolish interaction with the promoter.

172

CpxR box 1 cpxR cpx-F -35 a a t TTATTCATTGTTTAAATACCTCCGAGGCAGAAATTACGACATCAAGGCTATCTAATCCATGACTTTACGTTGTTTTACACC +1 -10

CpxR box 2 CpxR box 3 aaa t a tt cpx-R-FAM CCCTGACGCATGTTTGCAGCCTGAATCGTAGACTGTCTCTCGTTGAATCGCGACACGAAAGATTTTGGGAGCAAGTGATGC -35 -10 +1 cpxP

Figure 4.6. Nucleotide sequence and predicted regulatory elements in the intergenic region of divergently transcribed cpxP and cpxR. The locations of CpxR binding sites are boxed, and point mutations made to each are indicated in bold above each sequence. The -35 and -10 elements of the sigma-70 promoters are underlined, transcription start sites are indicated in bold italics and start codons are in bold. The primer binding sites used for EMSAs are marked with arrows. The locations of promoter elements and CpxR binding sites are inferred from orthologous sequences in the E. coli cpx promoters, which share a high degree of sequence conservation.

173

A 100 B 100 * * * * 10 10 * * * *

1 1

* * RelaSve Expression RelaSve Expression 0.1 0.1 TRP PHE TYR M9 TRP PHE TYR

Figure 4.7. TyrR downregulates expression of the cpx operons. Relative expression of cpxR (white bars) and cpxP (grey bars) was determined in E. cloacae wild-type (A) and the isogenic tyrR mutant (B) grown in M9 minimal media supplemented with or without 1 mM of each aromatic amino acid by RT-qPCR as described in Materials and Methods. Relative expression was determined using the ΔΔCt method by comparing to the respective wild-type transcripts in M9 minimal media alone (A) or to the wild type transcripts in each growth condition (B). Expression was normalized to expression levels of the recA housekeeping gene. Data represents the average of at least three biological replicates, error bars represent the standard error of the mean. The asterisks indicate significant differences (p<0.05) from the control treatment.

174

Probe WT Δbox 1 Δbox 2 Δbox 3 TyrR (µM) - 0.08 0.87 2.19 4.39 - 0.08 4.39 - 0.08 4.39 - 0.08 4.39 B

Figure 4.8. Sequence specific binding of TyrR to the cpxR-cpxP promoter region. Electrophoretic mobility shift assays (EMSAs) were performed with 100 ng of 6-FAM labeled DNA fragments spanning the wild type cpxP-cpxR intergenic region, or the same region containing point mutations made to each of the CpxR binding sites, and with increasing concentrations (0-4.39 µM) of purified TyrR. The location of free unbound DNA (F), and shifted protein-DNA complexes (B) are indicated. Only mutations made within the box 1 sequence reduce TyrR binding to the promoter.

175

4.8 Supplementary information

4.8.1 List of supplementary tables

Supplementary Table S4.1. Bacterial strains and plasmids used in this study...... 178

Supplementary Table S4.2. Oligonucleotides used in this study...... 179

Supplementary Table S4.3. Sequences of the synthesized cpx and dmpM promoters used for electrophoretic mobility shift assays...... 180

Supplementary Table S4.4. Genes that were significantly downregulated in the tyrR mutant strain of E. cloacae relative to the wild type...... 181

Supplementary Table S4.5. Genes that were significantly derepressed in the tyrR mutant strain of E. cloacae relative to the wild type...... 183

Supplementary Table S4.6. HHPred results with DmpM query sequence...... 185

4.8.2 List of supplementary figures

Supplemental Figure S4.1. TyrR is necessary for expression of ipdC and production of

IAA...... 186

Supplemental Figure S4.2. Multiple sequence alignment of the folA promoter region of four Enterobacteriacea strains...... 187

Supplemental Figure S4.3. Strong correlation of gene expression between quantitative

RT-PCR and RNA-Seq data...... 188

Supplemental Figure S4.4. Enriched GO terms in the genes differentially expressed in the tyrR mutant compaired to wild-type E. cloacae...... 189

176

Supplemental Figure S4.5. The paa catabolic operon for phenylacetate degradation in E. cloacae...... 190

177

Supplementary Table S4.1. Bacterial strains and plasmids used in this study.

Strain or Plasmid Relevant characteristics Reference E. coli JM109 Cloning host Promega S17.1λpir Cloning host; TpR SmR recA thi p10 hsdR-M+ RP4:2-Tc:Mu:KmR (2) Tn7λpir A118 Protein expression host; E. coli K-12 lacZ StR (pREP4, pQEtyrR) (3)

E. cloacae UW5 Wild type strain; AmpR (3)

J126 ΔtyrR; AmpR This study Plasmids pKD4 Source of FRT-flanked Km gene; rep R6K AmpR FRT-KmR-FRT (4)

pJQ200SK Low copy number suicide vector; GmR sacB (5) pCP20 Temperature-sensitive replication and induction of FLP-recombinase; (4) ts R R rep pSC101 Amp Cm cI857 λPR pGEM-T easy Cloning vector; AmpR Promega pUCIDT-AMP Cloning vector; AmpR IDT pTUD pGEM-T easy; 1.5 kb insert of tyrR upstream and downstream This study flanking regions. pTUD-Km pTUD; FRT-KmR-FRT This study pJQTUD-Km pJQ200SK; This study

178

Supplementary Table S4.2. Oligonucleotides used in this study. Restriction sites are in bold, fluorescent modifications are in square brackets.

Primer Sequence 5’-3’ Reference TUF-PstI AAAAAACTGCAGGGGATCTGCTCCACAGTCAC This study TUR-XbaI CGAAAAATGGCTCTAGAGGGAAATTCACCGTTTTAAG This study TDF-XbaI GTGAATTTCCCTCTAGACGGTAAAAAGCCTCTGTAAAC This study TDR-SacI AAAAAAGAGCTCTGTATCGCAGGTTGAAGTGG This study PKDP1 GTGTAGGCTGGAGCTGCTTC (4) PKDP2 CATATGAATATCCTCCTTAG (4) TUF GATCTGTTGCGACCGAGTG This study TUR TTTCTACGTGTTCCGCTTCC This study TDF CAAAAGCGCTCTGAAGTTCC This study TDR CCAAATCTACACGCTTCACG This study ICRTIF TCGAACTCAGCAAACAGCAC (3) ICRTIR AGGTTTGCAACGTTCTCCAG (3) recAF2 GCTGGACCCTGTTTATGCTC This study recAR1 GCCTTCGATTTCAGCTTTTG This study proP35 ACTATAAAACCCTCACGCTCG This study proP160 GCAACCACTCCGGGATATC This study eco145 GTCATTCAGTTACCCGCTCAG This study eco269 TCCAGGGTTTTGCTTTCCAG This study aroG576 CATTAAGGTGGCAATTGACGC This study aroG670 TGGTATTCACAATGGCGGAG This study yagU384 ACTCTGGCAAGGTTTACTGG This study yagU518 AAATGCCCCACGATCTCTG This study cpxP356 AGATGTTCCACCTGCTAACG This study cpxP494 CGGGTACTGCTATTGCTACTG This study cpxR209 ACCAGACGCCCGTAATTATG This study cpxR302 GGCTTTGGTAAATAGTCATCCG This study dmpM161 CCGATATGCTGAAACTTGACG This study dmpM294 GTGGTCCTTACAGAGAAAGTGG This study dmpM-F CAACCGTTAATTTCGCATTC This study dmpM-R-FAM [6FAM]GCTACCGGGTACAGTTAAACC This study cpx-F CCTCCGAGGCAGAAATTACG This study cpx-R-FAM [6FAM]AAATCTTTCGTGTCGCGATT This study 179

Supplementary Table S4.3. Sequences of the synthesized cpx and dmpM promoters used for electrophoretic mobility shift assays. Each insert in pUCIDT-AMP is flanked by EcoRI cut sites (bold) and primer binding locations are indicated in italics. The dmpM fragment is flanked outside the EcoRI sites by GC adapters (lower case), which were necessary to improve the synthesis reaction due to low GC content of the target sequence. Putative TyrR boxes (dmpM) and CpxR sites (cpx) are underlined, and mutated nucleotides are in bold lowercase. Insert Characteristics Sequence 5’-3’ dmpM-wt Wild type promoter ctcacttgtagaacggtgatcagcctgtgctctagagcctgatagttgagc gatacacacGAATTCCAACCGTTAATTTCGCATTCAGATATATATAACAAT ATTTACAGGCCACACCCGGAAATGAATTGAAAAACGCATGATCTTTAACGG TATTAGTTACCAGCATTAAAAAACGGTGACTTAATGTCCGGTTTTATTTAC AGGAGTCTGTCTTATGTAACTTAAATGTTACTCCTGTTTTCTGGGAGGGGG CGTTACATATGAAGTAATAAAGGTTTAACTGTACCCGGTAGCTGTAAATAA GAAATTCACATTTTTTAATCCTTCAGGAGCTAATGAATTCtgatcgttgaa gtcgacctacatcgagtgcgcactatcaagagtgttccagtcacgcgat dmpM-mut All three TyrR boxes ctcacttgtagaacggtgatcagcctgtgctctagagcctgatagttgagc mutated gatacacacGAATTCCAACCGTTAATTTCGCATTCAGATATATATAACAAT ATTTAtAGGCCACACCCGGAAATGAATTGAAAAACGCATGATCTTTAACGG TATTAGTTACCAGCATTAAAAAACGGTGACTTAATaTCCGGTTTTATTTAt AGGAGTCTGTCTTATaTAACTTAAATGTTAtTCCTGTTTTCTGGGAGGGGG CGTTACATATGAAGTAATAAAGGTTTAACTGTACCCGGTAGCTGTAAATAA GAAATTCACATTTTTTAATCCTTCAGGAGCTAATGAATTCtgatcgttgaa gtcgacctacatcgagtgcgcactatcaagagtgttccagtcacgcgat cpx-wt Wild type promoter GAATTCTTGTTTAAATACCTCCGAGGCAGAAATTACGACATCAAGGCTAT CTAATCCATGACTTTACGTTGTTTTACACCCCCTGACGCATGTTTGCAGCC TGAATCGTAGACTGTCTCTCGTTGAATCGCGACACGAAAGATTTTGGGAGC AAGTGGAATTC cpx-mut1 CpxR site 1 mutated GAATTCTTGTTTAAATACCTCCGAGGCAGAAATTACGACATCAAGGCTAT CTAATCCATGACTTTACGTTGTTTTACACCCCCTGACGCATGTTTGCAGCC TGAATCaTAGACTGTCTCTttTTGAATCGCGACACGAAAGATTTTGGGAGC AAGTGGAATTC cpx-mut2 CpxR site 2 mutated GAATTCTTGTTTAAATACCTCCGAGGCAGAAATTACGACATCAAGGCTAT CTAATCCATGACTTTACGTTGTTTTACACCaaaTGACGCATGTTTGtAGCC TGAATCGTAGACTGTCTCTCGTTGAATCGCGACACGAAAGATTTTGGGAGC AAGTGGAATTC cpx-mut3 CpxR site 3 mutated GAATTCTTGTTTAAATACCTCCGAGGCAGAAATTACGACATCAAGGCTAT CTAATCCATaAaTTTACGTTGTTTTAtACCCCCTGACGCATGTTTGCAGCC TGAATCGTAGACTGTCTCTCGTTGAATCGCGACACGAAAGATTTTGGGAGC AAGTGGAATTC

180

Supplementary Table S4.4. Genes that were significantly downregulated in the tyrR mutant strain of E. cloacae relative to the wild type. UW5 gene ID Gene Gene description log2FC padj ABT55_00195 nirD Nitrite reductase [NAD(P)H] small subunit -3.22 1.27E-16 ABT55_00205 yhfL Small lipoprotein -2.09 2.31E-11 ABT55_00335 feoC Ferrous iron transport protein C -2.40 9.60E-23 ABT55_00400 glpD Aerobic glycerol-3-phosphate -2.32 3.37E-05 dehydrogenase ABT55_00775 arnB UDP-4-amino-4-deoxy-L-arabinose-- -2.15 4.25E-18 oxoglutarate aminotransferase ABT55_02285 emrD Multidrug transporter -2.14 5.06E-61 ABT55_03070 yhjX Pyruvate-inducible inner membrane protein -3.88 5.64E-17 ABT55_03610 fdhF Formate dehydrogenase H subunit alpha -2.05 1.69E-32 ABT55_04495 arcB Ornithine carbamoyltransferase, catabolic -2.35 1.15E-12 ABT55_04500 arcC Carbamate kinase -3.16 4.25E-12 ABT55_04505 arcA Arginine deiminase -2.91 1.98E-08 ABT55_04870 ABT55_04870 DeoR faimly transcriptional regulator -3.21 3.94E-129 ABT55_04990 yjjB Threonine/serine exporter -2.20 2.06E-19 ABT55_05085 yjjW Putative pyruvate formate lyase activating -2.96 3.60E-13 enzyme ABT55_05090 yjjI DUF3029 family protein, putative glycine -2.05 5.35E-24 radical enzyme ABT55_05100 deoA Thymidine phosphorylase -2.21 2.12E-35 ABT55_07295 fimA Type I fimbriae protein subunit -2.06 3.77E-106 ABT55_07325 fimZ Fimbriae biosynthesis transcriptional -2.55 9.95E-158 regulator ABT55_07800 dmpM S-adenosylmethionine O-methyltransferase -5.36 7.81E-233 ABT55_07870 ABT55_07870 Extracellular serine protease -2.15 8.78E-121 ABT55_07875 ABT55_07875 Hypothetical protein -3.18 3.14E-53 ABT55_08065 budB Acetolactate synthase, catabolic -2.60 4.04E-22 ABT55_08070 budC Diacetyl reductase [(S)-acetoin forming] -2.57 9.16E-31 ABT55_08325 dcuC Anaerobic C4-dicarboxylate transporter -2.64 2.56E-27 dcuC ABT55_08600 speF Ornithine decarboxylase, inducible -2.43 3.55E-37 ABT55_08730 cydA Cytochrome d ubiquinol oxidase subunit 1 -2.19 2.52E-23 ABT55_08735 cydB Cytochrome d ubiquinol oxidase subunit 2 -2.85 9.70E-53 ABT55_08740 cydX Cytochrome bd-I oxidase subunit -2.12 8.45E-15 ABT55_08745 ybgE Cyd operon protein -2.11 1.12E-22 ABT55_09620 hcr NADH oxidoreductase hcr -2.60 5.28E-10 ABT55_09625 hcp Hydroxylamine reductase -2.76 3.47E-15 ABT55_09730 dmsA Anaerobic dimethyl sulfoxide reductase -2.43 1.01E-21 chain A ABT55_09735 dmsB Anaerobic dimethyl sulfoxide reductase -2.41 4.15E-30 chain B ABT55_09760 pflB Pyruvate formate-lyase -2.10 6.06E-30 ABT55_09835 ycbJ Putative phosphotransferase -2.25 1.22E-25 ABT55_10470 efeU Ferrous iron permease efeU -2.91 2.78E-77 ABT55_10475 efeO Ferrous iron transport system protein -3.90 2.35E-254 ABT55_10480 efeB Heme-containing -3.62 1.09E-171 peroxidase/deferrochelatase ABT55_10955 trg Methyl-accepting chemotaxis protein III -2.95 8.50E-136 ABT55_10960 yuxH HDOD domain-containing protein -2.95 1.97E-86

181

ABT55_12585 tcp Methyl-accepting chemotaxis citrate -2.60 1.50E-134 transducer ABT55_13045 pla Omptin family outer membrane protease -2.73 0.00E+00 ABT55_13320 trg Methyl-accepting chemotaxis protein III -2.26 3.59E-33 ABT55_14080 fdnI Formate dehydrogenase, nitrate-inducible, -2.34 3.93E-29 cytochrome b556(fdn) subunit ABT55_14085 fdnH Formate dehydrogenase, nitrate-inducible, -2.67 9.25E-36 iron-sulfur subunit ABT55_14090 fdnG Formate dehydrogenase, nitrate-inducible, -2.38 3.17E-27 major subunit ABT55_14470 tyrR Aromatic amino acid regulatory protein -4.95 0.00E+00 ABT55_15240 narH Respiratory nitrate reductase 1 beta chain -2.29 4.71E-33 ABT55_15385 ychH DUF2583 domain-containing protein -2.25 1.28E-40 ABT55_16160 tap Methyl-accepting chemotaxis protein IV -2.07 4.12E-55 ABT55_16165 tar Methyl-accepting chemotaxis protein II -2.36 2.07E-177 ABT55_16270 dcuB Anaerobic C4-dicarboxylate transporter -2.03 3.54E-23 ABT55_16345 cspB Cold shock-like protein -2.58 7.34E-42 ABT55_16350 ABT55_16350 Cold-Shock Protein -3.11 9.71E-11 ABT55_16500 fliD Flagellar filament capping protein -2.05 8.00E-42 ABT55_16685 ttdT L-tartrate/succinate antiporter -3.02 1.73E-52 ABT55_16690 fumB Fumarate hydratase class I, anaerobic -2.35 1.05E-46 ABT55_16925 ABT55_16925 Hypothetical protein -2.24 2.34E-11 ABT55_17380 gmd GDP-mannose 4,6-dehydratase -2.19 4.10E-36 ABT55_17390 wcaE Colanic acid biosynthesis -2.14 3.73E-05 glycosyltransferase ABT55_17500 matB Fimbrillin -2.03 2.73E-12 ABT55_17855 fruA PTS fructose transporter subunit IIBC -2.03 1.65E-25 ABT55_18110 glpA Anaerobic glycerol-3-phosphate -3.23 1.61E-14 dehydrogenase subunit A ABT55_18115 glpB Anaerobic glycerol-3-phosphate -4.15 1.39E-16 dehydrogenase subunit B ABT55_18120 glpC Anaerobic glycerol-3-phosphate -4.98 3.22E-20 dehydrogenase subunit C ABT55_18205 cheV Chemotaxis protein -2.30 3.07E-201 ABT55_18590 ipdC Indole-3-pyruvate decarboxylase -4.28 1.24E-216 ABT55_20230 hypE Hydrogenase isoenzymes formation protein -2.17 6.96E-23 ABT55_20235 flhA Formate hydrogenlyase transcriptional -2.11 1.17E-22 activator ABT55_21460 trg Methyl-accepting chemotaxis protein III -2.24 4.70E-152 ABT55_22185 tdcC Threonine/serine transporter -2.34 4.34E-06 ABT55_22190 tdcB Threonine dehydratase catabolic -2.85 3.71E-06 ABT55_22195 tdcA HTH-type transcriptional regulator -2.87 1.66E-07 ABT55_22775 yagU Inner membrane protein -3.63 2.52E-230 ABT55_22865 cah Carbonic anhydrase -2.55 5.17E-73

182

Supplementary Table S4.5. Genes that were significantly derepressed in the tyrR mutant strain of E. cloacae relative to the wild type. UW5 gene ID Gene Gene description log2FC padj ABT55_00940 dctA C4-dicarboxylate transport protein 2.27 1.03E-33 Fatty acid oxidation complex alpha ABT55_01055 fadB subunit 2.32 3.05E-44 ABT55_01060 fadA 3-ketoacyl-CoA thiolase 2.27 7.80E-24 ABT55_01650 cpxP Periplasmic protein CpxP 6.03 0 ABT55_01660 cpxA Sensory histidine kinase 2.15 1.50E-21 ABT55_01980 rbsD D-ribose pyranase 3.84 2.15E-154 ABT55_02335 mhpC Alpha/beta hydrolase 2.40 1.57E-25 ABT55_02780 lldD L-lactate dehydrogenase [cytochrome] 2.90 2.43E-38 Putative L-lactate dehydrogenase operon ABT55_02785 lldR regulatory protein 2.53 8.87E-31 ABT55_02790 lldP L-lactate permease 2.85 6.39E-54 ABT55_03585 actP Acetate / glycolate : cation symporter 2.94 2.91E-35 ABT55_03590 yjcH Inner membrane protein 2.73 9.22E-85 ABT55_03595 acs Acetyl-coenzyme A synthetase 2.69 1.84E-29 ABT55_04225 ytfK DUF1107 domain-containing protein 2.05 1.79E-26 Succinate-semialdehyde dehydrogenase ABT55_04835 ssdA [NADP+] 2.69 8.91E-36 ABT55_05285 proP Putative proline/betaine transporter 3.17 0 Aromatic amino acid transport protein ABT55_05725 aroP AroP 5.09 0 ABT55_05730 ABT55_05730 Hypothetical protein 3.73 1.37E-36 ABT55_05940 degP Serine endoprotease 2.22 4.73E-30 Putative ABC transporter ATP-binding ABT55_07220 ybbA protein 3.05 4.91E-203 Putative ABC transporter membrane ABT55_07225 ybbP subunit 3.70 0 Proofreading thioesterase in enterobactin ABT55_08180 entH biosynthesis 2.53 9.46E-46 Phospho-2-dehydro-3-deoxyheptonate ABT55_08825 aroG aldolase, Phe-sensitive 2.75 1.42E-75 ABT55_08920 hutH Histidine ammonia-lyase 2.01 3.03E-25 ABT55_09085 ybiJ DUF1471 domain-containing protein 3.88 6.13E-165 ABT55_09290 yliI Aldose sugar dehydrogenase 2.12 1.97E-44 ABT55_10065 yccA FtsH protease modulator 2.61 1.84E-22 ABT55_10630 yceJ Cytochrome b561-like protein 2 2.39 1.49E-71 ABT55_10915 ycfS (ldtC) L,D-transpeptidase YcfS 5.14 5.31E-225 ABT55_11735 spy Spheroplast protein Y 3.33 2.23E-246 Succinyl-CoA:3-ketoacid-coenzyme A ABT55_11935 lpsI transferase subunit A 2.26 1.67E-36 ABT55_11940 ybhD LysR family transcriptional regulator 2.48 2.24E-18 ABT55_12420 rnfC (rsxC) Electron transport complex protein rnfC 2.04 2.40E-09 ABT55_12645 mdtJ Multidrug/spermidine export protein mdtJ 2.26 4.09E-13 ABT55_12855 uspG Universal stress protein G 2.50 8.48E-31 Phenylacetyl-CoA 1,2-epoxidase, ABT55_12990 paaC structural subunit 2.24 1.67E-25 Putative phenylacetate-CoA oxygenase ABT55_12995 paaD subunit 2.50 8.05E-117 Phenylacetyl-CoA 1,2-epoxidase, ABT55_13000 paaE reductase subunit 2.19 5.71E-07 183

ABT55_13005 paaF 2,3-dehydroadipyl-CoA hydratase 2.57 5.09E-69 Ring 1,2-epoxyphenylacetyl-CoA ABT55_13010 paaG isomerase (oxepin-CoA forming) 2.68 4.59E-268 ABT55_13015 paaH 3-hydroxyadipyl-CoA dehydrogenase 2.55 2.18E-23 ABT55_13020 paaI Phenylacetyl-CoA thioesterase 2.63 2.64E-70 ABT55_13160 ABT55_13160 DUF2878 domain-containing protein 2.82 1.45E-36 ABT55_13165 ABT55_13165 Hypothetical protein 2.53 1.18E-32 ABT55_13330 ydcI LysR family transcriptional regulator 3.24 1.34E-31 ABT55_13885 dsbD Thiol:disulfide interchange protein DsbD 2.63 1.60E-69 ABT55_14525 ycjP Carbohydrate ABC transporter permease 2.29 2.74E-10 ABT55_15850 ftsI Peptidoglycan glycosyltransferase 3.13 7.16E-73 ABT55_15895 htpX Zinc dependent endoprotease 2.92 6.91E-238 ABT55_15985 yebE DUF533 family inner membrane protein 6.16 0 ABT55_16305 tyrP Tyrosine-specific transport protein 4.31 7.14E-131 Holocytochrome c synthetase - ABT55_17605 dsbE (ccmG) thiol:disulfide oxidoreductase 2.66 2.97E-33 ABT55_18015 eco Serine protease inhibitor ecotin 3.04 3.95E-186 ABT55_18135 tctE Sensor histidine kinase 2.53 1.98E-41 ABT55_18140 tctD Transcriptional regulator 2.15 3.87E-93 Tripartite tricarboxylate transporter ABT55_18145 tctC substrate binding protein 3.28 2.47E-50 Triartite tricarboxylate transporter family ABT55_18150 tctB protein 3.12 1.30E-22 Tripartite tricarboxylate transporter ABT55_18155 tcta permease 2.06 6.63E-65 ABT55_18420 cvpA Colicin V production protein 3.55 0 ABT55_18515 sixA Phosphohistidine phosphatase 2.08 1.83E-33 ABT55_18830 acrD Putative aminoglycoside efflux pump 2.03 2.29E-81 ABT55_19015 iolC 5-dehydro-2-deoxygluconokinase 2.15 1.34E-17 ABT55_19020 yjhC Putative oxidoreductase 2.17 1.69E-24 ABT55_19435 kgtP (pcaT) α-ketoglutarate:H+ symporter 2.19 1.14E-30 Fused chorismate mutase/prephenate ABT55_19495 tyrA dehydrogenase 5.03 0 Phospho-2-dehydro-3-deoxyheptonate ABT55_19500 aroF aldolase, Tyr-sensitive 6.04 0 ABT55_19895 yqaE Pmp3 family membrane protein 3.25 1.09E-130 Hydrogenase maturation protein, ABT55_20140 hypF carbamoyltransferase 2.58 0 ABT55_20755 yqeF Acetyl-CoA C-acetyltransferase 2.10 2.59E-19 Ferric transport system permease protein ABT55_20810 fbpB fbpB 2.93 3.72E-25 Fe3+/spermidine/putrescine ABC ABT55_20815 potA transporter ATP-binding protein 2.42 2.22E-19 ABT55_21175 mhpC Alpha/beta hydrolase 2.09 1.60E-225 ABT55_21185 dgoT MFS transporter 2.96 5.33E-16 Pyridine nucleotide-disulfide ABT55_21190 thcD oxidoreductase 3.32 2.54E-34 ABT55_21195 mcpII Glyoxalase 2.61 4.83E-17 ABT55_21200 pcaR IclR family transcriptional regulator 2.48 2.12E-15 ABT55_21205 ABT55_21205 SDR family oxidoreductase 2.68 6.63E-25 Aromatic ring-hydroxylating dioxygenase ABT55_21210 vanA subunit alpha 2.72 5.44E-22 ABT55_21215 ABT55_21215 Hypothetical protein 2.48 8.35E-29 ABT55_21220 ndoA Non-heme iron oxygenase ferredoxin subunit 2.44 1.32E-17 184

Supplementary Table S4.6. HHPred results with DmpM query sequence. The probability column is the percent probability of a database match being homologous to the query sequence. The E-value is defined similar to BLAST (as the expected number of false positives per database search with a score at least as good as the score of this sequence match) and identity is the percent sequence identity. PDB Product Species Probability E-value Identity % 3GWZ mitomycin 7-o-methyltransferase Streptomyces lavendulae 100.0 5.20E-30 27 3LST orsellinic acid methyltransferase Micromonospora 100.0 1.50E-30 36 echinospora 5EEG carminomycin-4-o- Streptomyces peucetius 99.95 1.10E-27 23 methyltransferase 1FP2 isoflavone o-methyltransferase Medicago sativa 100.0 4.50E-32 25 5XOH bergaptol o-methyltransferase Peucedanum 100.0 1.80E-31 25 praeruptorum 3P9C caffeic acid o-methyltransferase Lolium perenne 99.95 9.30E-28 23

185

800 600 400 [0-800] WT 200 B A 0 800 600 16 400 [0-800] WT 200 0 800 600 WT 400 [0-800] WT 200 0 800 600 12 400 [0-800] WT 200 0

5' 3' 3' ipdC akr 5' 3.964 mb 3.965 mb 3.966 mb 3.967 mb 3.968 mb 800 8 600 [0-800] 400

tyrR 200

0 (ug/ml/OD) IAA 800 600 400 [0-800]

tyrR 200 0 tyrR 800 4 600 Δ 400 [0-800]

tyrR 200 0 800 600 400 [0-800]

tyrR 200 0 0 WT ΔtyrR

Supplemental Figure S4.1. TyrR is necessary for expression of ipdC and production of IAA. (A) Histogram of RNA-Seq read coverage of the E. cloacae genome per base. RNA-Seq reads in the wild type strain (blue) indicate transcription beginning at 5’ end of the ipdC gene and covering the length of the coding sequence, which is absent in the tyrR mutant (red). The result was similar for all biological replicates, highlighted in yellow. The maximal depth of read coverage was the same across all samples (in square brackets). (B) Effect of the tyrR deletion on IAA production in M9 minimal medium alone (grey bars) or supplemented with 1 mM L-tryptophan (black bars). Error bars indicate the standard error of the mean of three biological replicates.

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STOP TyrR Box II Eco_K12 1 TGAACCGGAAACGAAACCCTCATCCTAAT----AAAGAGTGACGTAAATCACACTTTACA 60 Sen_LT2 1 TGAACCCGAGGTCAAACCGTCAATCTAAAGTAAAAAATGTGATGTTCTGCAAACTTTACT 60 Ecl_UW5 1 CGAACCGGAGAGTAAACCTTCCGCGTGAAGTGG------60 Ecl_WSU1 1 CGAACCGGAGAGTAAACCTTCCGCGTGAAGTGG------60 ***** ** ***** ** * *

TyrR Box I Eco_K12 61 GCTAACTGTTTGTTTTTGTTTCATTGTAATGCGGCGAGTCCAGGGAGAGAGCGTGGACTC 120 Sen_LT2 61 GCTAATTGGCTGTTTTTGAACTACTGTAATGCTGGCGCTCCACATCAAATGAGTGGCGTC 120 Ecl_UW5 61 ------AGTT 120 Ecl_WSU1 61 ------AGTT 120 * IHF -35 Eco_K12 121 GCCAGCAGAATATAAAATTTTCCTCAACATCATCCTCGCACCAGTCGACGACGGTTTACG 180 Sen_LT2 121 GCCAGCAGAACGAAAAATTTTCGTGCTCATCCTCTTTTCGTCAGTCGACGAAAGATTGCG 180 Ecl_UW5 121 GCCAGCAATATTAAAAAATTTCTCTATCTTTACTCAGCCGCCAGTCGACGAAAGATTAAG 180 Ecl_WSU1 121 GCCAGCAATATTAAAAAATTTCTCTATCTTTACTCAGCCGCCAGTCGACGAAAGATTAAG 180 ******* * **** **** * * * ********** * ** *

-10 +1 MET Eco_K12 181 CTTTACGTATAGTGGCGACAATTTTTTTTATC---GGGAAATCTCAATGATCAGTCTGAT 240 Sen_LT2 181 CTTTACGTATAGTGGCGGCAATTTTTTTTGTATCAGGAAATAATTAATGATCAGTCTGAT 240 Ecl_UW5 181 CTTTCCGTATAGTGGCGGCAATTTTTTGCA-TCCGGGAAATTTTCAATGATCAGTCTGAT 240 Ecl_WSU1 181 CTTTCCGTATAGTGGCGGCAATTTTTTGCA-TCCGGGAAATTTTCAATGATCAGTCTGAT 240 **** ************ ********* ** ** * ***************

Supplemental Figure S4.2. Multiple sequence alignment of the folA promoter region of four Enterobacteriacea strains. The locations of TyrR and IHF binding sites in E. coli are boxed, and the promoter elements underlined. Protein coding sequences are in bold, with the kefC and folA stop and start codons, respectively, labeled above. Conserved nucleotides are indicated with an asterisk. E. coli K12 (Eco K12), S. enterica sv. Typhiimurium str. LT2 (Sen LT2), E. cloacae UW5 (Ecl UW5), E. cloacae WSU1 (Ecl WSU1).

187

6 R = 0.9753 4 P = 1.82E-4 2 0 -2 -4 Seq (log2 Fold Change) Fold (log2 Seq - -6 RNA -8 -6 -4 -2 0 2 4 6 8

RT-qPCR (log2 Fold Change)

Supplemental Figure S4.3. Strong correlation of gene expression between quantitative RT-PCR and RNA-Seq data. Comparison of log2-fold changes for seven genes, each represented by a blue circle. The Pearson correlation coefficient (R), p-value (P) and the line of best fit from and ordinary least-squares linear regressions are indicated. A strong correlation is shown between the expression levels measured by RNA-Seq and quantitative RT-PCR.

188

Up Regulated Genes Down Regulated Genes

organic carboxylic acid formate monocarboxylic chorismate hydroxy transmembrane alditol lipid compound acid metabolic metabolic transport metabolic energy positive xenobiotic response to oxidation transport aerotaxis phosphate process process anaerobic formate chemotaxis glycerol−3−phosphate aromatic anion process taxis metabolic metabolic process catabolic process xenobiotic stimulus amino acid respiration oxidation glycerol−3−phosphate processmetabolism organic organictransport hydroxytransport oxidation−reduction monocarboxylicsingle−organism process response to response to acid metabolic response process compound endogenous nitrogen thymidine 'de novo' GDP−L−fucose aromatic metabolicacid metabolism small aspartate GDP−L−fucose movement energy taxis metabolic transport to acid metabolic biosynthetic family stimulus process response process alcohol molecule compound process cellular oxidation−reduction acetate organophosphate of cell or process compound catabolic process amino acid aromatic process catabolic metabolic subcellular chemical to small catabolic catabolic response to component cellular response C4−dicarboxylate response to amino acid process molecule catabolic process process oxygen−containing localization transport acetyl−CoA metabolic glycolate response to response to xenobiotic stimuluschemical family acetate tellurite processanaerobicorganic acidcarbohydrate respirationsmall aerobic response compound BP 3−phenylpropionate biosynthetic process to organic localizationof cell chemical biosynthetic transport transport transport derivative molecule electron process metabolic process process from substance nitrate metabolic catabolic catabolic transport stimulus process acetate to amino response to L−serine stimulus organic cyclic cellular process iron ionof cell fatty acid metabolic process process chain external transport post−translational anaerobic cellular response protein threonine response protein allantoin acid compound protein electron arginine monocarboxylic stimulus to cadmium ion transport metabolic autophosphorylation deacetylation process acetylation metabolic catabolic deiminase acid metabolic polyamine to benzene−containing L−threonine transport process catabolic process process pathway process chain biosynthetic nitrogen stimulus catabolic generation single−organism compound polyamine locomotion cycle process protein response arginine signaling process monocarboxylic to process to diol process metabolism single−organism cellular of drug metabolism stimulus metabolic biosynthesis autophosphorylation propionate cellular process metabolic single−organism amide regulation of polyamine acid catabolic cellular lipid metabolic precursor process process catabolic biosynthetic process lipid export threonine process reactive regulation negative short−chain process metabolites response organic process modification cellular protein ornithine glycol nitrogen cell catabolic regulation of fatty acid hydroxy lactate modification catabolic auxin to of apoptotic endopeptidase metabolic metabolic and energy species communication compound metabolic cellular carbohydrate stimulus process activity catabolism metabolism process oxidation process process process process process catabolic process metabolism metabolism

intrinsic component plasma intrinsic respiratory of the cytoplasmic membrane component of side of the plasma external side chain respiratory of plasma integral component integral component glycerol−3−phosphate fatty acid oxidoreductase membrane chain membrane of plasma membrane of membrane beta−oxidation dehydrogenase periplasmic space complex complex intrinsic component of multienzyme the cytoplasmic side high−affinity cell complex plasma of the plasma membraneiron integral component of plasma membrane oxidoreductase complex membrane CC permease part complex integral cytochrome macromolecular component of plasma complex intrinsic component formate complex catalytic plasma membrane plasma membrane dehydrogenase membrane of membrane complex cell division site complex protein cell cell partcell part cell surface membrane complex periphery

CoA glycolate carboxylic acid phosphohistidine metal thymidine solute:proton protein kinase molybdenum pyrimidine−nucleoside phosphorylase phosphatase glycerol−3−phosphate formate oxidoreductase activity, phosphorylase activity transmembrane symporter transmembrane 3−oxoadipyl−CoA hydrolase acting on the CH−OH ion activity symporter 3−deoxy−7−phosphoheptulonate activity ion binding transmembrane synthase activity dehydrogenase dehydrogenase group of donors, quinone thymidine transporter thiolase activity proteinbinding kinase molybdenumbinding signaling transporter activity activity or similar compound as phosphorylase activity activity activity (NAD+) activity acceptor receptor 3−deoxy−7−phosphoheptulonate CoA hydrolase ion binding activity activity activity binding amino carbamate ornithine synthase activity enzyme cation kinase carbamoyltransferase activity acid activity active protein thiolester binding activity secondaryglycolate active transmembrane 1,4−dihydroxy−2−naphthoyl−CoA oxidoreductase binding transmembrane acetyl−CoA deacetylase hydrolase thioesterase activity binding C−acyltransferase transmembrane transporter C−acyltransferase activity, acting on NADH activity activity activity glycerol−3−phosphatedehydrogenase transporter activity other nitrogenous C4−dicarboxylate heme transporter activity activity activity compounds as donors transmembrane signal activity acetate dehydrogenase activity bindingheme transporter transducer transmembrane formate electron carrier MF organic hydroxy transporter selenate oxidoreductase activity tetrapyrrolebinding activity tellurite uptake intramolecular anion dehydrogenase oxidoreductase activity, acting on compound transmembrane 2 iron, activity transmembrane activity 3−hydroxybutyryl−CoA serine−type reductase activity, acting diphenols and binding tellurite transporter lyase activity epimerase activity (quinone) on NAD(P)H related substances coenzyme transmembrane activity intramolecular endopeptidase 2 sulfur transporter transmembrane activity activity as donors molecular transporter transporter inhibitor cluster coenzymebinding activity dodecenoyllyase−CoA activity peptidase oxidoreductase activity, dimethyl transducer activity oxidoreductase activity activity binding acting on the aldehyde or acetoin oxidoreductase racemase and epimerase regulator sulfoxide activity, acting on binding activity delta−isomerase oxo group of donors, with diphenols and related dehydrogenase activity 3−hydroxybutyryl−CoA activity, acting on hydroxy substances as donors, a quinone or similar reductase oxygen as acceptor activity dehydrogenase activity acids and derivatives activity cofactor compound as acceptor activity 3−hydroxyacyl−CoA 3−hydroxyacyl−CoAactivity oxidoreductase metal cluster activity, acting 3 iron, binding dehydrogenase on the CH−NH acetate−CoA GDP−mannose activity oxidoreductase group of donors enoyl−CoA isomerase ornithine dehydrogenase activity ligase 4 sulfur binding 4,6−dehydratase arginine activity, acting on the activity iron−sulfur cluster binding decarboxylase activity deiminase CH−NH group of donors, hydratase activity cluster activity inhibitor activity NAD or NADP as acceptor binding activity

Supplemental Figure S4.4. Enriched GO terms in the genes differentially expressed in the tyrR mutant compared to wild-type E. cloacae. REVIGO TreeMap visualization summary of enriched GO terms at p < 0.05 determined by GOSeq for up- and downregulated genes in the tyrR mutant. GO terms are presented according to the three main classifications: biological processes (BP), cellular components (CC) and molecular function (MF). Semantically similar GO terms are grouped together by colour (SimRel = 0.7) (1) and box size correlates to the -log10 p-value calculated by GOSeq.

189

Pz Pa Px A paaZ paaA paaB paaC paaD paaE paaF paaG paaH paaI paaJ paaK paaX paaY E. coli K12

E. cloacae

1 kb

300 B 250 200 150 WT 100 50 0

tynA ABT55_12975 paaA paaB ABT55_12990ABT55_12995 ABT55_13000 ABT55_13010 ABT55_13015 ABT55_13025 ABT55_13030 ABT55_13035 ABT55_13040 ABT55_13045 ABT55_13050

tynA paaZ paaA paaB paaC paaD paaE ABT55_13005paaF paaG paaH ABT55_13020paaH paaJ paaK paaX paaY ABT55_13050 ABT55_13045

5' 3' 3' 5' 2.735 mb 2.74 mb 2.745 mb 300 250 200 150

tyrR 100 50 0 C MET Eco_K12 CCAGGTACCGGATAAGAAACTGGCTAACTGCTGCATCGCTACTCTCCAGATGTTTCACAT 60 Ecl_WSU1 CCAGGCACCGGACAAGAAGCTGGCTAACTGCTGCATCGCTTATCTCCAAGTGTTACGTAA 60 Ecl_UW5 CCAGGTGCCGGACAAGAAGCTGGCTAACTGCTGCATCGCTTATCTCCAAGTGTTACGTAA 60 ***** ***** ***** ********************* ****** **** * *

+1 Pz PaaX box Eco_K12 TTCTGTTGCTAATAGTTAAATCGCGAATCATAAAAAGCAAAGGATCTTTTAACGAAATGT 120 Ecl_WSU1 TTTCACTATCAATAGTTTAATCATGAATCATAAAAAGCAAGCTGAGTTTTAATGAATTGT 120 Ecl_UW5 TTTCACTATCAATAGTTTAATCATGAATCATAAAAAGCAAGCTGAGTTTTAATGAATTGT 120 ** * ******* **** **************** ****** *** ***

IHF Eco_K12 TAACTATGCGATCTGTATAGCAACTGCGGAAAACATTAATGCACTGATAAATAATGATTT 180 Ecl_WSU1 TAACATTGTGATCCAGCGTAATGAAGTGGAAGGCCGGAAGC-CCCGTTATGCAGGCCTTG 179 Ecl_UW5 TAACATTGTGATCCAGCGTAATGAAGTGGAAGGCCGGAAGC-CCCGTTGTGCAGGCCTTG 179 **** ** **** * **** * ** * * * * **

CRP -35 Eco_K12 ATAAAAATAGGGTGCGAAATCCGTCACAGTTCAAACATACAAAATTTGTGATTTTACTTA 240 Ecl_WSU1 TGCAAAAAGCCTTGCGAGCACCATCACAAAATAGGGAAACATTTCTTGGGGTTGTATTTA 239 Ecl_UW5 TGCAAAAAGCCTTGCGAGCACCATCACAAAATAGGGAAACATTTCTTGGGGTTATATTTA 239 **** ***** ** ***** * * *** *** * ** ** ***

-10 +1 Pa PaaX box Eco_K12 ACTATTGTGTAACTTTCATAAAACAATGTGATTCGTGT-TTTTAATTAATTCACGAAAAC 299 Ecl_WSU1 ACCTTTGTGTAACTTTTAAGAAACGAAGTGATTCAGTATTTCACTTTAGGTGGTAAATAA 299 Ecl_UW5 ACCTTTGTGTAACTTTTAAGAAATGAAGTGATTCAGTATTTCACTTTAAGTGGTAAATAA 299 ** ************ * *** * ******* ** *** * ** *

MET Eco_K12 TGGAATCGTAAAGGTGATGACGTGACCCAAGAAGAACGCTTTGAGCAACGGATAGCCC 357 Ecl_WSU1 TCGAATCATAAAGGTGATGACGTGACGCAAGAACAACGCTTTGAGCAGCGCATAGCGC 357 Ecl_UW5 TCGAATCATAAAGGTGATGACGTGACGCAAGAACAACGCTTTGAGCAGCGCATAGCGC 357 * ***** ****************** ****** ************* ** ***** * Supplemental Figure S4.5. The paa catabolic operon for phenylacetate degradation in E. cloacae. (A) Genome synteny and organization of the paa genes of E. cloacae and E. coli K12. Genes encoding the ring hydroxylating complex are in red (paaABCDE), the beta-oxidation enzymes in green (paaFGHIJ), the phenylacetate-CoA ligase in orange (paaK), the transcriptional repressor and thioesterase in blue (paaX), and a ring opening enzyme in purple (paaZ). (B) RNA-Seq reads mapped to the paa operon in the wild-type (blue) and tyrR mutant (red). Read coverage is consistent across the length of the operon. (C) Multiple sequence alignment of the paaZ and paaA promoters of E. coli K12, E. cloacae UW5 and E. cloacae WSU1. The locations of PaaX, IHF and CRP binding sites are boxed and the promoter elements are in bold. Start codons for paaZ and paaA are in bold. Conserved nucleotides are indicated with an asterisk.

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4.8.3 References

1. Schlicker A, Domingues FS, Rahnenfüher J, Lengauer T. 2006. A new measure for functional similarity of gene products based on Gene Ontology. BMC Bioinfo. 7:302 2. Simon R, Priefer U, Pühler A. 1983. A broad host range mobilization system for in vivo genetic engineering: transposon system mutagenesis in gram negative bacteria. Nat Biotechnol 1:784-791. 3. Ryu RJ, Patten CL. 2008. Aromatic amino acid-dependent expression of indole-3- pyruvate decarboxylase is regulated by TyrR in Enterobacter cloacae UW5. J Bacteriol 190:7200–7208. 4. Datsenko KA, Wanner BL. 2000. One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc Natl Acad Sci USA 97:6640– 6645. 5. Quandt J, Hynes MF. 1993. Versatile suicide vectors which allow direct selection for gene replacement in Gram-negative bacteria. Gene 127:15-21.

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Chapter 5 General Discussion and Conclusions

The physiological responses of a bacterium to changing environmental conditions rely on the appropriate regulation of gene expression. Various membrane-bound and cytoplasmic sensors detect changing biotic and abiotic conditions and influence transcription factor activity to activate or repress genes and modulate many cellular functions. The entire genome must be tightly controlled at all times to avoid unnecessary energy expenditure for protein synthesis. The view that bacterial gene expression requires only a promoter, RNA polymerase, and possibly a single transcription factor was revised in light of the finding that bacteria employ sophisticated transcriptional responses involving cross talk of multiple transcription factors, small non-coding RNAs, and condensation of the bacterial nucleoid structure (1, 2). New technologies such as RNA- sequencing allow monitoring of thousands of transcripts simultaneously, and ChIP- sequencing may reveal transcription factor binding sites across the entire chromosome.

This genome-wide view has rapidly accelerated regulon discovery compared to earlier studies with a single gene focus. It has revealed that some transcription factors have highly flexible regulons that are reflective of the diverse environments inhabited by bacterial species. These new technologies supplement in silico methods for transcription factor binding site and regulon prediction.

TyrR is a dual acting transcription factor that is known in E. coli to regulate aromatic amino acid biosynthesis and uptake (3). TyrR is found in the

Enterobacteriaceae family of the gammaproteobacteria having been acquired around the divergence between the Pseudomonaceae and the Vibrionaceae/Pasteurellaceae (Figure

192

5.1A) (4). It is a member of the ATPases associated with various cellular activities

(AAA+) group of bacterial enhancer binding protein (bEBP) transcription factors such as

NtrC, which activate sigma-54 dependent promoters (5). However, TyrR lacks a critical

GAFTGA motif required for bEBPs to contact sigma-54 and instead activates sigma-70 dependent promoters (Figure 5.1B) (3, 5-7). TyrR may be derived from PhhR, which retains a GAFTGA-like motif and controls genes for aromatic amino acid catabolism rather than biosynthesis and transport despite sharing an identical DNA binding sequence

(4). While many genes of the TyrR regulon are conserved in gammaproteobacteria, some species-specific differences are observed; for example, ipdC of E. cloacae, and tpl of C. freundii (8). These genes are both involved in secondary metabolism of aromatic amino acids. TyrR is required for the activation of ipdC, an important gene for the biosynthesis of the plant hormone indole-3-acetic acid (IAA) (9, 10). This protein contributes to root growth-promoting capabilities of the bacterium on host plants (11). Therefore, TyrR regulates bacterial genes with important roles in environmental interactions, in addition to amino acid biosynthesis and uptake.

5.1 TyrR: Beyond amino acid synthesis

In the second chapter of this thesis the regulation of the genes ipdC and akr was investigated in E. cloacae UW5 and the mechanisms of TyrR control were determined using a combination of gene expression analysis and direct protein-DNA interaction assays. The activation of ipdC by TyrR fits with known transcriptional activation of other

TyrR controlled genes. By binding to a single strong TyrR box upstream of the ipdC -35 promoter element, TyrR strongly activates expression in conjunction with phenylalanine,

193

tyrosine and tryptophan (10). Similarly located TyrR boxes in the promoters of mtr, tyrP and folA also induce transcription in response to tyrosine and/or phenylalanine (12-15).

This occurs via class I transcriptional activation involving contacts between DNA-bound

TyrR, and the RNA polymerase alpha subunit carboxy-terminal domain (16).

The enzyme indolepyruvate decarboxylase, encoded by the ipdC gene catalyzes the rate-limiting step in IAA biosynthesis from tryptophan via the so-named indolepyruvate pathway (Figure 1.1, Chapter 1). TyrR is known in E. coli to primarily respond to tyrosine and phenylalanine as cofactors, each of which alter the oligomeric status of TyrR or its relative affinity for DNA elements, and influence the activity of TyrR at a given promoter. Although tryptophan is the precursor for the indolepyruvate pathway, it is not the strongest cofactor for TyrR induction of ipdC expression; rather, phenylalanine has the strongest effects (10). While tryptophan is a cofactor for TyrR, studies suggest that it is less important for modulating promoter activity than the other two aromatic amino acids (13). The tryptophan repressor TrpR is the primary regulator of genes involved in tryptophan metabolism in E. coli (17). The two genes co-regulated by TyrR and TrpR, mtr and aroL, are negatively regulated by TrpR in response to tryptophan (18, 19). In the absence of a trpR gene, TyrR activates mtr transcription (13). Therefore, tryptophan is a cofactor for TyrR in vivo, but it seems to be less important than tyrosine and phenylalanine for regulation.

The regulation of ipdC by TyrR may be conserved among Enterobacteriaceae that possess an ipdC gene. Examination of the ipdC promoter regions of several

Enterobacteriaceae revealed a predicted strong TyrR binding site. In contrast, in

Azospirillum, a member of the Rhodospirillaceae, the ppdC gene encoding 194

phenylpyruvate decarboxylase, which was reclassified from an indolepyruvate decarboxylase to phenylpyruvate decarboxylase does not appear to be regulated by TyrR

(20). Rather, ppdC of A. brasilense appears to be activated in response to IAA by a bacterial auxin-responsive element found in the promoter, although evidence for involvement of a transcription factor is not yet present (21). While the differences in enzyme activity of indolepyruvate decarboxylase and phenylpyruvate decarboxylase are due to amino acid substitutions (20), they are further underscored by differences in transcriptional regulation; regulation of ipdC is controlled by the substrate of the pathway, while the latter is controlled by the product. These differences may not be surprising given the evolutionary distance between the Enterobacteriaceae of the gammaproteobacteria lineage and the Rhodospirillacaeae of the alphaproteobacteria (22).

Therefore, at least in the Enterobacteriaceae, it appears TyrR has evolved to take control of ipdC expression and IAA production. The lack of an ipdC gene in E. coli is one example of the evolution of the TyrR regulon in this family.

This study also identified the akr gene, which is divergently transcribed from ipdC, as a member of the TyrR regulon. TyrR downregulates akr transcription by binding a weak TyrR box located in the promoter region of akr, independently of aromatic amino acid cofactors (10). While the adjacent strong box was required for TyrR binding in vitro, it does not seem to play a role in akr regulation in vivo. It is possible that additional DNA binding proteins contribute to regulation of akr in the cell. Although the weak TyrR box fits the consensus sequence for the binding site, the spacing of the palindromic arms was atypical. The orientation of a regular TyrR box consists of two inverted palindromic arms

(left, L; and right, R) each six base pairs long and separated by six bases (L > N6 < R). 195

However, the weak box has a double right arm organization (L > N2 N2 < R or

L > N8 < R. Either configuration would cause TyrR to bind the DNA with atypical spacing of the palindromic arms. This is the first time such DNA binding has been described for TyrR and raises the possibility that additional TyrR boxes exist in the genome with such atypical spacing. ‘Fuzziness’ in DNA-binding motifs is an important mechanism regulating the level of control that a transcription factor exerts at a given promoter (1). When multiple transcription factors bind a promoter, their respective binding motifs naturally become fuzzier due to overlapping sequence requirements. This allows a promoter to accumulate information from multiple inputs which may dictate transcription levels in a complex and variable response (1). The weak interaction of TyrR with the noncanonical weak box in the akr promoter is consistent with the observed weak downregulation of the gene.

In Enterobacteriaceae with an ipdC gene, an akr orthologue is found divergently transcribed from ipdC and with both TyrR binding sites conserved in the intergenic region. A BLAST search using the E. cloacae akr polypeptide as a query sequence returned the gpr gene from E. coli encoding L-glyceraldehyde 3-phosphate reductase as a close match (59.8% identity, 72.8% similarity) (23). The promoter of the E. coli gene does not contain apparent TyrR boxes. This suggests that in species with an ipdC gene, the simultaneous transcriptional control of ipdC and akr is important.

Aldo-keto reductases, including that encoded by the E. cloacae akr, share a conserved structural fold despite having low sequence conservation. In general, they catalyze the NADPH-dependant reduction of carbonyl-containing compounds that are 196

often cytotoxic (24, 25), although due to their promiscuous substrate specificity it is difficult to assign physiological functions to most aldo-keto reductases. The catalytic function and the role, if any, that the E. cloacae aldo-keto reductase plays in IAA biosynthesis and plant-bacterial interactions is not yet known. One possibility to be investigated is that it detoxifies toxic by-products of tryptophan biosynthesis. The final step in tryptophan biosynthesis is the condensation of L-serine and indoyl-glycerol 3- phosphate to yield L-tryptophan and D-glyceraldehyde 3-phosphate (Figure 5.2A). Non- enzymatic or enzymatic racemization of D-glyceraldehyde 3-phosphate yields L- glyceraldehyde 3-phosphate (Figure 5.2B), which is bactericidal (23, 26).

Triosephosphate isomerase normally converts D-glyceraldehyde 3-phosphate to hydroxyacetone phosphate which then enters the central carbon metabolic pathway

(Figure 5.2B) (27). The high affinity of the E. coli gpr gene product for L-glyceraldehyde

3-phosphate strongly suggests that this is the natural substrate of the enzyme, despite having activity towards other molecules (23, 28). Thus, the E. coli enzyme and its orthologue in E. cloacae may reduce accumulation of the toxic product during tryptophan biosynthesis. Since tryptophan does not have a noticeable effect on akr expression, it is possible that low levels of the reductase are synthesized constitutively to ensure the enzyme is available for detoxification.

5.2 Assembly of the E. cloacae genome

The research described in Chapter 2, which examined the mechanism of regulation of two newly-identified TyrR controlled genes in E. cloacae UW5, revealed that the

TyrR regulon was more diverse than previously thought. Delineation of the TyrR regulon

197

using a whole-genome approach first required the genome of E. cloacae to be sequenced and assembled to allow mapping of RNA-seq data (Chapter 3); the completed genome was deposited in the NCBI Genbank database (29). The 4.9 Mbp E. cloacae genome has an average GC content of 54% and contains 4,496 annotated protein coding genes and eight ribosomal RNA operons. Complete pathways for glycolysis, gluconeogenesis, the

Entner-Doudoroff pathways, and pentose phosphate pathways are present. Characteristic of many E. cloacae complex members, the genome contains genes encoding multidrug resistance mechanisms including many transporter and efflux pumps, heavy metal toxicity resistance proteins, and beta-lactamases conferring resistance to beta-lactam antibiotics.

Present in the genome are genes and pathways that may confer colonization or survival advantages to E. cloacae in the rhizosphere. These include genes for siderophore biosynthesis that allow the bacterium to acquire iron from the soil, a full complement of chemosensory proteins that enable the bacterium to detect nutrients and other chemicals in the environment, and flagellar machinery that confers motility. Multiple pathways are found for the metabolism of aromatic compounds, which allow plant-associated bacteria to exploit the abundant aromatic molecules found in the rhizosphere as carbon and energy sources. Organic and inorganic sulphur and phosphate assimilatory mechanisms are present, which allow the bacteria to use available sources of these essential elements or liberate them from organic molecules. Finally, E. cloacae has a robust capacity for aerobic and anaerobic carbohydrate metabolism. Pathways exist for utilization of mono-, di-, and oligosaccharides, organic acids and sugar alcohols. The ability to exploit multiple carbon sources is correlated with increased rhizosphere fitness in many species of 198

bacteria. The extensive set of monosaccharide uptake and metabolic genes would allow the E. cloacae to use these sugars that are often a major component of plant root exudates.

Assembly and annotation of the E. cloacae genome made it possible to view the metabolic capability of the organism and to study the bacterium on a genome wide scale.

The genome sequence may now serve as a template for primer design to amplify specific sequences, as a query to search for homologous genes, and as a reference sequence for genome-wide sequencing projects. Furthermore, the genome may be mined via bioinformatic tools, perhaps as part of a larger meta-genomic analysis. The relative ease by which bacterial genomes may now be sequenced and assembled is in contrast with more laborious genome assembly methods of only a few years prior to the completion of this thesis. No longer is sequencing restricted to large laboratory collaborations, but may be used as a teaching tool in university classrooms where an E. coli genome can be sequenced and assembled in a single day.

5.3 A more expansive role for TyrR?

A classic technique used to study the function of a gene is to inactivate it through genetic manipulation and examine the resulting phenotype (30). Building upon the initial investigation of TyrR regulation of ipdC and akr in Chapter 2, and sequencing of the E. cloacae genome in Chapter 3, the goal of Chapter 4 was to delineate the TyrR regulon by generating a tyrR knockout strain of E. cloacae and comparing the genome wide transcription profile to a wild-type strain via RNA-sequencing (31). Markerless deletion removed the entire tyrR coding sequence, and total RNA was extracted from wild-type

199

and tyrR mutants grown in LB medium. Analysis of the transcriptome data revealed over

155 genes with significant differential expression between the two genetic backgrounds.

This included 75 genes upregulated and 80 genes downregulated in the tyrR mutant vs. the wild type strain. Many TyrR regulon members (aroF, aroG, tyrP, aroL, mtr, tyrB) displayed up- or down-regulation of transcription in the tyrR mutant E. cloacae consistent with previously published findings in other bacteria (3). Validation of expression of several genes by RT-qPCR gives further confidence that the total RNA-seq data may be used to infer gene expression. The genes impacted by TyrR have diverse functional roles in the cell, some of which are xenobiotic catabolism, aromatic catabolism, carboxyl compound breakdown, anaerobic respiration, formate oxidation and chemotaxis. Many of these genes are likely to be indirectly regulated by TyrR; for example, their expression may be altered in response to changes in intracellular metabolites brought upon by enzymes directly controlled by TyrR, or by other transcription factors that are regulated by TyrR.

The strongest downregulated gene in the tyrR mutant RNA-seq dataset encoding a predicted SAM-dependent methyltransferase, is a newly identified, confirmed member of the TyrR regulon. The dmpM gene is repressed more than 30-fold in the tyrR mutant, and when measured by RT-qPCR is strongly activated by TyrR in conjunction with tyrosine and to a lesser extent phenylalanine. Two TyrR boxes located upstream of the dmpM promoter are bound by TyrR in vitro. These boxes are spaced upstream of the promoter consistent with boxes in other TyrR activated promoters. Therefore, the evidence demonstrates that dmpM is directly controlled by TyrR. It is difficult to assign biological function to SAM-dependent methyltransferases due to low conservation of amino acid 200

sequence and a wide range of substrates recognized by these enzymes. Structurally, the

DmpM polypeptide shares features with a class of plant oxygen-methyltransferases which act upon a number of substrates derived from tryptophan and phenylalanine. Given the strong induction of the gene in response to aromatic amino acids, the evidence suggests the E. cloacae enzyme may act upon one or both of these amino acids, their metabolic precursors, or possible derivatives.

Many genes that are differentially expressed in the tyrR mutant and wild type strains are also members of the CpxR envelope stress response regulon and two genes of the cpx operon, cpxP and cpxA, were strongly upregulated in the mutant. Despite the lack of obvious TyrR boxes in these promoters, we investigated TyrR transcriptional control of these genes. Measurement of cpxR and cpxP transcription via RT-qPCR show that both genes are normally repressed by TyrR, but repression of cpxP is alleviated by tyrosine. TyrR bound to the promoter region of these genes at a site overlapping one of the three CpxR boxes. While the DNA binding site for TyrR was not definitively mapped, TyrR does contribute to repression of cpx by binding the promoter region. It is not at this time clear what the significance of TyrR repression of the CpxR regulon is.

The CpxR regulon controls the envelope stress response including genes for chaperones, proteases, and integral inner-membrane proteins of mostly unknown functions. CpxR also control genes for motility, adhesion, and virulence in some bacteria, and regulates the aroL and aroG genes in two strains of E. coli. In addition to dmpM and the cpx operon, genes encoding proteins involved in chemotaxis and motility, aromatic catabolism, and anaerobic respiration were perturbed in the mutant. Whether these latter systems are directly regulated by TyrR or impacted by pleiotropic effects arising from the mutation is 201

not known. Many of the differentially-regulated genes are under the control of other transcription factors that may be responding to changes in the cell. Strong TyrR binding sites were only shown to be functional in the dmpM promoter, and a weaker site is present in the cpx promoters; in the latter case, the TyrR box is atypical like the TyrR box of the akr promoter. From the results of this chapter, it is obvious that the TyrR regulon differs in E. cloacae and E. coli. However, the extent to which there are additional TyrR- regulated genes in E. cloacae is yet not fully realized. Comprehensive RNA- and ChIP- seq experiments performed under multiple conditions of aromatic amino acid availability and growth stage will be required to completely map the full extent of the regulon. While sequencing costs are rapidly shrinking, this still remains a daunting experimental challenge both technically and financially.

An interesting prospective study would be to examine the TyrR regulons across the

Enterobacteriaceae. While a TyrR-like protein seems to be present in all

Enterobacteriaceae, a multiple sequence alignment revealed that the central GAFTGA domain, characteristic of sigma-54 activating bacterial enhancer binding proteins including PhhR, is not present in the TyrR homologues found in Escherichia,

Enterobacter, Salmonella, Citrobacter and Klebsiella lineages (Figure 5.1A). The TyrR homologues of Erwinia, Pantoea, Serratia and Yersinia genuses retain the GAFTGA loop, but with changes to the three central amino acids. The sequence in the remaining genera of the Enterobacteraceae contains two to four amino acid deletions, while the

Pasteurellaceae have an almost complete GAFTGA deletion along with complete truncation of the N-terminal domain (amino acids 1-191 in E. coil TyrR). While PhhR retains the GAFTGA motif, and the ability to regulate sigma-54 promoters, its absence is 202

one of the critical features differentiating it from TyrR. The finding that several lineages are missing some, or all, of the GAFTGA motif is interesting. In all other lineages a vestigial portion of the loop is still present; however, it is unknown whether these residual loops confer any function. It would therefore be interesting to compare the function of TyrR among these lineages.

5.5 Concluding thoughts

Regulon identification via in silico methods relies on the presence of a conserved transcription factor and transcription factor binding sites, yet it is only a prediction of the composition of the regulon (32). Transcription factors may be subject to rapid diversification (32), reflecting the need to rewrite the cells’ genetic circuitry in response to new ecological niches (33-35). Experimental validation is required to confirm the differences in regulatory networks among different bacterial species with conserved transcription factors (36). For example, despite the presence of a conserved DNA binding domain, the global transcription factor Lrp (leucine responsive regulatory protein) displays considerable differences in its regulon composition among E. coli, Vibrio spp. and Proteus spp., due to only a few amino acid differences in the amino-terminal tail which impacted DNA affinity (37, 38). In the age of big data, experimental validation of regulons can be integrated with other systems level information regarding bacterial physiology and host interactions. Having manually curated results helps to fill voids left by automated annotation of experimental results, which are often propagated across databases.

203

The majority of our knowledge of TyrR has come from investigations of E. coli using selectable phenotypes and classical forward genetic techniques. While this has reliably led to the identification of the TyrR core regulon in E. coli, there is evidence that additional DNA binding sequences interact with this transcription factor. Most recently, genomic SELEX predicted additional TyrR binding sites in E. coli (12). Even if these are not regulatory sites for TyrR their interaction with TyrR, even transiently, would alter levels of available TyrR and therefore affect regulation at bona fide TyrR-controlled promoters. Moreover, while high-throughput analysis of global gene regulation and protein-DNA binding via RNA- and ChIP-sequencing is expanding the known regulons of well-studied bacterial transcription factors, often these studies identify many

‘artifactual’ binding sites that deviate from known consensus sequences and therefore are not considered in further analyses. However, it is possible that these represent true, non- canonical transcription factor binding sites with motif fuzziness. Therefore, even with the broader view of gene regulation afforded by these new investigatory techniques, researchers may be underestimating the true extent of transcription factors interactions with the genome. As shown in Chapters 2 and 4, TyrR interacts with non-canonical

DNA-binding sites. These binding sites control the transcription of metabolic genes that have functions peripheral to, or unrelated to, aromatic amino acid biosynthesis, indicating an expanded function for TyrR.

The ubiquitous bacterium E. cloacae is an important human pathogen and a beneficial rhizobacterium. This bacterium can exist as both a free-living organism in soils, as a plant endosymbiont, or as an enteric commensal in an animal host. Its diverse lifestyles and ability to thrive in a variety of habitats make it a fascinating and applicable 204

species to study. Furthermore, it is closely related to the model bacterium E. coli, and is therefore amenable to manipulation by standard molecular biology techniques. To successfully inhabit the rhizosphere and the mammalian intestine, E. cloacae must be metabolically versatile and capable of exploiting a diverse array of available nutrients.

Genome sequencing has revealed that E. cloacae strains have varying genome sizes and metabolic capabilities. The production of IAA and plant growth promotion by E. cloacae

UW5 made this strain an attractive organism with which to examine the underlying molecular and genetic mechanisms required for plant interactions. The finding that the

TyrR regulon is substantially different in E. cloacae and E. coli reinforces its suitability for studying transcription factor network evolution among closely related organisms.

Despite continuing advances in agricultural technology, the yield per hectare of crops is beginning to plateau as the limits of growth become constrained by the biology of plants growth limits. The diverting of land usage towards livestock or bioethanol also impacts crop production. Furthermore, as climate change continues to alter local and regional environmental conditions, previously fertile land may become unusable in the future due to increased flooding, drought, or altered weather patterns. Thus, societies must look to every available tool to combat disruptions in food security in the coming decades as these combined effects impact food security. Research into plant-bacterial interactions, and specifically plant growth-promoting rhizobacteria, is one such tool that may be exploited to improve crop yields, decrease pathogen susceptibility, or increase resistance to abiotic stressors such as drought or salinity. There has been much interest in identifying strains of bacteria that may be used as commercial inoculants to provide these benefits. Application of such bacteria in field trials has often yielded mixed results. 205

Successful exploitation of these bacteria as biofertilizers for food crops will require a comprehensive understanding of the underlying mechanisms of the beneficial interactions.

5.6 References

1. van Hijum SA, Medema MH, Kuipers OP. 2009. Mechanisms and evolution of control logic in prokaryotic transcriptional regulation. Microbiol Mol Biol Rev 73:481–509. 2. Balleza E, López-Bojorquez LN, Martínez-Antonio A, Resendis-Antonio O, Lozada-Chávez I, Balderas-Martínez YI, Encarnación S, Collado-Vides J. 2009. Regulation by transcription factors in bacteria: beyond description. FEMS Microbiol Rev 33:133–151. 3. Pittard AJ, Camakaris H, Yang J. 2005. The TyrR regulon. Mol Microbiol 55:16–26. 4. Song J, Bonner CA, Wolinsky M, Jensen RA. 2005. The TyrA family of aromatic-pathway dehydrogenases in phylogenetic context. BMC Biol 3:13. 5. Bush M, Dixon R. 2012. The role of bacterial enhancer binding proteins as specialized activators of σ54-dependent transcription. Microbiol Mol Biol Rev 76:497–529. 6. Pittard AJ, Davidson BE. 1991. TyrR protein of Escherichia coli and its role as repressor and activator. Mol Microbiol 5:1585–1592. 7. Zhao S, Zhu Q, Somerville RL. 2000. The sigma(70) transcription factor TyrR has zinc-stimulated phosphatase activity that is inhibited by ATP and tyrosine. J Bacteriol 182:1053–1061. 8. Smith HQ, Somerville RL. 1997. The tpl promoter of Citrobacter freundii is activated by the TyrR protein. J Bacteriol 179:5914–5921. 9. Ryu RJ, Patten CL. 2008. Aromatic amino acid-dependent expression of indole-3- pyruvate decarboxylase is regulated by TyrR in Enterobacter cloacae UW5. J Bacteriol 190:7200–7208. 10. Coulson TJD, Patten CL. 2015. The TyrR Transcription Factor Regulates the Divergent akr-ipdC Operons of Enterobacter cloacae UW5. PLoS ONE 10:e0121241. 11. Patten CL, Blakney AJC, Coulson TJD. 2013. Activity, distribution and function of indole-3-acetic acid biosynthetic pathways in bacteria. Crit Rev Microbiol 39:395–415. 12. Yang J, Ogawa Y, Camakaris H, Shimada T, Ishihama A, Pittard AJ. 2007. folA, a new member of the TyrR regulon in Escherichia coli K-12. J Bacteriol 189:6080–6084. 13. Sarsero JP, Pittard AJ. 1991. Molecular analysis of the TyrR protein-mediated activation of mtr gene expression in Escherichia coli K-12. J Bacteriol 173:7701– 7704. 206

14. Sarsero JP, Wookey PJ, Pittard AJ. 1991. Regulation of expression of the Escherichia coli K-12 mtr gene by TyrR protein and Trp repressor. J Bacteriol 173:4133–4143. 15. Yang J, Hwang JS, Camakaris H, Irawaty W, Ishihama A, Pittard AJ. 2004. Mode of action of the TyrR protein: repression and activation of the tyrP promoter of Escherichia coli. Mol Microbiol 52:243–256. 16. Lawley B, Fujita N, Ishihama A, Pittard AJ. 1995. The TyrR protein of Escherichia coli is a class I transcription activator. J Bacteriol 177:238–241. 17. Pittard AJ, Yang J. 2008. Biosynthesis of the Aromatic Amino Acids. EcoSal Plus 3. 18. Heatwole VM, Somerville RL. 1992. Synergism between the Trp repressor and Tyr repressor in repression of the aroL promoter of Escherichia coli K-12. J Bacteriol 174:331–335. 19. Heatwole VM, Somerville RL. 1991. The tryptophan-specific permease gene, mtr, is differentially regulated by the tryptophan and tyrosine repressors in Escherichia coli K-12. J Bacteriol 173:3601–3604. 20. Spaepen S, Versées W, Gocke D, Pohl M, Steyaert J, Vanderleyden J. 2007. Characterization of phenylpyruvate decarboxylase, involved in auxin production of Azospirillum brasilense. J Bacteriol 189:7626–7633. 21. Broek AV, Gysegom P, Ona O, Hendrickx N, Prinsen E, Van Impe J, Vanderleyden J. 2005. Transcriptional analysis of the Azospirillum brasilense indole-3-pyruvate decarboxylase gene and identification of a cis-acting sequence involved in auxin responsive expression. MPMI 18:311–323. 22. Gupta RS. 2000. The phylogeny of proteobacteria: relationships to other eubacterial phyla and eukaryotes. FEMS Microbiol Rev24:367–402. 23. Desai KK, Miller BG. 2008. A metabolic bypass of the triosephosphate isomerase reaction. Biochemistry 47:7983–7985. 24. Jez JM, Bennett MJ, Schlegel BP, Lewis M, Penning TM. 1997. Comparative anatomy of the aldo-keto reductase superfamily. Biochem J 326 (Pt 3):625–636. 25. Mindnich RD, Penning TM. 2009. Aldo-keto reductase (AKR) superfamily: genomics and annotation. Hum Genomics 3:362–370. 26. Kalyananda MK, Engel R, Tropp BE. 1987. Metabolism of L-glyceraldehyde 3- phosphate in Escherichia coli. J Bacteriol 169:2488–2493. 27. Alber T, Banner DW, Bloomer AC, Petsko GA, Phillips D, Rivers PS, Wilson IA. 1981. On the three-dimensional structure and catalytic mechanism of triose phosphate isomerase. Philos Trans R Soc B Biol Sci 293:159–171. 28. Grant AW, Steel G, Waugh H, Ellis EM. 2003. A novel aldo-keto reductase from Escherichia coli can increase resistance to methylglyoxal toxicity. FEMS Microbiol Lett 218:93–99. 29. Coulson TJD, Patten CL. 2015. Complete genome sequence of Enterobacter cloacae UW5, a rhizobacterium capable of high levels of indole-3-acetic acid production. Genome Announcements 3:e00843–15. 30. Zhou D, Yang R. 2006. Global analysis of gene transcription regulation in prokaryotes. Cell Mol Life Sci 63:2260–2290.

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31. Wilhelm BT, Landry J-R. 2009. RNA-Seq—quantitative measurement of expression through massively parallel RNA-sequencing. METHODS 48:249–257. 32. Price MN, Dehal PS, Arkin AP. 2007. Orthologous transcription factors in bacteria have different functions and regulate different genes. PLoS Comput Biol 3:1739–1750. 33. Lozada-Chávez I, Janga SC, Collado-Vides J. 2006. Bacterial regulatory networks are extremely flexible in evolution. Nucleic Acids Res 34:3434–3445. 34. Damkiær S, Yang L, Molin S, Jelsbak L. 2013. Evolutionary remodeling of global regulatory networks during long-term bacterial adaptation to human hosts. Proc Natl Acad Sci USA 110:7766–7771. 35. Galán-Vásquez E, Sánchez-Osorio I, Martínez-Antonio A. 2016. Transcription Factors Exhibit Differential Conservation in Bacteria with Reduced Genomes. PLoS ONE 11:e0146901. 36. Kılıç S, Erill I. 2016. Assessment of transfer methods for comparative genomics of regulatory networks in bacteria. BMC Bioinformatics 17 Suppl 8:277. 37. Lintner RE, Mishra PK, Srivastava P, Martinez-Vaz BM, Khodursky AB, Blumenthal RM. 2008. Limited functional conservation of a global regulator among related bacterial genera: Lrp in Escherichia, Proteus and Vibrio. BMC Microbiol 8:60. 38. Hart BR, Mishra PK, Lintner RE, Hinerman JM, Herr AB, Blumenthal RM. 2011. Recognition of DNA by the helix-turn-helix global regulatory protein Lrp is modulated by the amino terminus. J Bacteriol 193:3794–3803.

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A B * * * G A F T G A * Escherichia fergusonii E L F G H A P ------E G K K G 81 Escherichia albertii E L F G H A P ------E G K K G

97 Escherichia coli E L F G H A P ------E G K K G 98 93 Shigella flexneri E L F G H A P ------E G K K G

55 Citrobacter rodentium E L F G H A P ------E G K K G Salmonella enterica E L F G H A P ------E G K K G 77 54 Citrobacter freundii E L F G H A P ------E G K K G 99 Citrobacter youngae E L F G H A G ------E G K K G Enterobacter sp. 638 E L F G H A P ------E G K K G 52 Enterobacter cancerogenus E L F G H A P ------E G K K G 99 99 Enterobacter cloacae UW5 E L F G H A P ------E G K K G 63 Enterobacter hormaechei E L F G H A P ------E G K K G

99 Cronobacter sakazakii E L F G H A P G A Y A N A - H E G K K G

100 Cronobacter turicensis E L F G H A P G A Y A N A - H E G K K G 49 Cronobacter universalis E L F G H A P G A Y A N A - H E G K K G Klebsiella aerogenes E L F G D A S ------Q G K K G 100 Klebsiella pneumoniae E L F G D A L ------Q G K K G Enterobacteriales 78 100 Erwinia amylovora E L F G H A A G A Y P N A - L Q G K K G 55 Erwinia pyrifoliae E L F G H A A G A Y P N A - L Q G K K G 99 Erwinia billingiae E L F G H A A G A Y P N A - L E G K K G Pantoea agglomerans E L F G H A A G A Y P N A - L E G K K G 88 Pantoea stewartii E L F G H A A G A Y P N A - L E G K K G

68 Pectobacterium carotovorum E L F G H A P G A Y L N A - Q E G K K G 41 100 Pectobacterium parmentieri E L F G H A P G A Y L N A - Q E G K K G

100 Pectobacterium atrosepticum E L F G H A S G A Y L N A - Q E G K K G 77 Serratia sp. S4 E L F G H A P G A Y P N A - L E G K K G 40 Serratia symbiotica E L F G H A P G A Y P N A - L E G K K G

99 Serratia sp. M24T3 E L F G Y A P G A Y P N A - V D G K K G Yersinia pestis E L F G Y A A G A Y P N A - I E G K K G 35 TyrR 70 99 Yersinia enterocolitica E L F G Y A A G A Y P N A - V E G K K G 98 Yersinia intermedia E L F G Y A A G A Y P N A - I E G K K G Proteus mirabilis E L F G Y A A G A Y P N A - I E G K K G

51 Photorhabdus asymbiotica E L F G Y A A G A Y P N A - I E G K K G 32 Providencia stuartii E L F G Y A A G A Y P N A - L D G K K G

79 Aeromonas molluscorum E L F G Y A P G A F G N N - T E G K R G

100 Aeromonas hydrophila Aeromonadales E L F G Y A P G A F G N N - T E G K R G 99 Aeromonas salmonicida E L F G Y A P G A F G N N - T E G K R G

100 Photobacterium angustum E L F G F A G S T T - - - - E P S K K G 100 Photobacterium leiognathi E L F G F A G S A T - - - - E P S K K G Photobacterium damselae E L F G E V A T I G - - - - E P S K K G Vibrionales 36 93 Vibrio harveyi E L F G H A P G S F N H - - E Q G H K G

100 Vibrio anguillarum E L F G H A P G S F N H - - Q Q G H K G 97 Vibrio cholerae E L F G H A P G S F N H - - Q Q G H K G

100 Pseudoalteromonas piscicida E L F G Y - A G Y E E N - - A A P K R G Pseudoalteromonas rubra E L F G Y - A G Y E E N - - S A P K R G

100 Glaciecola nitratireducens E L F G I E H G Q F N G - - S D S K P G 77 Glaciecola pallidula E L F G I E Q G E F S G - - A E T K T G 55 Glaciecola punicea E L F G Y A V G S A E E - - S E G K A G 95 100 Alteromonas macleodii E L F G Y A A G A F N Q - - P N A K V G 100 Alteromonas mediterranea 99 Alteromonadales E L F G Y A A G A F N Q - - P N A K V G Alteromonas naphthalenivorans E L F G Y A Q G A F N Q - - P N A K E G Paraglaciecola arctica E L F G Y I S E T S E Q - - T E N K V G 63 100 Paraglaciecola psychrophila E L F G H A A G I D G Q - - T D R K V G 59 98 Paraglaciecola polaris E L F G Y A A G A F G Q - - T S G K E G 100 Pseudoalteromonas atlantica E L F G Y A A G A F G Q - - S T G K E G Shewanella marina E L F G Y M N Q G Q - - - - - V L K R G

100 Shewanella halifaxensis Shewanellaceae E L F G F V S N G N - - - - - V V K R G 66 Shewanella oneidensis E L F G Y V S Q G K - - - - - V I K R G Pseudomonas syringae E L F G Y D P G A F E G A R A E G R L G

100 Pseudomonas aeruginosa Pseudomonadales E L F G Y G P G A F E G A R P E G K L G PhhR 83 Pseudomonas fluorescens E L F G Y G P G A F E G A R A E G K L G Actinobacillus pleuropneumoniae E M F G H R G N G ------K E N I G Pasteurella multocida E M F G H R E Q D ------K E Y V G 100 Pasteurellales TyrR 94 Haemophilus influenzae E M F G R K V G D ------S E T I G 100 Haemophilus parainfluenzae E M F G R K V G D ------S E T S G NtrC Escherichia coli E L F G H E K G A F T G - A N T I R Q G NtrC

Figure 5.1. Phylogenetic tree of TyrR and its homologue PhhR (A). Order names are given for major groups. TyrR homologues were retrieved by BLASTp against the GenBank nonredundant database and annotated PhhR proteins from Pseudomonas were selected. Sequences were aligned with CLUSTALW and phylogenetic trees built by the UPGMA method using MEGA7. The bootstrap consensus tree is inferred from 10000 replicates and the percentage of trees clustered together are shown next to the branches. Multiple alignment of amino acids corresponding to Glu274-Gly285 from E. coli TyrR (B). Conserved amino acids are denoted above by asterisks and the sigma-54 contacting GAFTGA domain of NtrC is indicated above. Proteins classified as TyrR, PhhR or NtrC are indicated. 209

Chorismate Anthranilate A Gln phosphoribosyl trpD transferase B

Glu + Pyruvate Anthranilate PRPP Triosephosphate Anthranilate Isomerase trpD phosphoribosyl PPi transferase D-glyceraldehyde-3-phosphate Dihydroxyacetone phosphate Phosphoribosyl anthranilate NADH Phosphoribosyl trpC anthranilate Glycerol 3-phosphate Isomerase Dehydrogenase

1-(O-carboxyphenylamino) NAD+ -1-deoxyribose phosphate

trpC Indolglycerol phosphate Synthase CO2

Indoleglycerol phosphate NADPH NADP+ Glyceraldehyde Serine trpA Tryptophan Synthase 3-phosphate reductase trpB L-glyceraldehyde-3-phosphate (YghZ) L-glycerol-3-phosphate D-GAP Tryptophan

Figure 5.2. The tryptophan terminal biosynthetic pathway (A) and the triosephosphate isomerase bypass (B). D-GAP, D-glyceraldehyde-3-phosphate; L- GAP, L-glyceraldehyde-3-phosphate; DHAP, Dihydroxyacetone phosphate; L-G3P, L- glycerol-3-phosphate; TIM, Triosephosphate isomerase; GRP, Glyceraldehyde-3- phosphate reductase; GPDH, glycerol-3-phosphate dehydrogenase.

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Curriculum Vitae

Candidate’s full name: Thomas Joseph Dana Coulson

Universities attended (with dates and degrees obtained):

B. Sc. (Honors), Department of Biology, University of New Brunswick, 2008

Publications:

Published

Coulson, T.J.D. and Patten, C.L. (2015) Complete genome sequence of Enterobacter cloacae UW5, a rhizobacterium capable of high levels of indole-3-acetic acid production. Genome Announcements, 3(4): e00843-15; doi10.1128/genomeA00843-15

Coulson, T.J.D. and Patten, C.L. (2015) The TyrR transcription factor regulated the divergent akr-ipdC operons of Enterobacter cloacae UW5. PLoS One, 10(3): e0121241; doi10.1371/journal.pone.0121241

Patten, C.L., Blakney, A.J. and Coulson, T.J.D. (2013) Activity, distribution and function of indole-3-acetic acid biosynthetic pathways in bacteria. Critical Reviews in Microbiology, 39(4): 395-415.

English, M.M., Coulson, T.J.D., Horsman, S.R. and Patten, C.L. (2010) Over expression of hns in the plant growth promoting bacterium Enterobacter cloacae UW5 increases root colonization. Journal of Applied Microbiology, 108(6): 2180-2190.

Mesanza, N., Crawford, B.D., Coulson, T.J.D., Iturritxa, E., Patten, C.L. 2018. Colonization of Pinus radiata D. don seedling roots by biocontrol bacterium Erwinia billingiae and Bacillus simplex. Forests, 10:522.

In Preparation

Coulson, T.J.D., Malenfant, R., Patten, C.L. 2018. Characterization of the TyrR regulon in the rhizosphere bacterium Enterobacter cloacae UW5 reveals overlap with the CpxR envelope stress response. Submitted, Journal of Bacteriology.

Conference Presentations:

Coulson, T.J.D. and Patten, C.L. (2018) Characterization of the aromatic amino acid responsive TyrR transcription factor in Enterobacter cloacae UW5 reveals overlap with the CpxR envelope stress response. UNB Graduate Student Research Conference. Fredericton, Canada.

Coulson, T.J.D. and Patten, C.L. (2012) Regulation of overlapping, divergent promoters by the global transcription factor TyrR. American Society of Microbiology 112th General meeting. San Francisco, USA.

Coulson, T.J.D. and Patten, C.L. (2011) Regulation of indole-3-acetic acid biosynthesis in Enterobacter cloacae UW5 by the transcription factor TyrR. Rhizosphere 3 International Conference. Perth, Australia.

Coulson, T.J.D. and Patten C.L. (2010) Characterization of the Enterobacter cloacae UW5 akr gene and its implications in indole-3-acetic acid production. Canadian Society of Microbiology 60th Annual Conference. Hamilton, Ontario.

Coulson, T.J.D. English, M.M., Horsman, S.R. and Patten C.L. (2009) Quantification of hnsA gene expression in the rhizosphere in a hyper-colonizing Enterobacter cloacae UW5 mutant. Canadian Society of Microbiology 59th Annual Conference. Montreal, Quebec.